View of Respiratory Disease 145, 669–674 (1992)
Allergen-induced chemokine release from the bronchial epithelium:
A novel lysosomal release mechanism
A dissertation submitted to the University of Cincinnati
In partial fulfillment of the requirements for the degree of
Doctor of Philosophy (Ph.D.)
In the graduate program of Immunobiology
Of the College of Medicine
2013
By
Mark Webb
B.S., Brigham Young University, 2007
Committee members
Chair: Simon P. Hogan, Ph.D.
Marshall Montrose, Ph.D.
Lee Denson, M.D.
Nives Zimmermann, M.D.
Marsha Wills-Karp, Ph.D. Abstract
Asthma is a chronic inflammatory disease of the lung. In recent years, the airway epithelium has been identified as an important contributor to the initial development of asthma after allergen exposure. The mechanisms by which epithelial cells respond to allergen to help drive downstream inflammation and the initiation of an immune response are poorly understood. In this dissertation we sought to understand the basic mechanisms that control release of chemokines and cytokines in general and CCL20 in particular. To do this, we measured levels of CCL20 and other cell mediators retained in cells and released from cells, and we visualized these mediators' intracellular localization via immunofluorescence microscopy. We found that a number of chemokines and cytokines are stored within bronchial epithelial cells under homeostatic conditions. In addition, CCL20 was stored in lysosomes, and its release was partially mediated through secretion of these lysosomes. In order to determine the intracellular localization of multiple chemokines and cytokines simultaneously, we again employed fluorescence microscopy. We also developed a novel technique to measure specific proteins in subcellular organelles using modified flow cytometry. We found that lysosomes containing
CCL20 are released after exposure to HDM. We also found that the secretory lysosomes containing CCL20 also appear to contain other chemokines and cytokines released in response to HDM. We also found that allergens other than HDM induce release of specific subsets of chemokines and cytokines. These studies demonstrate a complex storage and release mechanism for chemokines and cytokines of the bronchial epithelium. These findings improve our understanding of the initial interactions between allergen and the epithelium, and provide
i an interesting potential target for future treatments that will allow them to be both targeted and robust.
ii
Acknowledgments
I would like to thank my advisor, Dr. Marsha Wills-Karp for her time and effort in helping me refine my scientific understanding. In addition, she provided a lab staffed with excellent post- doctoral fellows and graduate students who have also impacted me tremendously as I completed my degree. In particular, Stephane Lajoie, Stacey Burgess, and Ian Lewkowich have always provided a listening ear and sound opinions about specific hypotheses as needed. Their insights proved valuable as I sought to uncover the mechanisms presented herein. I would also like to thank my wife for her support and encouragement throughout the dissertation-writing process.
iii Table of Contents
ABSTRACT ...... I ACKNOWLEDGMENTS ...... III TABLE OF CONTENTS ...... IV LIST OF ABBREVIATIONS ...... V CHAPTER 1 – GENERAL INTRODUCTION ...... 1 CHAPTER 2 – MECHANISM OF CCL20 RAPID RELEASE FROM ALLERGEN-EXPOSED EPITHELIAL CELLS ...... 56 CHAPTER 3 – SIGNALING PATHWAYS LEADING TO HDM-INDUCED CCL20 RELEASE FROM THE BRONCHIAL EPITHELIUM ...... 93 CHAPTER 4 – ALLERGEN-INDUCED RELEASE OF MANY PREFORMED CHEMOKINES AND CYTOKINES IS THROUGH SECRETORY LYSOSOMES ...... 139 CHAPTER 5 – SUMMARY AND DISCUSSION ...... 194
iv List of abbreviations
AHR Airway hyperresponsiveness APTI Airway pressure time index ATP Adenosine triphosphate AUB Ambient urban Baltimore particulate matter BAL Bronchoalveolar lavage BLAST Basic local alignment search tool CCL C-C chemokine ligand CCR C-C chemokine receptor CFTR Cystic fibrosis transmembrane conductance receptor COX Cyclooxygenase CXCL C-X-C chemokine ligand CXCR C-X-C chemokine receptor DC Dendritic cell ELISA Enzyme-linked immunosorbent assay ENaC Epithelial sodium channel ERK Extracellular signal-relgulated kinase FBS Fetal bovine serum FGF Fibroblast growth factor GM-CSF Granulocyte-macrophage colony stimulating factor HDM House dust mite HS Heparan sulfate HSPG Heparan sulfate proteoglycan IL Interleukin ILC Innate lymphoid cells i.t. intra trachea JNK c-Jun N-terminal kinase LAMP Lysosome associated membrane protein LD50 Median lethal dose LPS Lipopolysachharride mTEC Mouse tracheal epithelial cell NHBE Normal human bronchial epithelial cell NKCC Na-K-Cl cotransporter NLRP3 NOD-like receptor family, pyrin domain containing 3 NSAID Non-steroidal anti-inflammatory drug PAMP Pathogen-associated molecular pattern PBS Phosphate-buffered saline pH Potential of hydrogen (measure of acidity) PKC Protein kinase C PNS Post-nuclear supernatant PRR Pattern recognition receptor RAB Ras-related protein
v ROS Reactive oxygen species SOFA Single organelle fluorescence analysis SYK Spleen tyrosine kinase TCR T-cell receptor TGF-β Transforming growth factor beta TH Helper T-cell TNF-α Tumor necrosis factor alpha TSLP Thymic stromal lymphopoietin V-ATPase Vacuolar-type H+ adenosine triphosphatase
Common Ions Ca2+ Calcium ion Cl- Chloride ion K+ Potassium ion Mg2+ Magnesium ion Na+ Sodium ion 3- PO4 Phosphate ion 2- SO4 Sulfate ion
vi CHAPTER 1 – GENERAL INTRODUCTION
ASTHMA ...... 2
SYMPTOMS AND TREATMENT ...... 2 INCIDENCE AND COSTS ...... 3 ETIOLOGY ...... 4 MAJOR ALLERGENS ASSOCIATED WITH ALLERGIC ASTHMA ...... 7 PHYSIOLOGICAL CHANGES IN ASTHMATIC LUNGS ...... 10 CELLULAR AND MOLECULAR MECHANISMS OF DISEASE ...... 12
ALLERGIC ASTHMA IS MEDIATED BY TYPE -2 CELLS ...... 13 DEVELOPMENT OF A TH2 RESPONSE ...... 16 Dendritic cells in asthma ...... 17 THE BRONCHIAL EPITHELIUM IN ASTHMA ...... 20 Ion transport in bronchial epithelial cells ...... 21 The bronchial epithelium and immunity ...... 26 ROLE OF CHEMOKINES /CYTOKINES IN ALLERGIC ASTHMA ...... 27 Inflammatory chemokines ...... 28 CCL20 ...... 28 Other inflammatory chemokines and cytokines ...... 31 Secretory lysosomes ...... 33 Syndecans and heparin sulfate proteoglycans ...... 38 SUMMARY ...... 40 REFERENCES ...... 43
1 Chapter 1 – General introduction
Asthma
Asthma is a debilitating chronic disease of the airways characterized by chest tightness, difficulty breathing, coughing, and wheezing. 1. These symptoms are mainly caused by changes in the lungs that result in narrowing of the conducting airways.2,3
Breathing becomes more difficult because the constricted airways increase resistance to air flow.4
Symptoms and treatment
The changes in the airways that lead to many of the symptoms described above include increased mucous production due to goblet cell hyperplasia 5, airway inflammation, angiogenesis, and smooth muscle hypertrophy. 6 One of the primary treatments for asthma symptoms is inhaled corticosteroids. 7 Inhaled corticosteroids are
an inexpensive treatment that has proven effective at controlling more common, mild
cases of asthma since the 1960s. They work by reducing inflammation in the airways. 8
However, patients with more severe asthma show a decreased response to
corticosteroids, and increasingly higher doses are given to patients with greater disease
severity. 9 As the dose of corticosteroids increases, many negative side effects begin to manifest including bone loss, hyperglycemia, depression, and cataracts among others. 10–
12 The reason why some asthma sufferers develop mild symptoms, while others
2 develop severe disease have been poorly characterized in the past. Thus, much research has focused on better understanding the molecular mechanisms of asthma pathogenesis, and especially the differences between pathogenesis of mild and severe asthma, in the hopes of developing more effective strategies for treatment and control – particularly for those who do not respond well to current treatment strategies.
Incidence and costs
Asthma is among the most common chronic diseases in the world, with an estimated 300 million people affected worldwide.13 Annual worldwide deaths from
asthma are estimated at around 250,000 individuals. In the United States, the asthma
prevalence as of 2008 was 8.5% of the population among adults, and 9.0% of children.
In recent decades, incidence of asthma in developed countries has been
increasing, 14 with one study in the United States showing asthma rates doubling over a
20 year period from 1964to 1983. 15 Although both genetic and environmental triggers have been shown to contribute to the development of asthma, evidence suggests that the increased incidence of asthma we have observed over the past few decades is the result of changes in environment and lifestyle in developed societies, which can occur over the course of a few decades; as opposed to genetic changes, which tend to occur over much longer time periods. 16 One striking feature about elevated levels of asthma
is that they are localized mainly to economically developed countries, with developing
countries experiencing higher rates of asthma as they become more economically
3 developed. 17 This can be seen from the heat map of asthma depicted in figure 1. This
region-specific elevation in asthma prevalence has led to a number of questions about
why some individuals develop asthma, in an attempt to better understand how to
prevent further increases in the general development of asthma.
Figure 1. World map of the global burden of asthma. Adapted from GINA 2004.
Etiology
Although the exact etiology of asthma is currently unknown, it is thought to be a heterogeneous disease that is driven by both genetic 18–22 and environmental influences.16,23 Many genetic risk factors have been explored through the use of genome-wide association studies (GWAS), which have identified a number of genes that may play a role in asthma pathogenesis. For some of these, the potential
4 mechanisms are well characterized, such as with tumor necrosis factor (TNF)-α and interleukin (IL) 4. For others, potential functional implications, and the link to asthma pathogenesis, are only beginning to be elucidated – such as with ORMDL3 and gasdermin B.24,25 Although mutations in any one of these genes may increase the risk of asthma marginally, no one gene has been directly identified as causing asthma and as such, asthma has been identified as a polygenic disease. This is in contrast to monogenic diseases – such as cystic fibrosis, which is directly linked to mutations in the cystic fibrosis transmembrane conductance receptor (CFTR). 26 The genetic cause of cystic fibrosis is much more well-characterized, as it is the primary driver of disease.
This is not the case with asthma, where multiple genetic changes contribute to a greater susceptibility to developing the disease, and asthma is not the result of a single aberrant gene. This may be due to the apparent need to activate multiple pathways leading to the development of asthma.
Indeed, in recent years asthma has become increasingly sub-divided into different types, including exercise-induced asthma, atopic asthma, childhood asthma, and so on. Within atopic asthmatics, we also see multiple different types of exacerbations with mild, moderate, and severe asthmatics who are responsive or resistant to interventions that target different molecular pathways.27 This is similar to
5 categorization of different types of cancers, where cancers driven by different molecular pathways respond to different types of treatment.
As mentioned previously, the dramatic elevation in asthma prevalence in developed regions is too rapid to be driven by genetic changes and therefore a number of non-genetic explanations have been explored to better understand this phenomenon.28 Thus, in addition to genetic differences, a number of environmental changes have also been shown to contribute to asthma susceptibility. For instance, exposure to tobacco smoke causes increased wheezing in genetically susceptible children. 29 Socio-economic status is also a risk factor for asthma, with higher asthma incidence and hospitalization rates for individuals from poorer communities. 30
Increased use of antibiotics in recent decades has led to changes in the gut microbiota that may contribute to allergic sensitization. 31
Other major behavioral and environmental changes seen more in developed
nations include migration away from farms into cities. In addition, as populations
move into urban environments, they are exposed to airborne pollutants such as
particulate matter and nitrous acid, both of which may exacerbate their asthma
symptoms, 32–34 and contribute to a change in their microbiota. 35–37 Living in an urban environment has also lead to increasing amounts of time spent indoors.38–41 Even the
6 altered diets in developed nations may contribute to the development of asthma. 42 Our
understanding of how each of these changes may lead to increased risk of asthma has
improved in recent years. Better understanding of the specific mechanisms leading to
asthma development may lead to the development of better strategies to prevent
further increases in asthma incidence. Although many of these changes affect similar
downstream pathways related to the development of asthma, a better understanding of
how each change, and indeed each allergen, contributes uniquely to the development of
atopy differently is poorly understood.
Major allergens associated with allergic asthma
Atopic asthma is defined as the development of an allergen-specific IgE response in association with asthma. Another way to characterize atopic asthma is the development of an aberrant immune response to otherwise innocuous airborne stimuli.
Early exposure to specific allergens has been linked to development of atopy to those allergens. 43 However, not all potential antigens that enter the airways have the potential to become allergens. Instead, a relatively small number of antigens have been identified as capable of driving allergic asthma. For example, one of the most potent allergens is house dust mite (HDM), with HDM-specific IgE responses found in up to
80% of atopic asthmatics. 44 Common classes and features have been recognized among the many identified allergens in atopic individuals. Some classes include insect
7 allergens, such as HDM and cockroach feces (frass); 45,46 animal allergens, such as the major cat allergen Fel d 1; 47 various species of mold, including Aspergillus fumigatus; 48,49 and many species of pollen, notably maple and birch pollens. 50
Additionally particulate matter, such as diesel exhaust and urban particulate collections, have been identified as potent irritants that dramatically affect the airways. 32 These additional danger signals may contribute to increased asthma
incidence or severity, especially in urban environments.
Specific features of these organisms and chemical mixtures have been identified
that lead to their allergenicity. One such feature is the presence of active proteases,
which contribute to damage of the airway epithelium. This may contribute to their
allergenic behavior. 51–53 For example, the protease activity in Der p 1, found within
HDM, 54 contributes to the degradation of the tight junctions between airway epithelial cells, providing access to other proteins and molecules found in HDM to the sub- epithelial environment. 55 Other allergens; such as pollens from birch, maple, and
ragweed; have also been demonstrated to contain active proteases that may play a role
in allergic sensitization. 56 Thus, allergens often contain active proteases which may help provide a “danger” signal to the responding airway epithelium. 57
8 However, breakdown of the epithelial barrier itself may not be sufficient to induce an immune response, nor is it required. Allergens are often a complex combination of proteins, carbohydrates and other moieties whose combined effect is more potent at eliciting an asthmatic response than individual proteases, or other components, alone. HDM also contains high levels of chitins, which are also found in many insects and molds.58 These chitins may serve to signal through pattern recognition receptors (PRRs) to promote an asthma response. 59 In addition, HDM
contains many species of bacteria in its gut; 60 these bacteria aid in digestion of the mite’s food source (skin cells) and may also play a role in allergic sensitization. Upon exposure of the bacteria, pattern recognition receptors on the surface of the epithelial cells (such as toll-like receptor 4) recognize molecular patterns on bacteria such as lipopolysaccharide (LPS).61,62 This provides an initiating signal the promotes the further development of an immune response against the antigen as a whole. HDM is a prime example of the complex composition of many allergens. As such, it is also a good model for studying the complex interactions that contribute to the development of atopy. In this work, HDM was chosen as a model to study the effect of allergens on eliciting multiple different types of responses from bronchial epithelial cells. In addition, the divergent effects of other allergens on the bronchial epithelium are briefly explored.
9 Physiological changes in asthmatic lungs
Smooth muscle surrounds the airway epithelium, and may contribute to narrowing of the airways when they contract. For example, when the lungs are exposed to cold air, contraction of the airways slows air flow, which allows the conducting airway to warm the incoming air prior to exposure to the blood in the alveoli. Excessive growth of smooth muscle cells, or hypertrophy, in asthmatics contributes to narrowing of the airways under homeostatic conditions. During an asthma attack the airway smooth muscle contracts, further narrowing the airways. 63
The cells lining the airways, bronchial epithelial cells, also contribute to lung
function by helping to clear the lung of particulate matter and debris. These cells
produce a mucous layer that coats the airways. As particulate matter travels along the
airway, turbulence causes contact with the mucous layer, which it adheres to. The
epithelial cells continuously push the mucous layer up, out of the lungs to the
esophagus, where it is swallowed. In this way, the mucous layer is constantly recycled
as new mucous is produced.64 In asthmatics, clearance of this mucous layer becomes impaired, and some airways may even become plugged with mucous, obstructing air flow completely. 65
10
Figure 2. Diagram of the conducting airways in healthy and asthmatic lungs.
Finally, in addition to the features mentioned above, sub-epithelial fibrosis is commonly observed in asthmatics.66 Fibrosis develops after repeated inflammation in the airways. Fibrotic tissue is deposited below the epithelial layer, which contributes to the thickening of the airway. The combined effect of this process is an overall remodeling of the airway, which results in permanently reduced lung capacity. 67 The gross anatomical manifestation of this is narrowing of the airways, which is the primary cause of the wheezing and shortness of breath that are characteristic of asthma.68,69
11 Pathophysiological manifestations of disease
Cellular and molecular mechanisms have been identified for each of the events described above that contribute to the thickening of the airways. Thickening of the airways is caused by collagen deposition and by hypertrophy of airway smooth muscle cells. 70,71 In addition, allergen exposure may induce constriction of smooth muscle cells, further narrowing the airways. 72
After repeated allergen exposure, fibrosis begins to occur. Fibrosis is driven by cells called fibroblasts, which proliferate and migrate into cites of tissue injury in response to the cytokine transforming growth factor (TGF)-β. Fibroblasts secrete a combination of collagen and fibronectin in the formation of fibrotic scar tissue.73
The pattern of airway inflammation in asthmatics includes the presence of increased numbers of eosinophils, neutrophils, and T lymphocytes, 74–78 although acute
airway inflammation in some severe asthmatics exhibits high levels of neutrophilia and
little or no eosinophilia. 79 This indicates that asthma may be a heterogeneous disease,
with differing inflammatory phenotypes. Whatever the inflammatory phenotype, T-
lymphocytes play a crucial role in regulating the behavior of inflammatory cells, and
thus the type of immune response that develops.
12 Allergic asthma is mediated by type-2 cells
T cells are central to cell-mediated immunity. T cells each display a unique T cell receptor (TCR). This receptor is generated in the thymus, where multiple TCR genes recombine at random to produce a receptor that can bind to a unique epitope, or recognition site. Millions of these unique cells are generated in the thymus, from which they migrate in a naïve state, circulating through the blood. These naïve cells are activated through interactions with dendritic cells carrying antigen specifically recognized by that cells TCR. Once activated, these cells have tremendous proliferative capacity, and help direct the immune system to respond to specific antigens recognized
by their unique TCR. This process of expanding previously generated populations of antigen-specific cells is known as Clonal Selection Theory. 80
After activation, CD 4 + T cells differentiate into specific subsets, instructed by the specific co-stimulatory molecules and cytokines supplied from the presenting DC. 81
Initially, these CD 4 + helper T cells were thought to differentiate into two distinct
subsets, T H1, and T H2 cells. In recent years, many different types of T cell subsets have
been identified, including T H1, T H2, regulatory T cells, and T H17 cells with additional
CD 4 + T cell subsets having been proposed.
Each T cell subset plays an important role in a number of immune responses. 82
In normal lungs, T H2 cells function to promote humoral defense, and defense against
13 83 parasites. In the pathogenesis of asthma, TH2-type T cells play a central role, and are enriched in the lungs of asthmatics. 84–87 They are characterized by production of many
interleukins (IL), including IL-4, IL-5, IL-6, IL-10, and IL-13. TH2 cells are characterized
by their activation of the transcription factor GATA-3, which helps preserve its
88 differentiation to the T H2 phenotype.
In asthma, IL-4, IL-5, and IL-13 are particularly important in helping to promote
many of the phenotypic changes associated with the disease.89 These interleukins together mediate B-cell class switching to IgE, eosinophilia, goblet cell metaplasia, and
bronchoconstriction. IL-4 is necessary for B cell class switching to IgE. 90 Additionally,
IL-4 signals to T H2 cells themselves to drive GATA-3. This positive feedback loop is
critical to maintain a T H2 phenotype. IL-5 is necessary for development of eosinophilia in mouse models of asthma. 91 IL-13 promotes mucous production and AHR.92
Previously, the importance of IL-13 in allergic asthma was highlighted by
demonstrating that IL-13 can directly induce experimental allergic asthma in mice.
Interestingly, mice deficient in IL-4 can still develop AHR in an OVA-induced asthma
mode, however these same mice do not develop AHR when IL-13 neutralizing
antibodies are administered.93,94 Additionally, IL-13 knockout mice do not develop
AHR in an allergen model of asthma. 95 Thus, IL-13 has been considered a central
mediator of allergic asthma.
14
Recent studies have highlighted a new cell type that is also capable of driving a type-2 immune response. These cells are a type of innate lymphoid cells (ILC) that produce type-2 cytokines (ILC2 cells). 96 They respond to cytokines secreted by the
bronchial epithelium, IL-25 and IL-33, by secreting large amounts of IL-5 and IL-13. 97
They are distinct from T H2 cells in that they do not express T-cell or B-cell markers,
though they are hematopoietic in origin. 98 Although these cells are not very abundant in tissues, their ability to secrete large amounts of cytokines highlights their potential importance in establishing a type-2 response.99
In addition to type-2 cells, our lab previously demonstrated a role for T H17 cell responses in asthma. While T H17 responses alone were insufficient to drive an asthmatic phenotype, we found that the T H17 cytokine IL-17 synergizes with the classical T H2 cytokines IL-4, IL-5, and IL-13 to produce exacerbated disease, including increased neutrophilia and increased airway hyperresponsiveness.100 Interestingly, IL-
17 alone was insufficient to drive increased AHR, but required the establishment of a
TH2 response to elicit its exacerbating effect on AHR. Thus, asthma dominated by type-
2 responses alone, whether driven by T H2 cells, or ILC2 cells, may represent a milder phenotype, while asthmatics that exhibit both a type-2 and a T H17-mediated response may represent a severe asthma phenotype.
15 Development of a T H2 response
When T cells migrate out of the thymus, they are characterized as naïve T-cells, and require stimulation in the presence of antigen to drive differentiation. Naïve T-cells circulate through secondary lymphoid organs, such as the spleen and lymph nodes, continuously sampling antigen that is presented on the surface of dendritic cells attached to major histocompatibility complex class II molecules. When it encounters the specific antigen-MHCII complex recognized by its TCR, the T-cell is arrested at the site of the DC. This first signal is accompanied by two additional signals that instruct T cell differentiation. The second signal provided by the DC is co-stimulatory molecules.
These are receptor/ligand pairs on the surface of both cells and include CD80/86 which
binds to CD28, and OX40 which binds to OX40L.101,102 In asthmatics, expression of
CD80 and CD86 are both up-regulated, and correlate with increased asthma
severity. 103,104 In a murine model of asthma, mice deficient in OX40L were unable to
mount a TH2 response after OVA challenge. 105 In addition to direct co-stimulation, T-
cells require a third signal that participates in T-cell skewing, secreted cytokines. In
allergic asthma, these signals have been shown to be provided by IL-25, IL-33, and TSLP
to drive a TH2 phenotype. 106–108 For example, T-cells stimulated with anti-CD28 and IL-
33 in vitro began to produce the TH2 cytokines IL-4, Il-5, and IL-13 whereas anti-CD28
or IL-33 alone did not induce production of these three signals. 109 T-cells exposed to
these three signals mature and begin to proliferate exponentially.
16
85 In asthmatics, allergen-specific T-cells differentiate into the T H2 lineage. These
T-cells then begin to proliferate over the course of the next several days, and form effector and memory T-cell lineages. T H2 effector T-cells in asthma mediate many of the events described above. Memory T-cells are long-lived cells that reside in secondary lymphoid organs and respond to subsequent antigen stimulation by DCs to produce additional effector T-cells. 110 Thus, the action of dendritic cells is critical to
development of a T H2 response, and asthma for atopic individuals.
Dendritic cells in asthma
Pulmonary dendritic cells (DCs) are important for T cell differentiation into a T H2 phenotype in allergic asthma. Knockout mice lacking the co-stimulatory molecules found on DCs display impaired development of experimental asthma 111,112 For example, PD-L1 knockout mice display reduced AHR and development of a T H2 response to an OVA model of asthma, while PD-L2 knockout mice display increased
AHR.113 Thus, different co-stimulatory molecules have varying effects on the
development of a T H2 response. Recently, two subsets of dendritic cells, myeloid DC
(mDCs) and plasmacytoid DCs (pDCs) have been identified, and have opposing effects
on T-cell differentiation. 114,115 Our lab has shown that mDCs are particularly important in promoting an asthma phenotype.116
17 Under homeostatic conditions, dendritic cells reside in the lung near the
basement membrane at a density of around 600 DCs per mm 2 (in histological examinations) in the upper airways. 117 This decreases toward the lower airway. These
DCs reside just below the bronchial epithelium, and continuously sample antigen in the
airway lumen through interdigitating processes. These processes travel between
epithelial cells to gain access to antigens in the airway.102 When the airways are exposed to a danger signal, additional DCs are recruited into the airways; this recruitment peaks around 24 hours after initial exposure at nearly 1000 DCs per mm 2.117
In addition to recruited DCs, resident DCs may become activated and both begin to take
up antigen. 118 Prior to antigen uptake, DCs are referred to as immature DCs. Immature
DCs express the chemokine receptor CCR6, which binds to the chemokine CCL20. This
binding to CCL20 induces DC recruitment toward the source of chemokine secretion.
CCR6 is required for DC recruitment in the lungs in mouse models of asthma. 119
After taking up antigen, DCs mature. They stop expressing CCR6, and begin expressing CCR7. 120 This signals DCs to migrate out of the lungs or other tissues, and
into draining lymph nodes. As they mature, DCs break down the antigen they took up
into short peptide sequences that are loaded onto MHC II and transported to the cell
surface. They also begin to express co-stimulatory molecules and secrete cytokines
induced by signaling from PRRs at the site of antigen uptake. 121 Once in the lymph
18 nodes, the mature DCs present the antigen they took up on their surface to T cells. The context in which the DC took up antigen contributes to the co-stimulatory molecules expressed by the DC during antigen presentation. These co-stimulatory molecules determine the type of T cell differentiation that results from antigen uptake. 122
Figure 3. Development of a T H2 response after allergen exposure.
19
The bronchial epithelium in asthma
There are two main types of epithelial cells in the conducting airways: goblet cells and ciliated cells. Goblet cells are the main source of production for the mucous layer that coats the airways. They produce various mucin proteins, and secrete them into the airway lumen. 123,124 Mucins are large, highly glycosylated proteins that form the basic component of mucus. 125 The mucous traps particles and bacteria from the air and prevents them from reaching the lower airways and the alveoli, thereby clearing most airborne debris from the lungs. Ciliated cells continuously move this mucous layer up toward the esophagus, where it is eventually swallowed. This mechanism is called the mucociliary escalator. A thin layer of liquid, called the sol layer, resides
between the cilia and the mucous layer. This, much less viscous, layer allows the cilia to move freely. On their forward stroke, cilia strike the mucous layer and propel it forward. On the reverse stroke, the sol layer allows the cilia to release the mucous layer and prepare for another forward stroke. 126 Bronchial epithelial cells play an important
role in regulating this periciliary fluid layer.
In asthmatics, several factors contribute to increased mucous, and even mucous
plugs, in the airways. The mucous itself is produced by specialized epithelial cells such
as goblet cells in the larger airways and Clara cells in the bronchioles. In asthmatics, the
20 balance between few goblet cells and many ciliated cells shifts as ciliated cells become, or are replaced by, goblet cells. 127 This leads to an increase in mucous production, and puts greater strain on the mucociliary escalator. Additionally, the periciliary fluid layer
becomes dysregulated, leading to stalled mucous secretions. This also results in impaired clearance of mucous from the airways.128,129 Interestingly, in addition to its role in T H2 cells, IL-13 has also been shown to directly induce mucous production. 130
Ion transport in bronchial epithelial cells
As the cell type covering the surface of the airways, bronchial epithelial cells are
responsible for maintaining a balanced luminal environment. They do this by
transporting various ions from the basolateral side of the cell membrane to the apical, or
luminal, side and vice-versa. By controlling solute transport, the epithelium can also
control movement of water, and luminal pH. However, ion transport also affects other,
precisely maintained aspects of cellular homeostasis, so a number of mechanisms have
evolved to handle this complex balance. These mechanisms involve the coordinated
function of multiple different ion transporters, outlined below. The work outlined in
later chapters further explores the role of ion transporters of the bronchial epithelium in
the development of downstream immune responses and an asthma phenotype.
Understanding how these mechanisms function in the normal lung environment is
21 important to appreciating how dysregulation of these mechanisms by allergen exposure may promote airway epithelial cells to release danger signals.
Under homeostatic conditions, sodium is maintained at a concentration near 140 mM in the interstitial fluid surrounding most cells, and located at the basolateral membrane of the bronchial epithelium. Intracellular sodium in most cell types is maintained at a concentration near 15 mM. This large difference in sodium concentrations generates a gradient across the cell membrane. Like a capacitor in electronic circuits stores energy as a voltage difference across a similar type of gap, this sodium gradient stores free energy that can be used to transport other ions or solutes across the cell membrane against smaller gradients. This same mechanism exists in reverse for potassium, where intracellular potassium concentrations are maintained at around 120 mM, and interstitial fluid contains approximately 5 mM potassium. The potassium gradient is formed in part through the function of the sodium-potassium adenosine triphosphatase (Na +/K +-ATPase). Na +/K +-ATPase transports three sodium ions out of the cell for every two potassium ions it transports into the cell, supporting
both gradients. The energy to drive this transport is driven by ATP hydrolysis.
Another major ion transporter involved in cell homeostasis is the sodium-potassium- chloride cotransporter (NKCC). NKCC is a symporter that transports one sodium, one potassium, and two chloride ions into the cell for each round of transport. This
22 transport is driven by the electrochemical gradient of sodium. Maintenance of specific cellular concentrations of sodium, potassium, and chloride is important to allow the cell to respond to many different stimuli, and perform homeostatic functions, including trans-epithelial ion transport.
Ion channels transport ions across the cell membrane through passive diffusion.
Two major ion channels active in many cell types, and important in bronchial epithelial cells, are the cystic fibrosis transmembrane conductance regulator (CFTR), and the epithelial sodium channel (ENaC). In bronchial epithelial cells, CFTR resides on the apical surface and exports chloride into the lumen. In contrast, ENaC also resides on the apical surface of the epithelium, but it imports sodium from the lumen.
In addition to ion transport directly through cells (across both the apical and
basolateral membranes for epithelial cells) passive diffusion between cells can also occur. Epithelial cells maintain a barrier through close association with one another.
Tight junctions between cells bring the membranes close together, and are continuous enough to exclude large molecules from diffusing between the cells. These tight
junctions are still permeable to ions, allowing some passive movement of ions across the epithelial barrier. As previously discussed, some active proteases in allergens, such as
Der p 1, break down these tight junctions, thus disrupting the tightly regulated flow of
23 both ions and water into and out of the airway lumen. This disruption potentially impacts the electrochemical balance surrounding and within bronchial epithelial cells, and could potentiate a response from these cells.
Another class of cell junction proteins also plays a role in ion transport. Gap
junctions form between cells and allow movement of ions and some small molecules
between cells. Gap junctions are formed when two heterodimeric pore complexes, called connexons found on each adjoining cell, come together. Each connexon is a hemichannel made up of different connexin proteins. These hemichannels may be gated, allowing them to open or close based on the cell’s condition. This allows multiple cells to share similar electrochemical properties, and even second messengers, providing a more coordinated response to stimuli.
The function of multiple ion transporters working in concert is required for fluid secretion and absorption in the lumen. For example, water absorption begins with movement of sodium from the lumen, through the cell, to the basolateral membrane.
This is accomplished through ENaC on the apical surface, and Na +/K +-ATPase on the
basolateral surface (the extra intracellular potassium that results escapes through
basolateral potassium channels). Sodium transport through the cell induces paracellular transport of chloride through tight junctions. This in turn generates an
24 osmotic gradient across the epithelium, as more salt exits the airway, and drives water from the lumen into the interstitial space (figure 4).131
Fluid secretion occurs when CFTR exports chloride from the apical membrane
into the lumen. This is balanced on the basolateral surface by NKCC1 importing
additional chloride into the cell. Again, this is accompanied by passive diffusion of
sodium from the interstitial space through paracellular transport, followed by
water. 132,133 All these mechanisms work in concert to promote a balanced electrochemical environment in the airways under homeostatic conditions. However, many of these mechanisms are dysregulated in asthmatic airways.
The airways of acute asthmatics are markedly more acidic than non-asthmatic airways. 134 One mechanism by which luminal pH can be controlled is by transporting
bicarbonate (HCO 3-) into the airway lumen. Chloride-bicarbonate exchangers have
been proposed to regulate luminal pH. 135,136 Note that chloride movement is used to power bicarbonate transport. This is yet another way that careful control of ion transport and ion gradients can affect physiological changes in the airways.
25
Figure 4. Ion transport in ciliated bronchial epithelial cells.
The bronchial epithelium and immunity
Airway epithelial cells have long been recognized as aiding in immunity by maintaining the mucociliary escalator, and by their barrier function. In recent years, there has been greater appreciation for a more direct interaction between the airway epithelium and dedicated immune cells. In response to pathogen-associated molecular patterns (PAMPs), they release multiple chemokines and cytokines that help drive inflammation and clear infection. 137 For example, double-stranded RNA, an agonist for
TLR3, induced expression of IL-8 and CCL20 in the human lung epithelial cell line
26 BEAS-2B; while zymosan, an agonist for TLR2 and dectin-1, induced expression of GM-
CSF and CCL20 in these same cells.138 In asthmatics, epithelial cells release these
mediators in response to molecular patterns found within allergens, helping to drive an
inappropriate immune response. 139 As mentioned previously, airway epithelial cells release IL-33 and IL-25. These activate ILC2 cells and help drive a type-2 response, and stimulate DCs as well.106,107 In addition, the epithelium responds to signals from T H2 T-
cells. For example, the bronchial epithelial cells of asthmatics have been shown to
exhibit phosphorylated STAT-6, and they respond to IL-4 exposure by secreting IL-8. 140
Thus, in addition to their role in regulating the airway environment, bronchial epithelial
cells provide many initial signals that promote down-stream immune responses.
Role of chemokines/cytokines in allergic asthma
Cytokines are proteins released from cells that serve as intercellular signaling molecules. They differ from hormones in that they normally function at shorter distances where the target cells are generally within the microenvironment of the cell releasing the cytokine, as opposed to hormones which generally circulate systemically.
Cytokines that induce chemotaxis of leukocytes toward the chemokine source are called chemokines, because they act as chemo tactic cyto kines . Many chemokines and cytokines are critical in the development and pathogenesis of allergic asthma. Some important examples include cytokines such as IL-4, IL-5 and IL-13; and chemokines such as CCL20
27 and IL-8.89,141,142 The bronchial epithelium participates in the release of IL-8 and CCL20;
but not IL-4, IL-5, and IL-13.
Inflammatory chemokines
Bronchoalveolar lavage of asthmatic lungs reveals significant increases in
cellularity compared to non-asthmatics. 143 This same lavage fluid also contains elevated levels of a number of chemokines and cytokines that have inflammatory effects. 144 In recent years, this has led to work on a number of cytokine-directed therapies that are currently under development. 145 Some specific chemokines and cytokines that are important in the development of asthma are discussed below.
CCL20
One chemokine that is elevated in the BAL of asthmatics is CCL20. The only known receptor for CCL20 is the chemokine receptor CCR6, which also binds beta- defensins, although with much lower affinity.146,147 CCL20 was originally discovered in liver tissue and named liver and activation-regulated chemokine, or LARC. 148 It is constitutively expressed in multiple cell types,149 including epithelial cells of the skin, intestine, and airways. 150–153 In addition to this constitutive expression, production and
release of CCL20 may also be induced by various stimuli. 152,154 These include cytokines
such as TNF-α or IL-1α, as well as bacterial pathogens or other danger signals. 155 We have previously shown that bronchial epithelial cells release CCL20 when exposed to the allergen house dust mite (HDM) 156 . Indeed, this release can happen rapidly after
28 HDM exposure (Figure 5). CCL20 has also been shown to be released from bronchial epithelial cells in response to particulate matter.157
Figure 5. CCL20 release within 30 minutes of HDM exposure. (Nathan et al. 2009)
A number of studies have demonstrated that signaling of CCL20 through CCR6 contributes to the development of allergic asthma. Studies with CCR6 knockout mice showed that it is required for pulmonary inflammation after allergen challenge, and have linked it to recruitment of DCs and CD-4+ T-cells in asthmatic airways. 158–161
162 CCL20 has also been linked to the recruitment of T H17 cells, which also express
163 CCR6. As mentioned previously, T H17 cells may promote more severe, steroid
resistant asthma. Signaling through CCL20 may explain why we see elevated levels of
164 TH17 cells in asthmatic airways. Indeed, patients with steroid resistant asthma had
ten times as much CCL20 in their bronchoalveolar lavage fluid as patients with asthma
that responded to steroid treatment.165
29
In addition to its role in inflammation, CCL20, like many chemokines, is also a
166 potent antimicrobial peptide, with an LD 50 of 400 ng/mL for Escherichia coli . This
antimicrobial activity is thought to be particularly active in acidic environments 167 where dimeric CCL20 separates into monomers, thus exposing its carboxy-terminal end, which is thought to directly mediate the protein’s antimicrobial activity.168 This cell-free antimicrobial activity varies greatly depending on the microbial target (table 1) and is not limited to the chemokine CCL20 (table 2).
The antimicrobial activity of CCL20/MIP-3a against various strains of bacteria and fungi
Microorganism LD 50 (ug/mL) Gram - bacteria P. aeruginosa 1.1 M. catarrhalis 0.9 Gram + bacteria S. pyogenes 0.2 E. faecium 4.5 Fungi C. albicans 25 C. neoforman 80 Table 1. Adapted from Yang, et al. 2003.
30 The antimicrobial activity of selected chemokines Chemokine Antimicrobial activity (at 10 ug/mL) Family Member E. coli S. aureus CXC CXCL1/Gro-α ++++ + CXCL2/Gro-β +++ + CXCL3/Gro-γ +++ + CXCL6/GCP-2 – – CXCL8/IL-8 – – CXCL9/MIG ++++ ++++ CXCL10/IP-10 ++++ +++ CXCL11/I-TAC +++ +++ CXCL12/SDF-1 ++++ ++ CXCL13/BCA-1 ++++ ++ CXCL14/BRAK ++ + CX 3C CX 3CL1/fractalkine – – C XCL1/lymphotactin ++++ ++ CC CCL1/I-309 ++++ + CCL2/ MCP-1 – – CCL3/ MIP-1α – – CCL5/RANTES – – CCL7/ MCP-3 – – CCL8/ MCP-2 + – CCL11/eotaxin +++ ++ CCL13/MCP-4 +++ – CCL16/LEC – – CCL17/TARC +++ ++ CCL18/PARC ++++ +++ CCL19/ MIP-3β ++++ – CCL20/MIP-3α ++++ ++ CCL21/SLC ++ +++ CCL22/MDC ++++ +++ CCL25/TECK +++ ++ CCL27/CTAK – – Table 2. Adapted from Yang, et al. 2003.
31 Other inflammatory chemokines and cytokines
Many other chemokines have been shown to be released in the BAL of asthmatics in addition to CCL20. Among others, these chemokines include Gro-α, IL-8, and to a lesser extent IP-10. 144 These chemokines help to recruit many of the cell types seen in asthmatics. Gro-α and IL-8 recruit neutrophils by signaling through the chemokine receptors CXCR2 (for both Gro-α and IL-8) and CXCR1 (for IL-8). 169–171 IP-
10 is responsible for the recruitment of a variety of cell types, including T-cells and DCs,
through its receptor CXCR3. 172,173
Cytokines reported in the BAL of asthmatics include granulocyte macrophage colony-stimulating factor (GM-CSF), IL-1β, IL-2, IL-5, IL-6, IL-13, and TNF-α.144,174
Additionally, other cytokines that have been reported to be released by the bronchial epithelium, such as thymic stromal lymphopoietin (TSLP) and IL-33, have also been reported to play a significant role in asthma pathogenesis. 175–177 Major roles for each of these cytokines in asthma are listed in table 3. In addition to de novo production after allergic stimuli, production of many chemokines and cytokines by the bronchial epithelium has been reported under homeostatic conditions. 178 Thus a number of
32 proteins work in concert to promote a T H2 response, with many redundant signals leading to some important phenotypic changes, such as neutrophilia.
The role of cytokines active in the pathogenesis of asthma Cytokine Role in asthma GM-CSF Promotes DC maturation and differentiation. 179 IL-1β Processed from proIL-1β through inflammasome activation. Leads to production of other inflammatory cytokines, such as IL-6. 180 80,88 IL-4 Promotes the development and maintenance of T H2 cells. IL-5 Promotes eosinophilia. 91 181 IL-6 Prevents T reg development, promotes T H17 development. IL-13 Necessary for goblet cell metaplasia, 182 and the development of asthma. 93,95 183 IL-25 Produced by BECs. Promotes T H2 differentiation. 184 IL-33 Produced by BECs. Promotes T H2 differentiation. TNF-α Airway remodeling, 185 T-cell activation, 186 neutrophil and eosinophil recruitment. 187 TSLP Produced by BECs. Helps drive DC activation and development of a T H2 response. 188,189 Table 3.
Secretory lysosomes
One cellular compartment that is identified in this thesis as a major storage location for chemokines and cytokines is in lysosomes. Lysosomes are acidic organelles within the cell that serve as a primary site of digestion for various cell components. 190
The structure of lysosomes is surprisingly complex, and reflects the multiple roles they play beyond recycling of cellular components traditionally thought to be their main function. Lysosomes are highly acidic, with a pH of around 4.3.191 This acidification is
driven by the vacuolar-type H+ ATPase, or V-ATPase. 192 V-ATPase is a proton pump
33 that uses the energy from ATP hydrolysis to transport protons from the cytosol into lysosomes. V-ATPase function can be inhibited by a number of molecules, including
bafilomycin A.193 Since many of the enzymes within lysosomes require low pH to function, inhibition of V-ATPase may affect both the development of new lysosomes, and the function of existing lysosomes.
The action of V-ATPase presents an interesting physiological problem. As protons are pumped from the cytosol into the lysosome, they are moving from an environment that holds a net negative charge into an environment whose positive charge increases with each additional proton. If this mechanism acted alone, acidification would soon cease as the energy from ATP hydrolysis soon became insufficient to drive new protons into the lysosome, and therefore the low lysosomal pH could not be achieved. In recent years, a number of papers have been published showing that the charge balance problem can be overcome in lysosomes by allowing negatively charge chloride ions into the organelle. Proton-chloride exchangers, such as
CLC-7 and CLC-5, are found on lysosomes and developing endosomes, respectively.194,195 These exchangers use the free energy from allowing the protons to escape through the transporter to drive chloride movement into the lysosome (figure 6).
The net result of the action of both V-ATPase and Cl -/H + exchanger action is an acidic lysosome with a high chloride concentration. 196 This high chloride concentration
34 further contributes to cytotoxic killing by lysosomes. These structural components of lysosomes will be explored in later chapters as the localization of intracellular chemokines and cytokines is investigated.
A B
Figure 6. (A) Diagram of lysosome, showing the electron-dense protein core, ion transporters involved in acidification, the membrane protein lamp-1, and membrane voltage (V) that is balanced by chloride ions. (B) Double labeling of Lamp-1 (10 nm gold) and granzyme B (15 nm gold) in cytotoxic T lymphocytes. Granzyme B labeling is mainly present within the electron-dense core. Lamp-1 is predominantly present at the outer membrane of the granules. The plasma membrane is labeled (p). Adapted from Peters et al 1991.
Many proteins are found within lysosomes in various cell types, including some chemokines and cytokines in cells of the hematopoietic lineage. These proteins are not solely associated with lysosomal localization, however. Some lysosome-specific
35 proteins include proteases such as cathepsins, and membrane-associated proteins such as LAMP-1.197 The aggressive proteases found in lysosomes are inactive at the higher
pH within the cell, or at lower chloride concentrations, 198,199 which is a protective
mechanism against inappropriate digestion during enzyme transport. Not all proteins
found in lysosomes are immune to the caustic environment there. These proteins are
targeted to a protective structure in lysosomes that, when visualized by electron
microscopy, appears as an electron dense core within the lysosome. 200 This core is made up of proteins bound to sulfated proteoglycans that help to protect the proteins from degradation. 201–205 In secretory lysosomes these complexes of proteins and
proteoglycans are released upon exocytosis. 206,207
Secretory lysosomes are a specific class of Lamp-1—containing lysosomal
vesicles that are capable of fusing with the plasma membrane and releasing their
content into the extracellular milieu. 200 Secretory lysosomes have been studied
predominantly in cells of the hematopoietic lineage, but they have been identified in
other cell types as well – including epithelial cells. 208 In these various cell types, secretory lysosomes have been shown to perform a diverse variety of functions. For example, skin pigment-producing cells, called melanocytes, produce lysosome-like organelles, called melanosomes, through which they transfer pigment to the surrounding epithelial cells. 209 A specialized type of macrophage, called osteoclasts,
36 use enzymes stored in secretory lysosomes to break down and recycle bone matrix. 210
And in many specialized immune cell types, cytotoxic and antimicrobial peptides and
mediators are stored in secretory lysosomes. These can then be released in a context-
dependent manner. For example, upon cross-linking of IgE mast cells release histamine
and other mediators that can attack invading pathogens.211 Additionally, natural killer cells and cytotoxic T cells store granzyme and perforin, which they use to kill specific target cells that may be cancerous or virally infected.200 Thus, secretory lysosomes operate in a variety of different cell types, and their release allows for context-specific responses to a variety of stimuli.
Finally, in some cell types, such as neutrophils, multiple types of secretory lysosomes have been identified within a single cell. 212 This allows the cell to respond to different stimuli by releasing the contents of different lysosomes. 213 Many different chemokines and cytokines have been shown to reside within these vesicles. For example, the chemokine IL-8 is stored and secreted from Weibel-palade vesicles in endothelial cells. 214 Although much research has contributed to a better understanding
of the complex loading and release mechanisms of different types of secretory
lysosomes in cells of the hematopoietic lineage, identification of potential secretory
lysosomes and their contents in epithelial cell types, such as the bronchial epithelium,
has yet to be explored.
37
Syndecans and heparin sulfate proteoglycans
Another potential storage site for chemokines and cytokines is on structural molecules attached to the cell membrane on the extracellular side. On the baso-lateral surface of the cell, chemokines and other proteins can be stored on syndecans. 215
Syndecans are proteoglycans, where the trans-membrane protein stalk is modified with
chains of polysaccharides called glycans. The syndecan protein consists of a small
intracellular domain, a trans-membrane domain, and a larger extracellular domain
where the glycan modifications are attached. The glycan linkages on syndecans
contain of heparin sulfate (HS). These side chains bind to other elements of the
extracellular matrix and help anchor the cell. 216
In addition to their structural role, syndecans also participate in cell signaling.
Many chemokines, cytokines, and growth factors bind to HS, like that found on syndecan, including: interferon-γ, epidermal growth factor, hepatocyte growth factor, vascular endothelial growth factor, fibroblast growth factor family members, IL-1α, IL-
1β, IL-2, IL-3, IL-4, IL-6, IL-7, IL-8, GM-CSF, and Gro-α.217–227 When the side chains are
released, the bound chemokines and cytokines are released as well, providing a rapid
storage and release mechanism on the cell surface.228 Near the base of the extracellular domain of syndecans are cleavage sites where syndecans may be cut by matrix
38 metalloproteinases. 229–231 In addition to cleavage of the protein stalk, the bonds holding
side chains themselves together may be broken by a variety of mechanisms, including
hydrolysis by nitric oxide, and degradation by release of heparanase which is secreted
from lysosomes. 232 Through cleavage of the protein stalk or by direct cleavage of HS
the proteins associated with the side chains of syndecan may be released from the cell
surface. In the case of chemokines, like IL-8, this release forms a gradient that can help
recruit inflammatory cells, such as neutrophils.
In asthma, HS in general and syndecans in particular play an important role in
the development of an allergic response. Mice deficient in HS develop less severe
airway responses in an ova-induced model of asthma.233 In contrast, mice deficient in
syndecan-1 displayed increased airway inflammation in a mold model of asthma. Thus
it appears that the specific agonist used to induce an allergic response determines the
role of syndecans and HS in either promoting or attenuating the response.
39
Figure 7. Syndecan structure and shedding.
Summary
Asthma is a chronic inflammatory disease affecting millions of people worldwide. In affected individuals, it causes narrowing of the airways through airway smooth muscle hypertrophy and contraction, increased airway mucous, inflammation, and eventually permanent changes in airway structure brought about by fibrosis. These features are primarily mediated through type-2 responses, whether through adaptive
TH2 cell responses or through the newly identified ILCs. The TH2 response in asthmatics develops as DCs in the lungs take up allergens in the lungs and
40 subsequently drive the development of TH2 cells. Dendritic cells receive signals to migrate into the sub-epithelial space and take up antigen by chemokines and cytokines that may be secreted by bronchial epithelial cells. Many allergens have been shown to induce the secretion of these chemokines and cytokines into the lavage fluid of asthmatics.
Airway epithelial cells manage the mucociliary escalator, airway pH, and fluid in the airways. Much of this regulation is maintained by balancing ion transport across the epithelium. Additionally, airway epithelial cells release chemokines and cytokines in response to allergens and other agonists.
Previously, we and others found that CCL20, an important chemokine for immature dendritic cell recruitment, is strongly released directly from bronchial epithelial cells after HDM stimulation. However, the mechanisms driving production and release of protein mediators from the bronchial epithelium are poorly understood.
Many mechanisms have been identified in other cell types for the storage and secretion of chemokines and cytokines. These include storage and secretion from intracellular compartments such as secretory vesicles and secretory lysosomes. In addition, chemokines and cytokines may be stored on the cell surface on syndecans. They are then released by enzymes that degrade the heparin sulfate side chains, or protein stalks
41 found on syndecans. Finally, chemokines and cytokines may be produced and secreted de novo after cell stimulation.
Understanding the mechanism and kinetics of chemokine release in general, and
CCL20 release specifically, from bronchial epithelial cells will help in our understanding of the initial processes driving sensitization. In addition, this may lead to better methods of intervention in the treatment of asthmatics. We hypothesize that rapid CCL20 release from bronchial epithelial cells is driven through the release of pre-formed extracellular and intracellular stores.
The specific aims of this dissertation are:
1. Determine the kinetics and localization of CCL20 chemokine release from BECs
from both intracellular and extracellular stores.
2. Determine the signaling pathways that lead to mobilization and inhibition of
CCL20 stores.
3. Identify the allergen selectivity of CCL20 release.
4. Identify other chemokines and cytokines that are stored in BECs and rapidly
released after allergen stimulation.
42 References
1. GINA Report: Global strategy for asthma management and prevention . (Global Initiative for Asthma, 2011). at
43 23. Strachan, D. P. Hay fever, hygiene, and household size. BMJ 299, 1259–1260 (1989). 24. Worgall, T. S. et al. Impaired sphingolipid synthesis in the respiratory tract induces airway hyperreactivity. Sci Transl Med 5, 186ra67 (2013). 25. Li, X. et al. Genome-wide association studies of asthma indicate opposite immunopathogenesis direction from autoimmune diseases. J. Allergy Clin. Immunol. 130, 861–868.e7 (2012). 26. Defective intracellular transport and processing of CFTR is the molecular basis of most cystic fibrosis. Cell 63, 827–834 (1990). 27. Moore, W. C. et al. Identification of Asthma Phenotypes Using Cluster Analysis in the Severe Asthma Research Program. Am J Respir Crit Care Med 181, 315–323 (2010). 28. Van der Heide, S., De Monchy, J. G., De Vries, K., Dubois, A. E. & Kauffman, H. F. Seasonal differences in airway hyperresponsiveness in asthmatic patients: relationship with allergen exposure and sensitization to house dust mites. Clin Exp Allergy 27, 627–33 (1997). 29. Dezateux, C., Stocks, J., Dundas, I. & Fletcher, M. E. Impaired Airway Function and Wheezing in Infancy The Influence of Maternal Smoking and a Genetic Predisposition to Asthma. Am. J. Respir. Crit. Care Med. 159, 403–410 (1999). 30. Claudio, L., Tulton, L., Doucette, J. & Landrigan, P. J. Socioeconomic factors and asthma hospitalization rates in New York City. J Asthma 36, 343–350 (1999). 31. Kozyrskyj, A. L., Ernst, P. & Becker, A. B. Increased Risk of Childhood Asthma From Antibiotic Use in Early Life*. Chest 131, 1753–1759 (2007). 32. Gavett, S. H. & Koren, H. S. The role of particulate matter in exacerbation of atopic asthma. Int. Arch. Allergy Immunol. 124, 109–112 (2001). 33. Jarvis, D. L., Leaderer, B. P., Chinn, S. & Burney, P. G. Indoor Nitrous Acid and Respiratory Symptoms and Lung Function in Adults. Thorax 60, 474–479 (2005). 34. Yeatts, K. et al. Coarse particulate matter (PM2.5-10) affects heart rate variability, blood lipids, and circulating eosinophils in adults with asthma. Environ. Health Perspect. 115, 709–714 (2007). 35. Van Nimwegen, F. A. et al. Mode and place of delivery, gastrointestinal microbiota, and their influence on asthma and atopy. J Allergy Clin Immunol doi:S0091-6749(11)01148-1 [pii] 10.1016/j.jaci.2011.07.027 36. Ege, M. J. et al. Exposure to environmental microorganisms and childhood asthma. N Engl J Med 364, 701–9 37. Hilty, M. et al. Disordered microbial communities in asthmatic airways. PLoS One 5, e8578 (2010). 38. Gelber, L. E. et al. Sensitization and Exposure to Indoor Allergens as Risk Factors for Asthma Among Patients Presenting to Hospital. Am. J. Respir. Crit. Care Med. 147, 573– 578 (1993). 39. Sporik, R., Holgate, S. T., Platts-Mills, T. A. & Cogswell, J. J. Exposure to house-dust mite allergen (Der p I) and the development of asthma in childhood. A prospective study. N. Engl. J. Med. 323, 502–507 (1990). 40. Wahn, U. et al. Indoor allergen exposure is a risk factor for sensitization during the first three years of life. Journal of Allergy and Clinical Immunology 99, 763–769 (1997). 41. Huss, K. et al. House dust mite and cockroach exposure are strong risk factors for positive allergy skin test responses in the Childhood Asthma Management Program*. Journal of Allergy and Clinical Immunology 107, 48–54 (2001).
44 42. Sandhu, M. S. & Casale, T. B. The role of vitamin D in asthma. Ann. Allergy Asthma Immunol. 105, 191–199; quiz 200–202, 217 (2010). 43. Simpson, B. M. et al. NAC Manchester Asthma and Allergy Study (NACMAAS): risk factors for asthma and allergic disorders in adults. Clin Exp Allergy 31, 391–399 (2001). 44. Miraglia del Giudice, M. et al. Atopy and house dust mite sensitization as risk factors for asthma in children. Allergy 57, 169–172 (2008). 45. Feinberg, A. R., Feinberg, S. M. & Benaim-Pinto, C. Asthma and rhinitis from insect allergens: I. Clinical importance. Journal of Allergy 27, 437–444 (1956). 46. Bernton, H. S., McMahon, T. F. & Brown, H. Cockroach asthma. British Journal of Diseases of the Chest 66, 61–66 (1972). 47. Arbes Jr., S. J. et al. Dog allergen (Can f 1) and cat allergen (Fel d 1) in US homes: Results from the National Survey of Lead and Allergens in Housing. Journal of Allergy and Clinical Immunology 114, 111–117 (2004). 48. Gent, J. F. et al. Levels of household mold associated with respiratory symptoms in the first year of life in a cohort at risk for asthma. Environ. Health Perspect. 110, A781–786 (2002). 49. Thorn, J., Brisman, J. & Torén, K. Adult ‐onset asthma is associated with self ‐reported mold or environmental tobacco smoke exposures in the home. Allergy 56, 287–292 (2002). 50. Citron, K. M., Frankland, A. W. & Sinclair, J. D. Inhalation Tests of Bronchial Hypersensitivity in Pollen Asthma. Thorax 13, 229–232 (1958). 51. Gough, L., Schulz, O., Sewell, H. F. & Shakib, F. The Cysteine Protease Activity of the Major Dust Mite Allergen Der P 1 Selectively Enhances the Immunoglobulin E Antibody Response. J Exp Med 190, 1897–1902 (1999). 52. Page, K., Strunk, V. S. & Hershenson, M. B. Cockroach proteases increase IL-8 expression in human bronchial epithelial cells via activation of protease-activated receptor (PAR)–2 and extracellular-signal-regulated kinase. Journal of Allergy and Clinical Immunology 112, 1112–1118 (2003). 53. Paton, J. B. Pollen and Pollen Enzymes. American Journal of Botany 8, 471–501 (1921). 54. Gough, L., Sewell, H. F. & Shakib, F. The proteolytic activity of the major dust mite allergen Der p 1 enhances the IgE antibody response to a bystander antigen. Clin. Exp. Allergy 31, 1594–1598 (2001). 55. Wan, H. et al. Der p 1 facilitates transepithelial allergen delivery by disruption of tight junctions. Journal of Clinical Investigation 104, 123–133 (1999). 56. Vinhas, R. et al. Pollen proteases compromise the airway epithelial barrier through degradation of transmembrane adhesion proteins and lung bioactive peptides. Allergy 66, 1088–1098 (2011). 57. Matzinger, P. Friendly and dangerous signals: is the tissue in control? Nat. Immunol. 8, 11– 13 (2007). 58. Neville, A. C., Parry, D. A. & Woodhead-Galloway, J. The Chitin Crystallite in Arthropod Cuticle. J Cell Sci 21, 73–82 (1976). 59. Wills-Karp, M. & Karp, C. L. Chitin checking--novel insights into asthma. N. Engl. J. Med. 351, 1455–1457 (2004). 60. Valerio, C., Murray, P., Arlian, L. & Slater, J. Bacterial 16S ribosomal DNA in house dust mite cultures. Journal of Allergy and Clinical Immunology 116, 1296–1300 (2005). 61. Michel, O. et al. Severity of Asthma Is Related to Endotoxin in House Dust. Am. J. Respir. Crit. Care Med. 154, 1641–1646 (1996).
45 62. Hammad, H. et al. House dust mite allergen induces asthma via Toll-like receptor 4 triggering of airway structural cells. Nature Medicine 15, 410–416 (2009). 63. James, A. & Carroll, N. Airway Smooth Muscle in Health and Disease; Methods of Measurement and Relation to Function. Eur Respir J 15, 782–789 (2000). 64. PhD, W. F. B. M. & MD, E. L. B. Medical Physiology, Updated Edition: With STUDENT CONSULT Online Access, 1e . (Saunders, 2004). 65. Pavia, D., Bateman, J. R., Sheahan, N. F., Agnew, J. E. & Clarke, S. W. Tracheobronchial Mucociliary Clearance in Asthma: Impairment During Remission. Thorax 40, 171–175 (1985). 66. Brewster, C. E. P. et al. Myofibroblasts and Subepithelial Fibrosis in Bronchial Asthma. Am. J. Respir. Cell Mol. Biol. 3, 507–511 (1990). 67. Elias, J. A., Zhu, Z., Chupp, G. & Homer, R. J. Airway remodeling in asthma. Journal of Clinical Investigation 104, 1001–1006 (1999). 68. Dunnill, M. S., Massarella, G. R. & Anderson, J. A. A Comparison of the Quantitative Anatomy of the Bronchi in Normal Subjects, in Status Asthmaticus, in Chronic Bronchitis, and in Emphysema. Thorax 24, 176–179 (1969). 69. Cutz, E., Levison, H. & Cooper, D. M. Ultrastructure of airways in children with asthma. Histopathology 2, 407–421 (1978). 70. Ebina, M., Takahashi, T., Chiba, T. & Motomiya, M. Cellular Hypertrophy and Hyperplasia of Airway Smooth Muscles Underlying Bronchial Asthma: A 3-D Morphometric Study. Am. J. Respir. Crit. Care Med. 148, 720–726 (1993). 71. Woodruff, P. G. et al. Hyperplasia of Smooth Muscle in Mild to Moderate Asthma Without Changes in Cell Size or Gene Expression. Am. J. Respir. Crit. Care Med. 169, 1001–1006 (2004). 72. Schaafsma, D. et al. Allergic sensitization enhances the contribution of Rho ‐kinase to airway smooth muscle contraction. British Journal of Pharmacology 143, 477–484 (2009). 73. Tschumperlin, D. J., Shively, J. D., Kikuchi, T. & Drazen, J. M. Mechanical Stress Triggers Selective Release of Fibrotic Mediators from Bronchial Epithelium. Am. J. Respir. Cell Mol. Biol. 28, 142–149 (2003). 74. Montefort, S. et al. Bronchial biopsy evidence for leukocyte infiltration and upregulation of leukocyte-endothelial cell adhesion molecules 6 hours after local allergen challenge of sensitized asthmatic airways. Journal of Clinical Investigation 93, 1411–1421 (1994). 75. Lacoste, J.-Y. et al. Eosinophilic and neutrophilic inflammation in asthma, chronic bronchitis, and chronic obstructive pulmonary disease. Journal of Allergy and Clinical Immunology 92, 537–548 (1993). 76. Bousquet, J. et al. Eosinophilic inflammation in asthma. N. Engl. J. Med. 323, 1033–1039 (1990). 77. Bradley, B. L. et al. Eosinophils, T-lymphocytes, mast cells, neutrophils, and macrophages in bronchial biopsy specimens from atopic subjects with asthma: Comparison with biopsy specimens from atopic subjects without asthma and normal control subjects and relationship to bronchial hyperresponsiveness. Journal of Allergy and Clinical Immunology 88, 661–674 (1991). 78. Bousquet, J., Jeffery, P. K., Busse, W. W., Johnson, M. & Vignola, A. M. Asthma From Bronchoconstriction to Airways Inflammation and Remodeling. Am. J. Respir. Crit. Care Med. 161, 1720–1745 (2000).
46 79. Wenzel, S. E. et al. Evidence That Severe Asthma Can Be Divided Pathologically into Two Inflammatory Subtypes with Distinct Physiologic and Clinical Characteristics. Am. J. Respir. Crit. Care Med. 160, 1001–1008 (1999). 80. Murphy, K. Janeway’s Immunobiology (Immunobiology: The Immune System . (Garland Science, 2011). 81. Guermonprez, P., Valladeau, J., Zitvogel, L., Théry, C. & Amigorena, S. Antigen Presentation and T Cell Stimulation by Dendritic Cells. Annual Review of Immunology 20, 621–667 (2002). 82. Bluestone, J. A., Mackay, C. R., O’Shea, J. J. & Stockinger, B. The functional plasticity of T cell subsets. Nature Reviews Immunology 9, 811–816 (2009). 83. Kallmann, B. A. et al. Systemic Bias of Cytokine Production Toward Cell-Mediated Immune Regulation in IDDM and Toward Humoral Immunity in Graves’ Disease. Diabetes 46, 237– 243 (1997). 84. Ying, S., Durham, S. R., Corrigan, C. J., Hamid, Q. & Kay, A. B. Phenotype of Cells Expressing mRNA for TH2-Type (interleukin 4 and Interleukin 5) and TH1-Type (interleukin 2 and Interferon Gamma) Cytokines in Bronchoalveolar Lavage and Bronchial Biopsies from Atopic Asthmatic and Normal Control Subjects. Am. J. Respir. Cell Mol. Biol. 12, 477–487 (1995). 85. Robinson, D. S. et al. Predominant TH2-like bronchoalveolar T-lymphocyte population in atopic asthma. N. Engl. J. Med. 326, 298–304 (1992). 86. Robinson, D. et al. Activation of CD4+ T cells, increased TH2-type cytokine mRNA expression, and eosinophil recruitment in bronchoalveolar lavage after allergen inhalation challenge in patients with atopic asthma. Journal of Allergy and Clinical Immunology 92, 313–324 (1993). 87. Ying, S. et al. Expression of IL-4 and IL-5 mRNA and Protein Product by CD4+ and CD8+ T Cells, Eosinophils, and Mast Cells in Bronchial Biopsies Obtained from Atopic and Nonatopic (intrinsic) Asthmatics. J Immunol 158, 3539–3544 (1997). 88. Zheng, W. & Flavell, R. A. The Transcription Factor GATA-3 Is Necessary and Sufficient for Th2 Cytokine Gene Expression in CD4 T Cells. Cell 89, 587–596 (1997). 89. Chung, K. F. & Barnes, P. J. Cytokines in Asthma. Thorax 54, 825–857 (1999). 90. Oettgen, H. C. & Geha, R. S. IgE in asthma and atopy: cellular and molecular connections. Journal of Clinical Investigation 104, 829–835 (1999). 91. Foster, P. S., Hogan, S. P., Ramsay, A. J., Matthaei, K. I. & Young, I. G. Interleukin 5 Deficiency Abolishes Eosinophilia, Airways Hyperreactivity, and Lung Damage in a Mouse Asthma Model. J Exp Med 183, 195–201 (1996). 92. Whittaker, L. et al. Interleukin-13 mediates a fundamental pathway for airway epithelial mucus induced by CD4 T cells and interleukin-9. Am J Respir Cell Mol Biol 27, 593–602 (2002). 93. Wills-Karp, M. et al. Interleukin-13: Central Mediator of Allergic Asthma. Science 282, 2258–2261 (1998). 94. Grünig, G. et al. Requirement for IL-13 Independently of IL-4 in Experimental Asthma. Science 282, 2261–2263 (1998). 95. Walter, D. M. et al. Critical Role for IL-13 in the Development of Allergen-Induced Airway Hyperreactivity. J Immunol 167, 4668–4675 (2001). 96. Spits, H. & Santo, J. P. D. The expanding family of innate lymphoid cells: regulators and effectors of immunity and tissue remodeling. Nature Immunology 12, 21–27 (2011).
47 97. Neill, D. R. et al. Nuocytes represent a new innate effector leukocyte that mediates type-2 immunity. Nature 464, 1367–1370 (2010). 98. Saenz, S. A. et al. IL25 elicits a multipotent progenitor cell population that promotes T(H)2 cytokine responses. Nature 464, 1362–1366 (2010). 99. Price, A. E. et al. Systemically dispersed innate IL-13–expressing cells in type 2 immunity. PNAS (2010). doi:10.1073/pnas.1003988107 100. Lajoie, S. et al. Complement-mediated regulation of the IL-17A axis is a central genetic determinant of the severity of experimental allergic asthma. Nat Immunol 11, 928–35 (2010). 101. Stoll, S., Delon, J., Brotz, T. M. & Germain, R. N. Dynamic Imaging of T Cell-Dendritic Cell Interactions in Lymph Nodes. Science 296, 1873–1876 (2002). 102. Banchereau, J. & Steinman, R. M. Dendritic cells and the control of immunity. Nature 392, 245–252 (1998). 103. Burastero, S. E. et al. Increased expression of the CD80 accessory molecule by alveolar macrophages in asthmatic subjects and its functional involvement in allergen presentation to autologous TH2 lymphocytes. Journal of Allergy and Clinical Immunology 103, 1136–1142 (1999). 104. Deng, J.-M. et al. Effects of allergen inhalation and oral glucocorticoid on concentrations of serum-soluble CD86 in allergic asthmatics. Clinical Immunology 115, 178–183 (2005). 105. Hoshino, A. et al. Critical role for OX40 ligand in the development of pathogenic Th2 cells in a murine model of asthma. European Journal of Immunology 33, 861–869 (2003). 106. Rank, M. A. et al. IL-33–activated dendritic cells induce an atypical TH2-type response. Journal of Allergy and Clinical Immunology 123, 1047–1054 (2009). 107. Wang, Y.-H. et al. IL-25 augments type 2 immune responses by enhancing the expansion and functions of TSLP-DC–activated Th2 memory cells. J Exp Med 204, 1837–1847 (2007). 108. Rochman, I., Watanabe, N., Arima, K., Liu, Y.-J. & Leonard, W. J. Cutting Edge: Direct Action of Thymic Stromal Lymphopoietin on Activated Human CD4+ T Cells. J Immunol 178, 6720–6724 (2007). 109. Schmitz, J. et al. IL-33, an Interleukin-1-like Cytokine that Signals via the IL-1 Receptor- Related Protein ST2 and Induces T Helper Type 2-Associated Cytokines. Immunity 23, 479– 490 (2005). 110. Sallusto, F., Geginat, J. & Lanzavecchia, A. Central Memory and Effector Memory T Cell Subsets: Function, Generation, and Maintenance. Annual Review of Immunology 22, 745–763 (2004). 111. Van Rijt, L. S. et al. Essential role of dendritic cell CD80/CD86 costimulation in the induction, but not reactivation, of TH2 effector responses in a mouse model of asthma. Journal of Allergy and Clinical Immunology 114, 166–173 (2004). 112. Mark, D. A. et al. B7-1 (CD80) and B7-2 (CD86) Have Complementary Roles in Mediating Allergic Pulmonary Inflammation and Airway Hyperresponsiveness. Am. J. Respir. Cell Mol. Biol. 22, 265–271 (2000). 113. Akbari, O. et al. PD-L1 and PD-L2 modulate airway inflammation and iNKT-cell- dependent airway hyperreactivity in opposing directions. Mucosal Immunol 3, 81–91 (2009). 114. Barchet, W., Blasius, A., Cella, M. & Colonna, D. M. Plasmacytoid dendritic cells. Immunol Res 32, 75–83 (2005). 115. Lambrecht, B. N. et al. Myeloid dendritic cells induce Th2 responses to inhaled antigen, leading to eosinophilic airway inflammation. Journal of Clinical Investigation 106, 551–559 (2000).
48 116. Lewkowich, I. P. et al. Allergen uptake, activation, and IL-23 production by pulmonary myeloid DCs drives airway hyperresponsiveness in asthma-susceptible mice. PLoS One 3, e3879 (2008). 117. Schon-Hegrad, M. A., Oliver, J., McMenamin, P. G. & Holt, P. G. Studies on the Density, Distribution, and Surface Phenotype of Intraepithelial Class II Major Histocompatibility Complex Antigen (Ia)-Bearing Dendritic Cells (DC) in the Conducting Airways. J Exp Med 173, 1345–1356 (1991). 118. Holt, P. G., Haining, S., Nelson, D. J. & Sedgwick, J. D. Origin and Steady-State Turnover of Class II MHC-Bearing Dendritic Cells in the Epithelium of the Conducting Airways. J Immunol 153, 256–261 (1994). 119. Osterholzer, J. J. et al. CCR2 and CCR6, but Not Endothelial Selectins, Mediate the Accumulation of Immature Dendritic Cells Within the Lungs of Mice in Response to Particulate Antigen. J Immunol 175, 874–883 (2005). 120. Sallusto, F. et al. Rapid and coordinated switch in chemokine receptor expression during dendritic cell maturation. European Journal of Immunology 28, 2760–2769 (1998). 121. Sioud, M. & Fløisand, Y. TLR agonists induce the differentiation of human bone marrow CD34+ progenitors into CD11c+ CD80/86+ DC capable of inducing a Th1-type response. European Journal of Immunology 37, 2834–2846 (2007). 122. Bellou, A. & Finn, P. W. Costimulation: critical pathways in the immunologic regulation of asthma. Curr Allergy Asthma Rep 5, 149–154 (2005). 123. Rogers, D. F. Airway Goblet Cells: Responsive and Adaptable Front-Line Defenders. Eur Respir J 7, 1690–1706 (1994). 124. Lumsden, A. B., McLean, A. & Lamb, D. Goblet and Clara Cells of Human Distal Airways: Evidence for Smoking Induced Changes in Their Numbers. Thorax 39, 844–849 (1984). 125. Rogers, D. F. Physiology of airway mucus secretion and pathophysiology of hypersecretion. Respir Care 52, 1134–1146; discussion 1146–1149 (2007). 126. Wanner, A., Salathé, M. & O’Riordan, T. G. Mucociliary Clearance in the Airways. Am. J. Respir. Crit. Care Med. 154, 1868–1902 (1996). 127. Fahy, J. V. Goblet Cell and Mucin Gene Abnormalities in Asthma*. Chest 122, 320S– 326S (2002). 128. Houtmeyers, E., Gosselink, R., Gayan ‐Ramirez, G. & Decramer, M. Regulation of mucociliary clearance in health and disease. European Respiratory Journal 13, 1177–1188 (2001). 129. Randell, S. H. & Boucher, R. C. Effective Mucus Clearance Is Essential for Respiratory Health. Am. J. Respir. Cell Mol. Biol. 35, 20–28 (2006). 130. Kuperman, D. A. et al. Direct effects of interleukin-13 on epithelial cells cause airway hyperreactivity and mucus overproduction in asthma. Nat Med 8, 885–889 (2002). 131. Palmer, L. G. & Andersen, O. S. The Two-Membrane Model of Epithelial Transport: Koefoed-Johnsen and Ussing (1958). J Gen Physiol 132, 607–612 (2008). 132. Zabner, J., Smith, J. J., Karp, P. H., Widdicombe, J. H. & Welsh, M. J. Loss of CFTR Chloride Channels Alters Salt Absorption by Cystic Fibrosis Airway Epithelia In Vitro. Molecular Cell 2, 397–403 (1998). 133. Barrett, K. E. & Keely, S. J. Chloride secretion by the intestinal epithelium: molecular basis and regulatory aspects. Annu. Rev. Physiol. 62, 535–572 (2000).
49 134. Ricciardolo, F. L. M., Gaston, B. & Hunt, J. Acid stress in the pathology of asthma. J. Allergy Clin. Immunol. 113, 610–619 (2004). 135. Stetson, D. L., Beauwens, R., Palmisano, J., Mitchell, P. P. & Steinmetz, P. R. A double- membrane model for urinary bicarbonate secretion. Am J Physiol Renal Physiol 249, F546– F552 (1985). 136. Newmark, H. L. & Lupton, J. R. Determinants and consequences of colonic luminal pH: Implications for colon cancer. Nutrition and Cancer 14, 161–173 (1990). 137. Noah, T. L. & Becker, S. Respiratory Syncytial Virus-Induced Cytokine Production by a Human Bronchial Epithelial Cell Line. Am J Physiol Lung Cell Mol Physiol 265, L472–L478 (1993). 138. Sha, Q., Truong-Tran, A. Q., Plitt, J. R., Beck, L. A. & Schleimer, R. P. Activation of Airway Epithelial Cells by Toll-Like Receptor Agonists. American Journal of Respiratory Cell and Molecular Biology 31, 358–364 (2004). 139. Fujii, T. et al. Particulate Matter Induces Cytokine Expression in Human Bronchial Epithelial Cells. Am. J. Respir. Cell Mol. Biol. 25, 265–271 (2001). 140. Mullings, R. E. et al. Signal transducer and activator of transcription 6 (STAT-6) expression and function in asthmatic bronchial epithelium. Journal of Allergy and Clinical Immunology 108, 832–838 (2001). 141. Velazquez, J. & Teran, L. Chemokines and Their Receptors in the Allergic Airway Inflammatory Process. Clinical Reviews in Allergy and Immunology 41, 76–88 (2011). 142. Hershey, G. K., Friedrich, M. F., Esswein, L. A., Thomas, M. L. & Chatila, T. A. The association of atopy with a gain-of-function mutation in the alpha subunit of the interleukin- 4 receptor. N. Engl. J. Med. 337, 1720–1725 (1997). 143. Kelly, C. et al. Number and activity of inflammatory cells in bronchoalveolar lavage fluid in asthma and their relation to airway responsiveness. Thorax 43, 684–692 (1988). 144. Fitzpatrick, A. M., Higgins, M., Holguin, F., Brown, L. A. S. & Teague, W. G. The molecular phenotype of severe asthma in children. Journal of Allergy and Clinical Immunology 125, 851–857.e18 (2010). 145. Walsh, G. M. Novel Cytokine-directed Therapies for Asthma. Discovery Medicine 11, 283–291 (2011). 146. Yang, D. et al. Β-Defensins: Linking Innate and Adaptive Immunity Through Dendritic and T Cell CCR6. Science 286, 525–528 (1999). 147. Hoover, D. M. et al. The Structure of Human Macrophage Inflammatory Protein- 3α/CCL20 LINKING ANTIMICROBIAL AND CC CHEMOKINE RECEPTOR-6- BINDING ACTIVITIES WITH HUMAN Β-DEFENSINS. J. Biol. Chem. 277, 37647– 37654 (2002). 148. Hieshima, K. et al. Molecular Cloning of a Novel Human CC Chemokine Liver and Activation-Regulated Chemokine (LARC) Expressed in Liver CHEMOTACTIC ACTIVITY FOR LYMPHOCYTES AND GENE LOCALIZATION ON CHROMOSOME 2. J. Biol. Chem. 272, 5846–5853 (1997). 149. Tanaka, Y. et al. Selective expression of liver and activation-regulated chemokine (LARC) in intestinal epithelium in mice and humans. European Journal of Immunology 29, 633–642 (1999). 150. Rangel-Moreno, J. et al. Role of CXC Chemokine Ligand 13, CC Chemokine Ligand (CCL) 19, and CCL21 in the Organization and Function of Nasal-Associated Lymphoid Tissue. J Immunol 175, 4904–4913 (2005).
50 151. Nakayama, T. et al. Inducible Expression of a CC Chemokine Liver- and Activation- Regulated Chemokine (LARC)/Macrophage Inflammatory Protein (MIP)-3α/CCL20 by Epidermal Keratinocytes and Its Role in Atopic Dermatitis. Int. Immunol. 13, 95–103 (2001). 152. Izadpanah, A., Dwinell, M. B., Eckmann, L., Varki, N. M. & Kagnoff, M. F. Regulated MIP-3α/CCL20 Production by Human Intestinal Epithelium: Mechanism for Modulating Mucosal Immunity. Am J Physiol Gastrointest Liver Physiol 280, G710–G719 (2001). 153. Starner, T. D., Barker, C. K., Jia, H. P., Kang, Y. & McCray, P. B. CCL20 Is an Inducible Product of Human Airway Epithelia with Innate Immune Properties. Am. J. Respir. Cell Mol. Biol. 29, 627–633 (2003). 154. Kwon, J. H., Keates, S., Bassani, L., Mayer, L. F. & Keates, A. C. Colonic Epithelial Cells Are a Major Site of Macrophage Inflammatory Protein 3 α (MIP-3α) Production in Normal Colon and Inflammatory Bowel Disease. Gut 51, 818–826 (2002). 155. Fujiie, S. et al. Proinflammatory Cytokines Induce Liver and Activation-Regulated Chemokine/Macrophage Inflammatory Protein-3α/CCL20 in Mucosal Epithelial Cells Through NF-κB. Int. Immunol. 13, 1255–1263 (2001). 156. Nathan, A. T., Peterson, E. A., Chakir, J. & Wills-Karp, M. Innate immune responses of airway epithelium to house dust mite are mediated through beta-glucan-dependent pathways. J. Allergy Clin. Immunol. 123, 612–618 (2009). 157. Reibman, J., Hsu, Y., Chen, L. C., Bleck, B. & Gordon, T. Airway Epithelial Cells Release MIP-3α/CCL20 in Response to Cytokines and Ambient Particulate Matter. Am. J. Respir. Cell Mol. Biol. 28, 648–654 (2003). 158. Lukacs, N. W., Prosser, D. M., Wiekowski, M., Lira, S. A. & Cook, D. N. Requirement for the Chemokine Receptor Ccr6 in Allergic Pulmonary Inflammation. J Exp Med 194, 551–556 (2001). 159. Lundy, S. K. et al. Attenuation of Allergen-Induced Responses in CCR6–/– Mice Is Dependent Upon Altered Pulmonary T Lymphocyte Activation. J Immunol 174, 2054–2060 (2005). 160. Francis, J. N., Sabroe, I., Lloyd, C. M., Durham, S. R. & Till, S. J. Elevated CCR6+ CD4+ T lymphocytes in tissue compared with blood and induction of CCL20 during the asthmatic late response. Clin Exp Immunol 152, 440–447 (2008). 161. Cook, D. N. et al. CCR6 Mediates Dendritic Cell Localization, Lymphocyte Homeostasis, and Immune Responses in Mucosal Tissue. Immunity 12, 495–503 (2000). 162. Singh, S. P., Zhang, H. H., Foley, J. F., Hedrick, M. N. & Farber, J. M. Human T cells that are able to produce IL-17 express the chemokine receptor CCR6. J Immunol 180, 214– 21 (2008). 163. Annunziato, F. et al. Phenotypic and Functional Features of Human Th17 Cells. J Exp Med 204, 1849–1861 (2007). 164. Pène, J. et al. Chronically Inflamed Human Tissues Are Infiltrated by Highly Differentiated Th17 Lymphocytes. J Immunol 180, 7423–7430 (2008). 165. Goleva, E. et al. Corticosteroid-resistant asthma is associated with classical antimicrobial activation of airway macrophages. J. Allergy Clin. Immunol. 122, 550–559.e3 (2008). 166. Yang, D. et al. Many Chemokines Including CCL20/MIP-3α Display Antimicrobial Activity. J Leukoc Biol 74, 448–455 (2003). 167. Chan, D. I., Hunter, H. N., Tack, B. F. & Vogel, H. J. Human Macrophage Inflammatory Protein 3 α: Protein and Peptide Nuclear Magnetic Resonance Solution Structures,
51 Dimerization, Dynamics, and Anti-Infective Properties. Antimicrob. Agents Chemother. 52, 883–894 (2008). 168. Nguyen, L. T., Chan, D. I., Boszhard, L., Zaat, S. A. J. & Vogel, H. J. Structure-function studies of chemokine-derived carboxy-terminal antimicrobial peptides. Biochim. Biophys. Acta 1798, 1062–1072 (2010). 169. Schumacher, C., Clark-Lewis, I., Baggiolini, M. & Moser, B. High- and low-affinity binding of GRO alpha and neutrophil-activating peptide 2 to interleukin 8 receptors on human neutrophils. Proc Natl Acad Sci U S A 89, 10542–10546 (1992). 170. Moser, B., Clark-Lewis, I., Zwahlen, R. & Baggiolini, M. Neutrophil-activating properties of the melanoma growth-stimulatory activity. J. Exp. Med. 171, 1797–1802 (1990). 171. Kunkel, S. L., Standiford, T., Kasahara, K. & Strieter, R. M. Interleukin-8 (IL-8): the major neutrophil chemotactic factor in the lung. Exp. Lung Res. 17, 17–23 (1991). 172. Dufour, J. H. et al. IFN-Γ-Inducible Protein 10 (IP-10; CXCL10)-Deficient Mice Reveal a Role for IP-10 in Effector T Cell Generation and Trafficking. J Immunol 168, 3195–3204 (2002). 173. Booth, V., Keizer, D. W., Kamphuis, M. B., Clark-Lewis, I. & Sykes, B. D. The CXCR3 binding chemokine IP-10/CXCL10: structure and receptor interactions. Biochemistry 41, 10418–10425 (2002). 174. Broide, D. H. et al. Cytokines in symptomatic asthma airways. Journal of Allergy and Clinical Immunology 89, 958–967 (1992). 175. Al-Shami, A., Spolski, R., Kelly, J., Keane-Myers, A. & Leonard, W. J. A Role for TSLP in the Development of Inflammation in an Asthma Model. J Exp Med 202, 829–839 (2005). 176. Kondo, Y. et al. Administration of IL-33 Induces Airway Hyperresponsiveness and Goblet Cell Hyperplasia in the Lungs in the Absence of Adaptive Immune System. Int. Immunol. 20, 791–800 (2008). 177. Virchow, J. C. et al. T Cells and Cytokines in Bronchoalveolar Lavage Fluid After Segmental Allergen Provocation in Atopic Asthma. Am. J. Respir. Crit. Care Med. 151, 960–968 (1995). 178. Cromwell, O. et al. Expression and generation of interleukin-8, IL-6 and granulocyte- macrophage colony-stimulating factor by bronchial epithelial cells and enhancement by IL-1 beta and tumour necrosis factor-alpha. Immunology 77, 330–337 (1992). 179. Wang, J. et al. Transgenic Expression of Granulocyte-Macrophage Colony-Stimulating Factor Induces the Differentiation and Activation of a Novel Dendritic Cell Population in the Lung. Blood 95, 2337–2345 (2000). 180. Besnard, A.-G. et al. Inflammasome–IL-1–Th17 Response in Allergic Lung Inflammation. J Mol Cell Biol 4, 3–10 (2012). 181. Bettelli, E. et al. Reciprocal developmental pathways for the generation of pathogenic effector TH17 and regulatory T cells. Nature 441, 235–238 (2006). 182. Tyner, J. W. et al. Blocking airway mucous cell metaplasia by inhibiting EGFR antiapoptosis and IL-13 transdifferentiation signals. J. Clin. Invest. 116, 309–321 (2006). 183. Oliphant, C. J., Barlow, J. L. & McKenzie, A. N. J. Insights into the initiation of type 2 immune responses. Immunology 134, 378–385 (2011). 184. Kurowska-Stolarska, M. et al. IL-33 Amplifies the Polarization of Alternatively Activated Macrophages That Contribute to Airway Inflammation. J Immunol 183, 6469– 6477 (2009).
52 185. Sullivan, D. E., Ferris, M., Pociask, D. & Brody, A. R. Tumor Necrosis Factor-Α Induces Transforming Growth Factor-Β1 Expression in Lung Fibroblasts Through the Extracellular Signal–Regulated Kinase Pathway. Am. J. Respir. Cell Mol. Biol. 32, 342–349 (2005). 186. Scheurich, P., Thoma, B., Ucer, U. & Pfizenmaier, K. Immunoregulatory Activity of Recombinant Human Tumor Necrosis Factor (TNF)-Alpha: Induction of TNF Receptors on Human T Cells and TNF-Alpha-Mediated Enhancement of T Cell Responses. J Immunol 138, 1786–1790 (1987). 187. Lukacs, N. W., Strieter, R. M., Chensue, S. W., Widmer, M. & Kunkel, S. L. TNF-Alpha Mediates Recruitment of Neutrophils and Eosinophils During Airway Inflammation. J Immunol 154, 5411–5417 (1995). 188. Liu, Y.-J. Thymic Stromal Lymphopoietin: Master Switch for Allergic Inflammation. J Exp Med 203, 269–273 (2006). 189. Al-Shami, A. et al. A Role for Thymic Stromal Lymphopoietin in CD4+ T Cell Development. J Exp Med 200, 159–168 (2004). 190. Alberts, B. et al. Molecular Biology of the Cell, Fourth Edition . (Garland Science, 2002). 191. Nilsson, C., K �gedal, K., Johansson, U. & �llinger, K. Analysis of cytosolic and lysosomal pH in apoptotic cells by flow cytometry. Methods in Cell Science 25, 185–194 (2004). 192. Arai, K., Shimaya, A., Hiratani, N. & Ohkuma, S. Purification and Characterization of Lysosomal H(+)-ATPase. An Anion-Sensitive V-Type H(+)-ATPase from Rat Liver Lysosomes. J. Biol. Chem. 268, 5649–5660 (1993). 193. Dröse, S. & Altendorf, K. Bafilomycins and Concanamycins as Inhibitors of V-ATPases and P-ATPases. J Exp Biol 200, 1–8 (1997). 194. Jentsch, T. J. Chloride and the Endosomal–Lysosomal Pathway: Emerging Roles of CLC Chloride Transporters. J Physiol 578, 633–640 (2007). 195. Graves, A. R., Curran, P. K., Smith, C. L. & Mindell, J. A. The Cl-/H+ antiporter ClC-7 is the primary chloride permeation pathway in lysosomes. Nature 453, 788 (2008). 196. Smith, A. J. & Schwappach, B. Think Vesicular Chloride. Science 328, 1364–1365 (2010). 197. Page, L. J., Darmon, A. J., Uellner, R. & Griffiths, G. M. L is for lytic granules: lysosomes that kill. Biochimica et Biophysica Acta (BBA) - Molecular Cell Research 1401, 146–156 (1998). 198. McDonald, J. K., Reilly, T. J., Zeitman, B. B. & Ellis, S. Cathepsin C: a chloride- requiring enzyme. Biochem. Biophys. Res. Commun. 24, 771–775 (1966). 199. Cigi ć, B., Pain, R. H., Cigi ć, B. & Pain, R. H. Location of the binding site for chloride ion activation of cathepsin C, Location of the binding site for chloride ion activation of cathepsin C. European Journal of Biochemistry, European Journal of Biochemistry 264, 264, 944, 944–951, 951 (1999). 200. Peters, P. J. et al. Cytotoxic T Lymphocyte Granules Are Secretory Lysosomes, Containing Both Perforin and Granzymes. J Exp Med 173, 1099–1109 (1991). 201. MacDermott, R. P. et al. Proteoglycans in Cell-Mediated Cytotoxicity. Identification, Localization, and Exocytosis of a Chondroitin Sulfate Proteoglycan from Human Cloned Natural Killer Cells During Target Cell Lysis. J Exp Med 162, 1771–1787 (1985). 202. Schmidt, R. E. et al. Specific release of proteoglycans from human natural killer cells during target lysis. , Published online: 21 November 1985; | doi:10.1038/318289a0 318, 289–291 (1985).
53 203. Stevens, R. L., Fox, C. C., Lichtenstein, L. M. & Austen, K. F. Identification of chondroitin sulfate E proteoglycans and heparin proteoglycans in the secretory granules of human lung mast cells. Proc Natl Acad Sci U S A 85, 2284–2287 (1988). 204. Masson, D., Peters, P. J., Geuze, H. J., Borst, J. & Tschopp, J. Interaction of chondroitin sulfate with perforin and granzymes of cytolytic T-cells is dependent on pH. Biochemistry 29, 11229–11235 (1990). 205. Humphries, D. E. et al. Heparin is essential for the storage of specific granule proteases in mast cells. Nature 400, 769 (1999). 206. Serafin, W. E., Katz, H. R., Austen, K. F. & Stevens, R. L. Complexes of Heparin Proteoglycans, Chondroitin Sulfate E Proteoglycans, and [3H]diisopropyl Fluorophosphate- Binding Proteins Are Exocytosed from Activated Mouse Bone Marrow-Derived Mast Cells. J. Biol. Chem. 261, 15017–15021 (1986). 207. Metkar, S. S. et al. Cytotoxic Cell Granule-Mediated Apoptosis: Perforin Delivers Granzyme B-Serglycin Complexes into Target Cells without Plasma Membrane Pore Formation. Immunity 16, 417–428 (2002). 208. McKechnie, N. M., King, B. C. R., Fletcher, E. & Braun, G. Fas-ligand is stored in secretory lysosomes of ocular barrier epithelia and released with microvesicles. Experimental Eye Research 83, 304–314 (2006). 209. Orlow, S. J. Melanosomes Are Specialized Members of the Lysosomal Lineage of Organelles. Journal of Investigative Dermatology 105, 3–7 (1995). 210. Baron, R., Neff, L., Louvard, D. & Courtoy, P. J. Cell-Mediated Extracellular Acidification and Bone Resorption: Evidence for a Low pH in Resorbing Lacunae and Localization of a 100-kD Lysosomal Membrane Protein at the Osteoclast Ruffled Border. J Cell Biol 101, 2210–2222 (1985). 211. Prussin, C. & Metcalfe, D. D. 4. IgE, mast cells, basophils, and eosinophils. Journal of Allergy and Clinical Immunology 111, 486–494 (2003). 212. Wright, D. G. & Gallin, J. I. Secretory Responses of Human Neutrophils: Exocytosis of Specific (Secondary) Granules by Human Neutrophils During Adherence in Vitro and During Exudation in Vivo. J Immunol 123, 285–294 (1979). 213. Brumell, J. H. et al. Subcellular Distribution of Docking/Fusion Proteins in Neutrophils, Secretory Cells with Multiple Exocytic Compartments. J Immunol 155, 5750–5759 (1995). 214. Utgaard, J. O., Jahnsen, F. L., Bakka, A., Brandtzaeg, P. & Haraldsen, G. Rapid Secretion of Prestored Interleukin 8 from Weibel-Palade Bodies of Microvascular Endothelial Cells. J Exp Med 188, 1751–1756 (1998). 215. Lortat-Jacob, H., Grosdidier, A. & Imberty, A. Structural Diversity of Heparan Sulfate Binding Domains in Chemokines. PNAS 99, 1229–1234 (2002). 216. Saunders, S., Jalkanen, M., O’Farrell, S. & Bernfield, M. Molecular Cloning of Syndecan, an Integral Membrane Proteoglycan. J Cell Biol 108, 1547–1556 (1989). 217. Roberts, R. et al. Heparan sulphate bound growth factors: a mechanism for stromal cell mediated haemopoiesis. Nature 332, 376–378 (1988). 218. Burgess, W. H. & Maciag, T. The heparin-binding (fibroblast) growth factor family of proteins. Annu. Rev. Biochem. 58, 575–606 (1989). 219. Gitay-Goren, H., Soker, S., Vlodavsky, I. & Neufeld, G. The Binding of Vascular Endothelial Growth Factor to Its Receptors Is Dependent on Cell Surface-Associated Heparin-Like Molecules. J. Biol. Chem. 267, 6093–6098 (1992).
54 220. Lortat-Jacob, H. & Grimaud, J. A. Binding of interferon-gamma to heparan sulfate is restricted to the heparin-like domains and involves carboxylic--but not N-sulfated--groups. Biochim. Biophys. Acta 1117, 126–130 (1992). 221. Ramsden, L. & Rider, C. C. Selective and differential binding of interleukin (IL)-1 alpha, IL-1 beta, IL-2 and IL-6 to glycosaminoglycans. Eur. J. Immunol. 22, 3027–3031 (1992). 222. Webb, L. M., Ehrengruber, M. U., Clark-Lewis, I., Baggiolini, M. & Rot, A. Binding to Heparan Sulfate or Heparin Enhances Neutrophil Responses to Interleukin 8. PNAS 90, 7158–7162 (1993). 223. Lyon, M., Deakin, J. A., Mizuno, K., Nakamura, T. & Gallagher, J. T. Interaction of Hepatocyte Growth Factor with Heparan Sulfate. Elucidation of the Major Heparan Sulfate Structural Determinants. J. Biol. Chem. 269, 11216–11223 (1994). 224. Witt, D. P. & Lander, A. D. Differential binding of chemokines to glycosaminoglycan subpopulations. Curr. Biol. 4, 394–400 (1994). 225. Aviezer, D. & Yayon, A. Heparin-Dependent Binding and Autophosphorylation of Epidermal Growth Factor (EGF) Receptor by Heparin-Binding EGF-Like Growth Factor but Not by EGF. PNAS 91, 12173–12177 (1994). 226. Clarke, D., Katoh, O., Gibbs, R. V., Griffiths, S. D. & Gordon, M. Y. Interaction of interleukin 7 (IL-7) with glycosaminoglycans and its biological relevance. Cytokine 7, 325– 330 (1995). 227. Lortat-Jacob, H., Garrone, P., Banchereau, J. & Grimaud, J. A. Human interleukin 4 is a glycosaminoglycan-binding protein. Cytokine 9, 101–105 (1997). 228. Li, Q., Park, P. W., Wilson, C. L. & Parks, W. C. Matrilysin Shedding of Syndecan-1 Regulates Chemokine Mobilization and Transepithelial Efflux of Neutrophils in Acute Lung Injury. Cell 111, 635–646 (2002). 229. Götte, M. & Echtermeyer, F. Syndecan-1 as a Regulator of Chemokine Function. TheScientificWorldJOURNAL 3, 1327–1331 (2003). 230. Götte, M. Syndecans in Inflammation. FASEB J 17, 575–591 (2003). 231. Wang, Z., Götte, M., Bernfield, M. & Reizes, O. Constitutive- and accelerated shedding of murine syndecan-1 is mediated by cleavage of its core protein at a specific juxtamembrane site. Biochemistry 44, 12355–12361 (2005). 232. Shafat, I., Vlodavsky, I. & Ilan, N. Characterization of Mechanisms Involved in Secretion of Active Heparanase. J. Biol. Chem. 281, 23804–23811 (2006). 233. Zuberi, R. I. et al. Deficiency of Endothelial Heparan Sulfates Attenuates Allergic Airway Inflammation. J Immunol 183, 3971–3979 (2009).
55 Chapter 2. Mechanism of CCL20 rapid release from allergen exposed epithelial cells
Mark Webb 1, Naina Gour 1, Stacey Burgess 1, Stephane Lajoie 1,2 , Ian Lewkowich 1, Marsha Wills-
Karp 1,2
1 Division of Allergy and Immunology, Cincinnati Children’s Hospital Medical Center, 3333
Burnet Ave, Cincinnati, OH, 45229
2 Department of Environmental Health Sciences, Johns Hopkins School of Public Health, Baltimore, MD,
USA, 21205
Prepared for publication in the Journal of Biological Chemistry
56 Abstract
Rapid chemokine release by the bronchial epithelium is an important initial step in the development of an adaptive response against allergen. Previous studies demonstrated CCL20 release within 30 minutes of HDM exposure. We asked whether this rapid release of CCL20 is from pre-formed stores. We found that after allergen exposure, CCL20 is quickly secreted via degranulation from preformed intracellular lysosomal stores. Targeting the lysosome-specific
V-ATPase and CLC-7 ion channels decreased HDM-induced CCL20 secretion. Lysosomal
CCL20 release was dependent on signaling through syk and raf1 kinases. Our findings reveal a novel pathway by which the bronchial epithelium releases pre-formed intracellular chemokine stores.
57 Introduction
Allergic asthma is a chronic inflammatory disease of the airways whose prevalence continues to rise in developed nations. Although the origins of allergic asthma are complex, excessive activation of T helper type 2 (T H2) cells specific for normally innocuous environmental allergens drives disease pathology.1 However, the initiating events following allergen exposure leading to the development of a T H2 response remain to be elucidated.
After exposure to allergen, the development of a T H2 response is dependent on the recruitment of immature dendritic cells (iDCs) to the lungs. This recruitment is primarily mediated by the chemokine CCL20,2 a strong chemoattractant for iDCs. CCL20 binds uniquely
to CCR6, which is strongly expressed by iDCs, and is vital for their recruitment into the lungs
following allergen exposure.3,4 The epithelium appears to be the major source of CCL20 in the airways,5 and we and others have shown that its secretion is strongly induced by allergen.6
Interestingly, chemokine release from cells can happen within a few minutes after contact with pattern recognition receptors or other agonists,7 suggesting they are contained in pre-formed
stores. Rapid secretion of cytokines from preformed stores has typically been studied in
granulocytes – in which the mechanisms governing release have been examined extensively –
and this release often involves secretory lysosomes.8,9 Rapid release of chemokines from
structural cells has also been reported,10 but the mechanisms driving this release – including whether they are released from extracellular stores, intracellular stores, or both – remain to be explored.
58 Dendritic cell recruitment to the lungs has been shown to occur within a few hours after allergen exposure in both mice and humans. 11 As iDCs are recruited by the chemoattractant
CCL20, this suggests that CCL20 is released within this time period. We thus hypothesized that
bronchial epithelial cells contain preformed stores of CCL20 for quick release in response to
HDM exposure and this release leads to the initial DC recruitment. We demonstrate that pools of CCL20 are stored within lysosomes, and these pools rapidly degranulate upon exposure to
HDM. These data point to a previously unknown mechanism of chemokine/cytokine release by
AECs from secretory lysosomes, and suggest that the epithelium may be a source of pre-formed cytokines designed for rapid release. This newly identified mechanism could contribute to the initiating events that lead to allergic inflammation.
59 Materials and Methods
Bronchial Epithelial Cells: 16HBE14o- cells were seeded at 12,500 cells/cm2 in 24-well collagen-coated culture plates in DMEM including 10% FBS. At 75% confluence, cells were serum starved in DMEM plus 0.1% FBS overnight prior to cell stimulation. All inhibitors were added one hour prior to HDM addition. HDM was added to cell supernatants at a concentration of 100 ug/ml.
Inhibitors: Inhibitors were titrated to determine maximal efficacy without affecting cell viability. Concentrations used are: bafilomycin A (Sigma, 0.5 pM), piceatannol (4 uM), Bay 61-
3606 (Sigma, 2 uM), Raf 1 kinase inhibitor I (EMD Millipore, 10 uM), R0 31-8220 (Cayman
Chemical, 10 uM).
Separation of cellular components: Supernatants were removed directly from cells after
treatment and stored at -20°C to -80°C. Cell lysates were obtained by rinsing treated cells in PBS
followed by 1-2 rapid freeze-thaw cycles. RNA was obtained using TRIzol reagent (Invitrogen)
and isolated according to manufacturer’s protocol. First-strand cDNA was synthesized using
the Superscript II First-Strand Synthesis kit (Invitrogen).
ELISA: Human CCL20 (DuoSet, R&D Systems) and human IL-8 (OptEIA, BD) ELISAs
were performed according to their respective protocol specifications.
Degranulation: Alexa-fluor647-conjugated Lamp-1 antibody (eBioscience, clone#
eBioH4A3) was added to cell culture concurrently with allergen. After 30 minutes, cells were
60 rinsed with PBS, and removed from culture plate with trypsin. Antibody uptake was then analyzed by flow cytometry.
Tissue and cell slide preparations: Mouse lungs were fixed in formaldehyde, and paraffin embedded. 2 uM sections were cut, then deparaffinized in xylene, and rehydrated in a graded ethanol series followed by water. Antigen retrieval was done in citrate buffer (10 mM sodium citrate, 0.05% Tween-20) at 90°C for 20 min. Human 16HBE14o- cells were grown and treated in
8-well chamber slides. After treatment, slides were fixed in 4% paraformaldehyde (Electron
Microscopy Sciences) for 15 minutes and rinsed with PBS.
Immunofluorescence staining: Slides were permeabilized with 0.1% Triton-X 100, and
blocked in 10% species serum matching the host of the secondary antibody. Slides were incubated overnight at 4C in primary mouse CCL20 antibody (Abcam, ab9829) or human
CCL20 antibody (R&D Systems, AF360). Fluorescently labeled secondary antibodies
(Invitrogen) were incubated for one hour at RT; slides were then rinsed with PBS-Tween 0.1% and mounted with VectaShield (Vector Labs).
Live immunofluorescence: To visualize CCL20, we generated a CCL20-mCherry fusion protein by inserting the human CCL20 open reading frame (minus the endogenous stop codon) into a mammalian expression plasmid (BD Biosciences), upstream of mCherry. Lamp1-YFP was obtained from AddGene.12 Rab3a-GFP, and Rab7-GFP were obtained from Origene. 16HBE14o-
cells were grown in Fluorodish (WPI) confocal dishes, and plasmids were transfected with
FuGENE HD (Roche) overnight. Cells were then serum-starved (0.1% FBS) for 24h, before
stimulation with HDM. LysoTracker (Invitrogen), ERTracker (Invitrogen), and NBD-
61 C6ceramide (Invitrogen) dyes were individually loaded into cells 30 minutes prior to imaging according to manufacturer protocol. Images were taken with a Zeiss LSM 710 confocal microscope.
Immunofluorescence quenching: Cells transfected with CCL20-mCherry were fixed, but not permeablized. Cells were then stained with anti-RFP polyclonal antibody (Invitrogen, R10367), or control antibody prior to immunofluorescence imaging.
62 Results
Kinetics suggests release of CCL20 is derived from preformed stores.
To better understand the mechanisms controlling CCL20 release, we first sought to assess the kinetics of CCL20 release after stimulation with HDM. We treated the human
bronchial epithelial cell line, 16HBE14o-, with HDM and measured CCL20 and IL-8 release, as well as mRNA at various times. We found that HDM induced significant CCL20 release into the supernatant in as little as five minutes (Fig. 1A), which continued to increase throughout the first hour. We saw little further increase in CCL20 release from two hours to six hours. This suggests that early release of CCL20 came from preformed stores, and that these stores were depleted during the first one to two hours after HDM exposure. In support of this hypothesis, the early burst of CCL20 protein was followed by an increase in CCL20 message 6h after HDM exposure (Fig. 1C), after which we saw a further increase of CCL20 in the supernatant from 6 to
24 hours after HDM exposure (Fig. 1A). Treatment with cycloheximide, which inhibits translation of new protein from mRNA, for one hour prior to HDM exposure had no effect on
CCL20 release after 2 hours, suggesting early CCL20 release is independent of de novo synthesis
(Fig. 1E).
In addition to HDM-induced release at 24 hours, significant accumulation of baseline release also appears to contribute to the overall level of CCL20 in the supernatants of these cells.
This suggests that low levels of CCL20 are released from bronchial epithelial cells at baseline, with an increase induced after HDM exposre. The difference between CCL20 released from
HDM-exposed cells and CCL20 released from control-treated cells appears to represent the
63 contribution of HDM to the CCL20 released from bronchial epithelial cells. Interestingly, this difference (HDM minus control) at 24 hours of HDM exposure represents a doubling of cumulative release over the difference of HDM minus control after 2 hours. This suggests that the first two hours of CCL20 release that can be attributed to HDM exposure represents half of the total release in the first 24 hours, and that HDM-induced release slows after the first two hours of HDM exposure.
In contrast to the kinetics seen for CCL20, significant increases of IL-8 protein release into the supernatant only began to appear after 2h of HDM treatment (Fig. 1B). This suggests
IL-8 may be released through a different mechanism than that governing CCL20. Furthermore,
IL8 expression levels did not appear to increase before 24h of HDM exposure (Fig. 1D). Given that IL-8 is released prior to this time, these data suggest that the increase in IL-8 release is derived from preformed stores of IL-8 protein, or IL-8 mRNA that is later transcribed. To determine whether transcription of IL-8 mRNA is required for IL-8 protein release in response
to HDM, we pre-treated cells with cycloheximide one hour prior to exposing them to HDM for
2 hours. Again in contrast to results seen with CCL20, IL-8 release was dramatically reduced in
the presence of cycloheximide, suggesting secretion of IL-8 requires translation of previously
formed mRNA. Also, the ratio of HDM-induced IL-8 release to control IL-8 release also
remained the same at 2 and 24 hours, suggesting the rate of release did not decrease over time,
as was observed in HDM-induced CCL20 release. Taken together, our findings indicate that
CCL20 and IL-8 are regulated by two different mechanisms.
64 CCL20 is stored in intracellular and extracellular pools.
To determine whether CCL20 is stored in distinct cellular pools, we transfected
16HBE14o- cells with CCL20 that was conjugated with the RFP-derived fluorophore mCherry
(CCL20-mCherry). The CCL20 gene was inserted at the N-terminus, with mCherry at the C- terminus, in order to ensure that any signal peptide remained intact. Epithelial cells transfected with a CCL20-mCherry fusion construct show both diffuse cellular staining (compatible with the hypothesis that stores of chemokine are retained on the surface of the cell) and punctate, granule-like stores (consistent with the hypothesis that stores of chemokines are retained in intracellular vesicles); these results support previous reports indicating that chemokines can be stored both in intracellular and extracellular locations.13,14 To test whether intracellular CCL20-
mCherry fluorescence could be distinguished from extracellular fluorescence, cells were left not
permeabilized and incubated with an anti-RFP antibody, which has been shown to quench RFP
fluorescence upon binding.15 We observed that the diffuse staining seen in the control panel was
lost in cells treated with the quenching antibody (Fig. 2A), suggesting that fluorescence from
cell surface stores of CCL20-mCherry were quenched by the antibody, whereas the punctate
CCL20 stores were unaffected by the quenching antibody, as we expect that these intracellular
stores are inaccessible to the quenching antibody.
We next sought to distinguish the cellular pools of CCL20 that are specifically released
in response to HDM, and whether they are derived from intracellular or extracellular sources.
To do this, we removed surface CCL20 in order to isolate the effects of HDM on intracellular
CCL20 only. We digested surface CCL20 by treating cells (that had already been exposed to
PBS or HDM for two hours) with trypsin until cell adherence was lost. As a control for the
65 effects of a sudden loss of adherence, cells were also dissociated using a non-enzymatic method.
The trypsin was then neutralized with 10% FBS (which was also applied to dissociation buffer controls), and the concentration of CCL20 remaining in cell lysates was analyzed (Fig. 2B). As expected, we observed a reduction in preformed CCL20 in the lysates of HDM-treated cells, presumably because it is being released from the cells into the supernatant. Under control conditions, lysates from non-enzymatically lifted cells (CD) contained more CCL20 than lysates from trypsin-treated cells (trypsin), suggesting that significant stores of CCL20 are stored on the exterior of the cells, where they were exposed to the trypsin. Although this extracellular CCL20 may also contribute to HDM-induced release of CCL20 into the supernatant, our measure of intracellular CCL20 (the trypsinyzed cells) showed that remaining stores of CCL20 significantly decreased after HDM treatment, demonstrating that intracellular stores of CCL20 are an important source of HDM-induced release into the supernatant after allergen exposure (Fig.
2B).
To further demonstrate that the early burst of CCL20 is from ready-to-be-secreted stores and does not necessitate trafficking through the endoplasmic reticulum and the Golgi apparatus, we inhibited vesicular trafficking with brefeldin A (which inhibits ER trafficking) 16
or monensin (which inhibits golgi trafficking).17 These two molecules inhibit vesicular trafficking, from transcription and translation, through different mechanisms. We found that pretreatment of cells with these inhibitors did not prevent early (2h) CCL20 release after HDM exposure (Fig. 2C). In contrast, early IL-8 release was completely sensitive to brefeldin A for
both baseline release (control) and HDM induced release; whereas for monensin baseline
66 release remained unaffected and HDM-induced release was decreased by one third (Fig. 2D).
This suggests that the majority of IL-8 export is not routed through the golgi apparatus in a monensin-sensitive manner, but that it is processed through the ER. After 24 hours of HDM exposure, CCL20 was decreased by both brefeldin and monensin, suggesting de novo production and trafficking through the Golgi apparatus are necessary to replenish CCL20 stores that are released in response to the initial stimulus. Interestingly, brefeldin A again had a stronger impact on reducing both baseline chemokine release, as well as the ratio of HDM-induced release to baseline. This suggests that after 24 hours, substantial stores of CCL20 de novo
production are trafficked through similar pathways to those of IL-8. The IL-8 release
mechanism appeared to remain constant from 2 to 24 hours, affected solely by brefeldin A (Fig.
2E, F). A similar distribution of cellular CCL20 was observed in mouse lung tissues stained
with CCL20, suggesting this pattern applies in vivo , as well as in vitro (Fig. 2G).
We next sought to determine the specific intracellular organelle to which CCL20 is
targeted for storage in 16HBE14o- cells. To do so, we input the human CCL20 peptide sequence
into the SignalP 4.0 server hosted by the Center for Biological Sequence Analysis to identify
putative signal peptide cleavage sites in the CCL20 protein sequence. We found one putative
cleavage site at SEA-AS, between peptides 26 and 27. When we BLASTed the first 26 peptides,
MCCTKSLLLAALMSVLLLHLCGESEA, we found a number of proteins with a high degree of
sequence homology within their predicted signal peptide regions. One such protein that
contained a highly similar sequence to that of CCL20 within its own predicted signal peptide
was cathepsin C, a lysosomal enzyme (Table 1). This correlation suggested a similar cellular
67 trafficking of both CCL20 and cathepsin C to lysosomes. To test whether CCL20 is stored in lysosomes, we co-transfected cells with CCL20-mCherry and markers for various types of intracellular organelles, including traditionally classified secretory vesicles (Rab3a-GFP), late endosomes/lysosomes (Rab7-GFP), and lysosomes (Lamp-1-YFP). No overlay was observed
between CCL20-mCherry fluorescence and that of rab3a-GFP (Fig. 3A), suggesting that CCL20 does not traffic through secretory vesicles distinguished by the marker rab3a. Partial co- localization was observed with rab7-GFP, suggesting that CCL20 can traffic through the late endosomal compartment (Fig. 3B). However, CCL20 fluorescence was strongly correlated with lamp-1-YFP fluorescence, suggesting that the majority of intracellular CCL20 is contained within lysosomes (Fig. 3C). Our previous observation that at least half of the cellular CCL20 resides on the external cell membrane suggests that some external CCL20 should be expected.
However, this is likely to be distributed throughout the extracellular membrane, and may therefore be too diffuse to observe juxtaposed to more highly concentrated lysosomal pools of
CCL20. Given that these cells are not grown in a trans-well environment, they are not polarized and we would not expect them to specifically localize the extracellular CCL20 to the baso-lateral surface. Accordingly, during imaging, no such polarized surface concentration of CCL20 was observed.
To test whether lysosomal CCL20 is released in response to HDM, cells were transfected with CCL20-mCherry and stained with LysoTracker dye, which inserts into acidic vesicles, such as lysosomes. In un-stimulated cells, we observed that most CCL20-mCherry was localized to
68 LysoTracker-containing acidic organelles; but after HDM treatment, lysosomal CCL20 was dramatically decreased (Fig. 4A).
We next wanted to examine whether epithelial cells undergo degranulation after HDM treatment. During degranulation, lamp-1 (which is normally found on the lysosomal membrane) relocates to the cell surface and levels of surface lamp-1 can be measured by flow cytometry. We observed that HDM induced a significant increase in degranulation, as measured by lamp-1 uptake, compared to control (Fig. 4B) treatment.
To test whether functioning lysosomes are required for CCL20 release, we treated cells with bafilomycin A, which prevents formation of lysosomes by blocking their acidification via the V-type H +-ATPase (V-ATPase).18,19 We observed that cells treated with bafilomycin displayed reduced CCL20 release in response to HDM (Fig. 4C); suggesting functional lysosomes are necessary for CCL20 release. These data suggest CCL20 is stored and released from secretory lysosomes. V-ATPase has been shown to localize to the cell surface, as well as to lysosomes. To distinguish between the roles of lysosomal and cell surface V-ATPases in contributing to CCL20 release, we specifically blocked extracellular V-ATPase using concanamycin A, which does not penetrate the cell membrane and cannot inhibit lysosomal V-
ATPases. Concanamycin had no effect on HDM-induced CCL20 (Fig. 4D), demonstrating that extracellular V-ATPase does not contribute to CCL20 release. To further confirm whether intracellular stores of CCL20 are sensitive to lysosome inactivation, we looked at CCL20 retention in cells treated with the lysosome inhibitor bafilomycin A. To to this, we treated cells with bafilomycin A prior to HDM treatment; we then trypsinized cells (to remove extracellular
69 CCL20 stores) and measured CCL20 in cell lysates. We observed that in bafilomycin A-treated cells intracellular CCL20 was retained after HDM exposure, suggesting inhibition of lysosomes with bafilomycin A specifically inhibited release of intracellular (lysosomal) CCL20. In contrast, cells containing both intracellular and extracellular stores of CCL20 displayed decreased levels of CCL20 retained in the cell lysates after HDM exposure (Fig. 4E), suggesting that this difference was derived from extracellular CCL20 stores.
In addition to V-ATPase, recent studies have shown that function of the chloride-proton cotransporter CLCN7 is also necessary for lysosome acidification. To further verify that lysosome function is necessary for CCL20 release, Clcn7 was knocked down via siRNA. We observed a reduction of CCL20 release after HDM exposure in Clcn7 knockdown cells compared to scrambled controls (Fig. 4F). These data further support our hypothesis that functional lysosomes are necessary for CCL20 release. Taken together, these data suggest that
CCL20 is stored in lysosomes within the cell, and that HDM induces degranulation of these lysosomes.
CCL20 release is mediated through beta glucan and syk/raf1 kinase signaling.
Previously we reported that beta-glucan signaling is necessary for HDM-induced CCL20 release after 24 hours. To test whether beta-glucans are necessary for CCL20 release at the 2 hour time point we have explored in this study, we digested HDM with beta-glucanase prior to treatment of 16HBE14o- cells and added either the digested HDM or the untreated HDM with vehicle control to the cells for 2 or 24 hours. We observed reduced CCL20 release in cells treated with HDM where beta-glucans had been enzymatically removed in comparison to cells treated
70 with HDM plus vehicle control. This observation was consistent after both 2 hour (Fig. 5A) and
24 hour (Fig. 5B) exposures. This finding suggests that HDM-induced CCL20 release after 2 hours is mediated by a similar beta-glucan dependent mechanism to what has been explored previously. 20
As shown in this previous study, signaling through spleen tyrosine kinase (syk) is associated with CCL20 release, which has been shown by others to lead to airway inflammation in asthmatics.21 In addition to syk, other pathways that have been shown to be associated with
beta-glucan signaling include raf 1 and protein kinase C (PKC). To further determine which intracellular signaling pathways contribute to CCL20 release, we treated cells with inhibitors to spleen tyrosine kinase (Bay 61-3606 and piceatannol), raf 1 kinase (raf 1 kinase inhibitor I), and
PKC (R0 31-8220). We found that inhibition of syk or raf 1 kinase reduced CCL20 release, but that PKC had no effect on CCL20 release. For both syk and raf 1 kinase inhibitors this effect more pronounced on baseline release, with some reduction of HDM-induced release after piceatannol exposure.
We next sought to visualize the effect of inhibiting syk/raf 1 kinase signaling on lysosomal stores. Therefore, we transfected cells with CCL20-mCherry and loaded them with
LysoTracker dye. Cells were incubated with the inhibitors listed above for 1 hour, and HDM was then added for 2 hours prior to imaging. We observed a dramatic reduction in colocalization of CCL20-mCherry with LysoTracker dye in the presence of syk inhibitors (Fig.
4D), suggesting that syk signaling is important for maintaining CCL20 within lysosomes under homeostatic conditions. Additionally, while cells treated in the presence of raf 1 kinase inhibitor
71 showed no change in colocalization of CCL20-mCherry and LysoTracker, these cells also failed to mobilize their lysosomal stores in response to HDM, suggesting that raf 1 signaling is a necessary component for signaling the release of CCL20 from secretory lysosomes in response to HDM exposure. Thus, our data suggest that CCL20 release from secretory lysosomes requires both syk and raf 1 kinase signaling, but that these pathways mediate separate cellular events in regards to CCL20 storage and release. Another potential interpretation of these results is that syk inhibition reduces shedding of extracellular CCL20. Thus, as intracellular stores of
CCL20 are released, some release may be captured by the extracellular stores that remain intact.
This would account for the reduction in baseline release of CCL20 observed in figure 5C.
72 Discussion
The rapid recruitment of iDCs to the lungs after allergen exposure is a critical initiating event in the development of the allergic response. The necessary signals for this recruitment involve key epithelial-derived mediators like CCL20 and GM-CSF.
We found that preformed stores of CCL20 in bronchial epithelial cells were released within a short (2 hr.) period after exposure to HDM. We also found that early CCL20 release from bronchial epithelial cells in response to HDM is derived from intracellular stores within secretory lysosomes. Lysosomal release required beta-glucan signaling in a syk/raf 1 kinase- dependent manner. Syk signaling was required to maintain CCL20 within lysosomes, while raf
1 kinase was required for release of lysosomal stores.
Interestingly, syk inhibition not only reduced release of CCL20 into the extracellular environment, but also reduced visible pools of pre-formed CCL20, even in the absence of HDM.
This suggests that syk signaling may be necessary to prevent CCL20 degradation, perhaps by maintaining a lysosomal structure that allows CCL20 to remain bound within the electron- dense core of the lysosome and thereby avoid degradation. Further biochemical analysis is required to determine the precise mechanism by which syk prevents loss of CCL20 within
bronchial epithelial cells. In other cell types, a direct role for syk in lysosome formation and
function has been suggested.22,23 Our findings support these previous observations, and suggest
that the loss of lysosomal structure in the absence of syk results in a loss of CCL20 stores within
the cell that is not associated with chemokine release.
73 In contrast to the immediate release seen with CCL20, IL-8 was not released until two hours of HDM exposure. At this time point, HDM-induced IL-8 release was completely dependent on protein synthesis and required trafficking through the Golgi apparatus, suggesting the cells increased de novo production of IL-8 in response to HDM exposure.
Interestingly, increased transcription of IL-8 was not detectable within the first 24 hours of
HDM exposure, long after de novo production of IL-8, which was observed within 2 hours of
HDM exposure. This suggests that prior to HDM exposure, although IL-8 mRNA is present in the cells, it likely undergoes translational repression. One major mechanism for translational repression that has been in recent years is through microRNAs, which bind to mRNA, and depending on the binding energy of the microRNA, either repress transcription or induce mRNA degradation.24 Future studies will determine whether a similar mechanism may control
IL-8 translation. However IL-8 translation is controlled, our data suggest that the early release of IL-8 we observed within the first 24 hours of HDM exposure, similar to early release of
CCL20, is not regulated by de novo transcription.
Previously, the mechanisms controlling chemokine release in bronchial epithelial cells have been poorly defined. This release is often thought to be controlled by direct secretory trafficking through the golgi. 25 As such, chemokine release is often measured after 12 or 24 hours, and increased mRNA is used as a measure of increased chemokine release. However, our studies suggest that by the time an increase in chemokine mRNA is observed, much initial
CCL20 and IL-8 release has already occurred, with de novo synthesis functioning to replenish already-released cellular stores, not to produce the initially released stores themselves. We
74 noted that stores of CCL20 begin to replenish about six hours after HDM exposure. This suggests that measurements of CCL20 release at later time points measure a delayed mechanism of release, which may also be mediated through secretory lysosomes. However, our studies also suggest that lack of message at a 24 hour time point does not suggest a lack of release, or even a lack of production, as message levels that were high at 6 hours decreased to
baseline by 24 hours.
Indeed, given that IL-8 release was similarly via an increase in mRNA within this time frame, we might hypothesize that a number of chemokines that display no significant increase in transcription within the first 24 hours could nonetheless be released within this time frame.
This also makes attribution of chemokine release to a specific cell type more difficult, as elevated levels of any specific chemokine in the BAL cannot necessarily be attributed to cells in which increased transcription of that chemokine are observed. Much of the observed rapidly released chemokines and chemokines released in the BAL of asthmatics in response to allergen could come from release of either preformed stores of these proteins (as is the case with CCL20), or from preformed mRNA that is subsequently translated (as we observed with IL-8).
In the present study, we focused primarily on CCL20 release from intracellular stores.
However, our evidence suggests the existence of extracellular stores of CCL20 whose release may be controlled by separate mechanisms from those controlling release of intracellular stores.
For example, when cell lysates were measured after trypsin degradation, some CCL20 was lost suggesting this pool of CCL20 that was exposed to trypsin resides outside the cell. We would not expect this pool of CCL20 to be affected by disruptions to lysosomal stores. Indeed, when
75 we treated cells with bafilomycin we saw partial inhibition of CCL20 release, suggesting non- lysosomal stores, such as those positioned on the cell surface, were unaffected by inhibition of lysosomes.
Kinetics of CCL20 release showed that low levels of CCL20 are released at baseline, in the absence of HDM stimulation. This constitutive, low-level release of CCL20 may play a role in the maintenance of resident dendritic cells in the lungs. Interestingly, baseline release of
CCL20 was unaffected by bafilomycin treatment, suggesting this low-level, constitutive release of CCL20 is not derived from lysosomal stores.
In contrast to bafilomycin A inhibition, when we inhibited production of CLC-7 via siRNA to CLCN7 , we observed that both baseline and HDM-induced CCL20 release were inhibited. Additionally, ratio of HDM to control CCL20 in knockdown cells was similar to the ratio in control-treated cells. This suggests that release of all CCL20 (including extracellular
CCL20) is impacted by CLCN7 knockdown, whereas for bafilomycin A knockdown only lysosomal stores were impacted. One major difference between these two approaches to lysosome inhibition is that bafilomycin A was applied to cells one hour prior to HDM exposure, while knockdown of cells began three days prior to HDM exposure. As we have previously observed that extracellular CCL20 is constitutively shed at a low level, this pool would need to
be replenished through de novo production. Given that long-term inhibition of lysosome function ( CLCN7 knockdown) impacted both intracellular and extracellular stores, this may suggest that CCL20 is trafficked through lysosomes in order to replenish extracellular stores.
76 When we measured the effect of signaling pathway inhibition, we again saw incomplete inhibition of CCL20 release after HDM treatment. However, we did not observe residual CCL20 stores within lysosomes as would be expected if lysosomal stores were the sole contributor to
HDM-induced CCL20 release, suggesting the CCL20 released in the presence of lysosome inhibition is derived from a separate, possibly extracellular, pool. In contrast to bafilomycin treatment, inhibitors of syk and raf1 reduced baseline CCL20 release, which suggests that the impact of these inhibitors on CCL20 release may not be restricted to lysosomal storage and release, but that these inhibitors may also interfere with constitutive release from extracellular stores discussed previously.
The finding that some preformed stores of CCL20 localize to lysosomes reveals novel insights about its function in response to antigen. CCL20 has been shown to be a potent antibacterial protein (with an LD 50 of 400 ng/mL against E. coli ) in addition to its signaling role.26
That CCL20 is not inactivated in the acidic lysosomal environment is unsurprising, given that
previous studies have shown CCL20 exhibits stronger antimicrobial properties at acidic pH,
which exposes its antimicrobial C-terminal domain.27 Thus, rapid release of preformed CCL20
stores from secretory lysosomes may contribute to a first-line defense against microbes at the
bronchial epithelium, with CCL20 release subsequently serving to help recruit DCs and T H17 cells. Many other chemokines exhibit similar antimicrobial properties, and 9some, such as IL-18, have been shown to be rapidly released from the bronchial epithelium in response to allergen exposure, similar to CCL20. 28 Future experiments will determine whether these two proteins are
stored and released in a similar manner. For CCL20, release from secretory lysosomes may
77 suggest that its antimicrobial role is the more ancient mechanism, as airway epithelial cells protect themselves by rapidly releasing antimicrobial CCL20, allowing subsequent adaptations to later link this release of antimicrobial peptides to inflammation.
In the context of HDM exposure, this mechanism may provide a useful target for intervention. Our finding that CCL20 release requires syk and raf 1 kinase signaling suggests that blocking these targets may affect early sensitization, and possibly airway inflammation in general.
78 References
1. Robinson DS, Hamid Q, Ying S, et al. Predominant TH2-like bronchoalveolar T-lymphocyte population in atopic asthma. N. Engl. J. Med. 1992;326(5):298–304. 2. Stick SM, Holt PG. The Airway Epithelium as Immune Modulator The LARC Ascending. Am. J. Respir. Cell Mol. Biol. 2003;28(6):641–644. 3. Lukacs NW, Prosser DM, Wiekowski M, Lira SA, Cook DN. Requirement for the Chemokine Receptor Ccr6 in Allergic Pulmonary Inflammation. J Exp Med . 2001;194(4):551–556. 4. Lundy SK, Lira SA, Smit JJ, et al. Attenuation of Allergen-Induced Responses in CCR6–/– Mice Is Dependent Upon Altered Pulmonary T Lymphocyte Activation. J Immunol . 2005;174(4):2054–2060. 5. Starner TD, Barker CK, Jia HP, Kang Y, McCray PB. CCL20 Is an Inducible Product of Human Airway Epithelia with Innate Immune Properties. Am. J. Respir. Cell Mol. Biol. 2003;29(5):627–633. 6. Reibman J, Hsu Y, Chen LC, Bleck B, Gordon T. Airway Epithelial Cells Release MIP- 3α/CCL20 in Response to Cytokines and Ambient Particulate Matter. Am. J. Respir. Cell Mol. Biol. 2003;28(6):648–654. 7. Tapper H. The Secretion of Preformed Granules by Macrophages and Neutrophils. J Leukoc Biol . 1996;59(5):613–622. 8. Persson-Dajotoy T, Andersson P, Bjartell A, Calafat J, Egesten A. Expression and Production of the CXC Chemokine Growth-Related Oncogene-Α by Human Eosinophils. J Immunol . 2003;170(10):5309–5316. 9. Ludwig JW, Richards ML. Targeting the secretory pathway for anti-inflammatory drug development. Curr Top Med Chem . 2006;6(2):165–178. 10. Øynebråten I, Barois N, Hagelsteen K, et al. Characterization of a Novel Chemokine- Containing Storage Granule in Endothelial Cells: Evidence for Preferential Exocytosis Mediated by Protein Kinase A and Diacylglycerol. J Immunol . 2005;175(8):5358–5369. 11. Jahnsen FL, Moloney ED, Hogan T, et al. Rapid Dendritic Cell Recruitment to the Bronchial Mucosa of Patients with Atopic Asthma in Response to Local Allergen Challenge. Thorax . 2001;56(11):823–826. 12. Sherer NM, Lehmann MJ, Jimenez-Soto LF, et al. Visualization of retroviral replication in living cells reveals budding into multivesicular bodies. Traffic . 2003;4(11):785–801. 13. Utgaard JO, Jahnsen FL, Bakka A, Brandtzaeg P, Haraldsen G. Rapid Secretion of Prestored Interleukin 8 from Weibel-Palade Bodies of Microvascular Endothelial Cells. J Exp Med . 1998;188(9):1751–1756. 14. Li Q, Park PW, Wilson CL, Parks WC. Matrilysin Shedding of Syndecan-1 Regulates Chemokine Mobilization and Transepithelial Efflux of Neutrophils in Acute Lung Injury. Cell . 2002;111(5):635–646. 15. Darmon A, Bar-Noy S, Ginsburg H, Cabantchik ZI. Oriented reconstitution of red cell membrane proteins and assessment of their transmembrane disposition by immunoquenching of fluorescence. Biochimica et Biophysica Acta (BBA) - Biomembranes . 1985;817(2):238–248.
79 16. Fujiwara T, Oda K, Yokota S, Takatsuki A, Ikehara Y. Brefeldin A causes disassembly of the Golgi complex and accumulation of secretory proteins in the endoplasmic reticulum. J. Biol. Chem. 1988;263(34):18545–18552. 17. Mollenhauer HH, Morré DJ, Rowe LD. Alteration of intracellular traffic by monensin; mechanism, specificity and relationship to toxicity. Biochim. Biophys. Acta . 1990;1031(2):225– 246. 18. Dröse S, Altendorf K. Bafilomycins and Concanamycins as Inhibitors of V-ATPases and P-ATPases. J Exp Biol . 1997;200(1):1–8. 19. Yoshimori T, Yamamoto A, Moriyama Y, Futai M, Tashiro Y. Bafilomycin A1, a specific inhibitor of vacuolar-type H(+)-ATPase, inhibits acidification and protein degradation in lysosomes of cultured cells. J. Biol. Chem. 1991;266(26):17707–17712. 20. Nathan AT, Peterson EA, Chakir J, Wills-Karp M. Innate immune responses of airway epithelium to house dust mite are mediated through beta-glucan-dependent pathways. J. Allergy Clin. Immunol. 2009;123(3):612–618. 21. Francis JN, Sabroe I, Lloyd CM, Durham SR, Till SJ. Elevated CCR6+ CD4+ T lymphocytes in tissue compared with blood and induction of CCL20 during the asthmatic late response. Clin Exp Immunol . 2008;152(3):440–447. 22. Majeed M, Caveggion E, Lowell CA, Berton G. Role of Src kinases and Syk in Fcγ receptor-mediated phagocytosis and phagosome-lysosome fusion. J Leukoc Biol . 2001;70(5):801–811. 23. He J, Tohyama Y, Yamamoto K, et al. Lysosome is a primary organelle in B cell receptor- mediated apoptosis: an indispensable role of Syk in lysosomal function. Genes to Cells . 2005;10(1):23–35. 24. Doench JG, Sharp PA. Specificity of microRNA target selection in translational repression. Genes Dev. 2004;18(5):504–511. 25. Stanley AC, Lacy P. Pathways for Cytokine Secretion. Physiology . 2010;25(4):218–229. 26. Yang D, Chen Q, Hoover DM, et al. Many Chemokines Including CCL20/MIP-3α Display Antimicrobial Activity. J Leukoc Biol . 2003;74(3):448–455. 27. Chan DI, Hunter HN, Tack BF, Vogel HJ. Human Macrophage Inflammatory Protein 3α: Protein and Peptide Nuclear Magnetic Resonance Solution Structures, Dimerization, Dynamics, and Anti-Infective Properties. Antimicrob. Agents Chemother. 2008;52(3):883–894. 28. Murai H, Qi H, Choudhury B, et al. Alternaria-Induced Release of IL-18 from Damaged Airway Epithelial Cells: An NF-κB Dependent Mechanism of Th2 Differentiation? PLoS ONE . 2012;7(2):e30280.
80 Downstream signaling inhibitors used in figure 2 Inhibitor Target Concentration SP600126 JNK (inhibitor) 10 uM R031 -8220 PKC (inhibitor) 10 uM Ly294002 PI3Kinase (inhibitor) 10 uM Raf 1 kinase inhibitor Raf1 kinase (inhibitor) 10 uM Bafilomycin V-ATPase (inhibitor) 0.5 uM Pertussus toxin Increases cAMP 100 ng/mL ZM 336372 Raf1 (activator) 10 uM U0126 MEK1/2 (inhibitor) 10 uM SB216763 GSK -3 (inhibitor) 10 uM CPG 57380 MNK1 (inhibitor) 10 uM Wortmannin PI3Kinase (inhibitor) 1 uM Pimelic diphenylamide 106 HDAC 1, 2, 3, 8 (inhibitor) 2 uM PP242 mTOR (inhibitor) 1 uM Rapamycin mTOR (inhibitor) 0.1 uM MS -275 HDAC 1>3 (inhibitor) 0.5 uM Trichostatin HDAC 1, 3, 4, 6, 10 (inhibitor) 0.1 uM SAHA HDAC (inhibitor) 1 uM EX -527 SIRT1 (inhibitor) 0.1 uM CAY10603 HDAC 6 (inhibitor) 0.1 uM IKK2 inhibitor NFkappaB (inhibitor) 1 uM GSIxxx Gamma secretase/Notch (inhibitor) 1, 10, 100 nM Table 1
81 Signal peptide homology for selected chemokines and cytokines Chemokine Signal peptide homologue CCL20 CR1L, cathepsin C, β4 integrin, CD46, urocortin IL -8 CX 3CL1 β-defensin 2 CD9 IFN -α – IFN -β Cadherin 7 IFN -γ – IL -10 – TNF -α – IL -17a – IL -6 – GM -CSF – TSLP – CCL22 Cortexin -1 CCL18 MIP -1α, LY6H, CYP7B1 CXCL10 No matches XCL1 FGFR4, NMDAR, PLOD1 Table 2
82 Actin inhibitors and their functions Inhibitor Function Cytochalasin D Binds G -actin and prevents incorporation into filaments Jasplakinolide Prevents F -actin extension and polymerization Latrunculin B Prevents G -actin polymerization Swinholide A Inhibits actin polymerization, and severs F -actin Table 3
83 Supernatant mRNA of de novo protein synthesis, 16HBE cells were treated with cycloheximide for 1 hour prior to *** A 800 PBS C 5 HDM exposure for 2 hours, and (E), CCL20 and (F), IL-8 were measured in supernatants by HDM *** 4 ELISA. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001 compared to
) 600 L m 3 control. pg / (
400 0
2 2
L ** CCL20 mRNA
C **
C 200 ** ** 1 ** 0 0 5 20 40 60 2 24 0.5 1 2 4 6 24 minutes hours 5 mins time (h) B 250 PBS *** D 5 HDM 200 4 * ) L
m 150 3 pg / ( 100 2 L- 8 I IL8 mRNA 50 1
0 0 5 20 40 60 2 24 0.5 1 2 4 6 24 minutes hours 5 mins time (h)
Supernatant F E 100 ** ns 150 *** PBS *** HDM 80 *** ) L
m 100 / 60 (p g
40 50 ns IL-8 (pg/mL) CCL2 0 20
0 0 Ctrl CHX Ctrl CHX
Figure 1 – Rapid CCL20 release from bronchial epithelial cells is dependent on preformed stores. 16HBE cells were treated with HDM, and cells were harvested at di erent time points for determination of (A) CCL20 and (B) IL-8 secretion by ELISA. RNA was extracted, and (C), Ccl20 mRNA and (D), Cxcl1 mRNA were measured by real-time PCR. To determine the involvement of de novo protein synthesis, 16HBE cells were treated with cycloheximide for 1 hour prior to
HDM exposure for 2 hours, and (E), CCL20 and (F), IL-8 were measured in supernatants by
ELISA. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001 compared to control. n=3. anti-RFP antibody, or isotype control antibody. (B) HDM-treated 16HBE14o- cells were lifted A B Lysate with either an enzymatic (trypsin) or non-enzymatic (cell dissociation bu er; CD) method, 200 * PBS ** HDM lysed by freeze-thaw, then CCL20 content in lysates was analyzed by ELISA. Cells were treated
Control 150 ** with HDM in the presence of brefeldin A, monensin, or vehicle control; supernatants were 100 0-mCherry I
2 collected at 2h (C, D) and 24h (E, F) and tested for CCL20 and IL-8 via ELISA. (G) Lung sections CL20 (pg/mL) -RF P C 50 from naïve BALB/c mice stained with anti-CCL20 antibody and DAPI nuclear stain. Data are DA P CC L ant i 0 represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001. CD trypsin
C D ns *** ns *** 100 PBS 60 PBS *** *** HDM HDM 80 40 60
40 20 CL20 (pg/mL) IL-8 (pg/mL) 20 G DAPI 0 0 CCL20 rol A rol A
Cont eldin Cont eldin ref Monensin ref Monensin B B
E F
*** *** 1000 250 *** *** PBS HDM 800 *** PBS 200 HDM 600 150
lumen 400 100 IL-8 (pg/mL) CL20 (pg/mL) 200 50
0 0 rol A rol A
Cont eldin Cont eldin ref Monensin ref Monensin B B
Figure 2 – CCL20 is stored in both intracellular and extracellular compartments. (A) Fixed, but not permeablized 16HBE14o- cells transfected with CCL20-mCherry, and exposed to either an anti-RFP antibody, or isotype control antibody. (B) HDM-treated 16HBE14o- cells were lifted with either an enzymatic (trypsin) or non-enzymatic (cell dissociation buf er; CD) method, lysed by freeze-thaw, then CCL20 content in lysates was analyzed by ELISA. Cells were treated with HDM in the presence of brefeldin A, monensin, or vehicle control; supernatants were collected at 2h (C, D) and 24h (E, F) and tested for CCL20 and IL-8 via ELISA. (G) Lung sections from naïve BALB/c mice stained with anti-CCL20 antibody and DAPI nuclear stain. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001. n=3. A B C
rab 3a-GFP rab 7-GFP lamp-1-YFP
CCL20-mCherry CCL20-mCherry CCL20-mCherry
Overlay Overlay Overlay secretory late lysosomes vesicles endosomes
Figure 3 – Intracellular CCL20 is stored within lysosomes. Live cell confocal microscopy images of cells co-transfected with CCL20-mCherry and either (A) rab 3a-GFP, (B) rab 7-GFP, or (C) lamp-1-YFP. A PBS HDM B 80 PBS C 50 (F) Cells were electroporated with scramble control or CLCN7-targeting siRNAs and treated HDM ** * 60 *** 40 PBS with media or HDM for 2h, and CCL20 release into the supernatant was measured. (G) mRNA HDM 30 40 expression levels of F. *p<0.05, **p<0.01, ***p<0.001. Scale bar = 10 uM. 20 CL20 (pg/mL) Lamp-1 MFI
25 C LysoTracker 10
0 0 Ctrl Baf A PBS HDM
D ns 100 * ns PBS E 200 PBS - CD HDM HDM - CD CCL20-mCherry 80 *** 150 PBS - Tryp * 60 HDM - Tryp ns 100 40
CCL20 (pg/mL) 50 20 CCL20 (pg/mL)
0 0 Ctrl BafA Overlay Control
Concanamycin
F 120 G 15 * PBS ** *** HDM
80 10 PBS HDM
40 Clcn7 mRNA 5 CCL20 (pg/mL)
0 0 Scbl Clcn7 Scbl Clcn7 siRNA siRNA siRNA
Figure 4 – HDM induces release of CCL20 from secretory lysosomes. (A) Cells transfected with CCL20-mCherry and loaded with LysoTracker were treated for 2 hours with PBS, or HDM. (B) Cells were incubated with Lamp-1—AF647 antibody and HDM for 30 minutes. Antibody uptake was then assessed via ow cytometry. (C, D) Cells were pretreated with ba lomycin A or concanamycin A for 1h, prior to adding media or HDM for 2h, and measuring CCL20 release into the supernatant. (E) Cells were treated with vehicle control or ba lomycin A for one hour prior to addition of HDM for 2 hours; CCL20 retention was measured as described in g. 2B. (F) Cells were electroporated with scramble control or CLCN7-targeting siRNAs and treated with media or HDM for 2h, and CCL20 release into the supernatant was measured. (G) mRNA expression levels of F. *p<0.05, **p<0.01, ***p<0.001. Scale bar = 10 uM. B-E n=3, F-G n=6. A B 80 2 hrs. 600 24 hrs. C 100 PBS *** ** *** HDM *** 80 *** 60
400 mL ) 60
40 (pg / ††† ††† 40 200 ††† 20 CCL20 CCL20 (pg/mL) CCL20 (pg/mL) 20 * ** 0 0 0
BS BS P P
HDM+veh HDM+veh HDM+b-gluc HDM+b-gluc Veh. ControlBay 61-3606Piceatannol Ro 31-8220 Raf 1 k inhibitor
Control Ba lomycin A Piceatannol Raf 1k inh. D
LysoTracker PBS
CCL20-mCherry
Overlay
LysoTracker HDM
CCL20-mCherry
Overlay Figure 5 – Beta-glucan-dependent release of lysosomal CCL20 is, through a syk/Raf 1 kinase- dependent pathway. A and B, Cells were treated with HDM that was pre-treated with either beta-glucanase (HDM+b-gluc) or vehicle control (HDM+veh) for 2 hours A or 24 hours B. C, Cells were treated with various signaling pathway inhibitors, including piceatannol (syk), raf 1 kinase inhibitor I (raf 1 kinase), and Ro 31-8220 (PKC) and CCL20 release into the supernatant was measured. D, Live confocal microscopy of cells transfected with CCL20-mCherry, loaded with LysoTracker dye, and treated with the inhibitors mentioned above, or baf lomycin A, then treated with PBS, or HDM for 2 hours. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001 vs. PBS control PBS; †p<0.05, ††p<0.01, †††p<0.001 vs. HDM control. Scale bar = 5 uM n=3. Chapter 3. Signaling pathways leading to HDM-induced CCL20 release from the bronchial epithelium
Mark Webb*, Naina Gour*, Stacey Burgess*, Stephane Lajoie**, Ian Lewkowich*, Marsha Wills-
Karp**
* Division of Allergy and Immunology, Cincinnati Children’s Hospital Medical Center, 3333
Burnet Ave, Cincinnati, OH, 45229
** Department of Environmental Health Sciences, Johns Hopkins School of Public Health, Baltimore,
MD, USA, 21205
93 Abstract
A number of intracellular signaling pathways have been identified as important in the generation of allergic asthma. General inhibition of many of these – such as reactive oxygen species generation, ion transport, and the NLRP3 inflammasome – have been shown to reduce airway hyperresponsiveness in animal models of asthma. We hypothesized that these same signaling pathways may contribute to HDM-induced CCL20 release from bronchial epithelial cells. We treated bronchial epithelial cells with inhibitors to a number of intracellular signaling pathways prior to treatment with HDM and measured CCL20 release. We found that CCL20 release does not require glucocorticoid signaling, as do many other chemokines and cytokines.
Additionally, we found that while chloride transport is necessary for HDM-induced CCL20 release, excess potassium was sufficient to reverse this effect. Other signaling contributors to
CCL20 release included gap junctions, reactive oxygen species, and cyclooxygenase. These findings demonstrate that CCL20 release is downstream of a number of pathways that are active in asthma, suggesting that inhibition of these pathways in asthmatics may have broader effects on inflammation and airway hyperresponsiveness.
94 Introduction
Chemokines and cytokines have been shown to be stored within a number of different cell types; including cells of the hematopoietic lineage, 1 as well as non-hematopoietic cells, such as endothelial cells. 2 These chemokines and cytokines mediate a diverse set of responses. For example, IL-8 and gro-α (found in endothelial cells) promote neutrophil infiltration, 3,4 while IL-4
(found in T H2 cells) promotes type-2 immune responses.5 The release of these different molecules is controlled by a diverse set of signaling pathways.
Different signaling pathways have been shown to play important roles in either the development or treatment of allergic asthma, such as signaling through the inflammasome, magnesium ion transport, reactive oxygen species (ROS) generation, and prostaglandin synthesis through cyclooxygenases. For example, activation of the NLRP3 inflammasome has
been shown to be critical to the development of T H2 and T H17 responses in murine asthma
models.6,7 Magnesium (Mg 2+ ) treatment may help reduce asthma symptoms by relaxing the
airway smooth muscle or by attenuating respiratory burst. 8,9 For this reason, magnesium is used as a treatment for patients suffering acute asthma attacks. 10 In severe asthmatics, increased
neutrophil inflammation results in release of reactive oxygen species and prostaglandins. 11–13
The bronchial epithelium has been shown to directly produce both reactive oxygen species, and prostaglandins.11,14,15 Each of these pathways has been shown to play a distinct role in inflammation.
One of the primary drivers of inflammation is the release of chemokines, which are critical for the recruitment of migrating cells. 16 We hypothesized that signaling pathways
95 already shown to actively contribute to airway hyperresponsiveness in asthmatics may also control release of CCL20 from bronchial epithelial cells. Many of these same mechanisms have already been shown to be active in bronchial epithelial cells following allergen exposure.
Others have been shown to contribute to airway inflammation, which suggests they may play a role in chemokine release from airway cells.
One source of inflammatory cytokines in allergic asthma is the inflammasome. 6,17
Activation of caspase 1 through NALP3 induces the release of IL-1β and IL-18. 18 This signaling
cascade can be initiated by a number of factors, including potassium export through pannexin-
1, external ATP, uric acid crystals, 19 and bacterial toxins such as nigericin (a potassium ionophore).20,21 Interestingly, potassium export has also been shown to be necessary for secretion of TNF-α, and IL-8. Inhibition of potassium export reduced immediate IL-8 and TNF- a release, but this effect began to diminish after 2 hours. 22
Signaling through other ions has also been shown to be involved in cytokine release.
For example, magnesium deficiency leads to early increases in IL-623 and TNF-α release.24
Indeed, intracellular magnesium levels have been shown to regulate lysosomal pH, 25 possibly through their effect on lysosomal chloride transport. 26 This may be one route by which
magnesium regulates release of intracellular chemokines and cytokines.
If individual ions contribute to signaling of CCL20 release, intercellular transport of
these ions may serve to enhance this signaling as changes in ion concentration spread from cell
to cell. One mechanism by which ions are transported between adjacent cells is by diffusing
through gap junctions. Gap junction proteins form gated pores between cells. 27 These pores are
96 large enough to allow transport of ions and small molecules, but not larger proteins. 28 In this way, signaling that is initiated in one cell may propagate to neighboring cells. For example, cells infected with limiting numbers of intracellular bacteria promoted NF-κB nuclear
translocation and IL-8 release in neighboring, uninfected cells through a mechanism dependent
on gap junction function. 29 A similar mechanism may help promote CCL20 release from
bronchial epithelial cells that are not directly contacted by allergen.
Another signaling pathway that has been shown to induce chemokine release in some cell types is induction of reactive oxygen species. ROS are highly reactive oxygen-containing molecules (such as H2O2) that help protect against foreign microorganisms, and participate in cell signaling. They are often generated in times of cell stress through a process referred to as respiratory burst.30,31 During respiratory burst, NADPH oxidase helps to convert oxygen to
H2O2.32 Although this pathway has traditionally been linked to metabolic processes within the
cell, recent studies have demonstrated that ROS participate in downstream cell signaling events
as well.33 This is accomplished through the oxidation, mainly of cysteine residues, of target proteins. A variety of different protein oxidation targets have been identified, including calcium and potassium ion channels, 34,35 MMPs and ADAMs, 36 JNK and ERK, 37–39 among others.
We have previously identified many of these signaling pathways as important in mediating
CCL20 release through either intracellular or extracellular mechanisms. Generation of reactive oxygen species has also been associated with release of the proinflammatory cytokines IL-1β and IL-18 in macrophages. 40 Given the many roles of ROS in mediating signaling pathways
97 previously identified as important in CCL20 release, as well as their influence in asthma, we hypothesized that ROS generation may play a role in HDM-induced CCL20 release.
Prostaglandins are lipophilic, paracrine signaling molecules produced from arachadonic acid. 41 Cyclooxygenase enzymes mediate the synthesis of prostaglandin H 2 from arachadonic acid. 42 From this point, other prostaglandins are synthesized. Three isoforms make up the
cyclooxygenase family: COX-1, COX-2, and COX-3. 43,44 Inhibition of COX-1 and COX-2 is a major function of aspirin, a common non-steroidal anti-inflammatory drug (NSAID). 45
Although prostaglandin release in asthmatic airways has been reported, 13 in some asthmatics
aspirin treatment exacerbates airway hyperresponsiveness (AHR), suggesting prostaglandin
synthesis may mitigate AHR in these individuals.46 Specificity of cyclooxygenase inhibition also seems to be important, as inhibitors of COX-2, but not COX-1 do not affect AHR, suggesting aspirin’s role in exacerbating AHR is due to its role in inhibiting COX-1, not COX-2. 47
Prostaglandin signaling has also been linked to cytokine secretion. Prostaglandin E2 (PGE2) in
T cells suppressed IFN-g, but increased production of IL-5. 48 In addition, inhibition of JNK or
ERK signaling reduced PGE2 production and IL-8 release. 49 Thus, the function of
prostaglandins in asthmatic airways is complex, with some prostaglandins promoting a T H2
response, and others antagonizing it.
Many of the pathways we have reviewed have been demonstrated to be important in
both airway hyperresponsiveness, and intracellular signaling events. We hypothesize that
some of these pathways may promote CCL20 release, which would stimulate inflammation and
98 AHR. To investigate which pathways are important for CCL20 release, we will block various pathways and analyze HDM-induced CCL20 release into the supernatant.
99 Materials and Methods
Cell culture and treatment: The 16HBE14o- human bronchial epithelial cell line was cultured in
10-15 cm round culture dishes that had been collagen coated. Cells were passaged by applying trypsin (0.05% with EDTA) for 10 min. Treatments were performed in 24-well collagen-coated culture plates in DMEM including 10% FBS, penicillin/streptomycin, and L-glutamine. Upon reaching 70-80% confluence, media was replaced and FBS reduced to 0.1% overnight. All inhibitors or balanced culture solutions were added one hour prior to HDM or allergen addition. Primary normal human bronchial epithelial cells (NHBEs) were thawed directly from storage in liquid nitrogen into 24-well collagen-coated culture plates. NHBEs were grown in
Bronchial Epithelial Cell Growth Medium (BEGM) from Lonza (CC-3170) with supplied supplements, including bovine pituitary extract (BPE), insulin, hydrocortisone (HC), GA-1000, retinoic acid, transferrin, triiodothyronine, epinephrine, and human epidermal growth factor
(hEGF). Upon reaching 70-80% confluence, media was replaced with BEGM deficient in BPE,
HC, and hEGF.
Inhibitors: Reagents used as inhibitors were obtained from SigmaAldrich, unless specified, and were used at the following concentrations: forskolin, gadolinium, carbenoxolone (50 uM), phloretin, 1-octanol, valinomycin (50 uM), SB203580 (10 uM), SP600125 (JNK inh.) (10 uM),
Bafilomycin (500 pM), Ly294002 (10 uM), R031-8220 (PKC inh.) (10 uM), Raf1 kinase inh. (10 uM), DIDS (10 uM), Pertussus toxin (100 ng/mL), ZM 336372 (10 uM), U0126 (10 uM), SB216763
(10 uM), CPG 57380 (10 uM), Wormannin (1 uM), PP242 (1 uM), Pimelic Diphenylamide 106 (2 uM), Dexamethasone (1 uM), Rapamycin (0.1 uM), MS-275 (0.5 uM), EX-527 (0.1 uM), SAHA (1
100 uM), CAY10603 (0.1 uM), IKK2 inh. (1 uM), Trichostatin (TSA) (0.1 uM), Monensin, brefeldin A, cytochalasin D (10 uM), Jasplakinolide (10 uM), Latrunculin (10 uM), Swinholide (10 uM).
HDM treatment: Whole HDM extract was obtained from Greer (B70). 100-200 ug/mL HDM was added to cell supernatant for two hours prior to harvesting cells.
Solutions : Chloride sufficient media is a Krebs-Ringer solution consisting of the following (in mM): 120 NaCl, 25 NaHCO 3, 11 Glucose, 4.7 KCl, 2.5 CaCl 2, 1.2 MgSO 4, and 1.2 KH 2PO 4. In anion replacement solutions, 100 mM of NaCl was replaced with 100 mM sodium gluconate.
For sulfate-free solutions, MgSO 4 was replaced with Na 2SO 4. For magnesium-free solutions,
MgSO 4 was replaced with MgCl 2. For potassium-free solutions, KCl was replaced with NaCl,
KH 2PO 4 and was replaced with NaH 2PO 4.
ELISA : Mouse and human ELISAs were performed in half-well plates using DuoSet ELISA kits
from R&D Systems according to their specifications. Multiplex quantification was performed
via Luminex using proprietary kits and 50 uL sample volume. Samples were from BAL or cell
supernatants that had been frozen at -80 oC.
Intracellular chloride measurement : Intracellular chloride measurements are modified from the
protocol reported in (Coskun T, Baumgartner HK, Chu S & Montrose MH (2002). Coordinated
regulation of gastric chloride secretion with both acid and alkali secretion. Am J Physiol
Gastrointest Liver Physiol 283, G1147–G1155 ). After HDM treatment for 30 min, cells were rapidly washed three times in ice cold 140 mM Sodium gluconate solution during continuous aspiration. Samples were allowed to dry before reconstitution in 2N sulfuric acid and 1%
101 Tween 20. Cells were subsequently diluted to 0.67 N sulfuric acid before chloride concentrations were measured using a Labconco Chloridometer, and compared with a standard curve of known chloride concentrations.
Mouse asthma model: Sensitization to HDM was achieved by intra tracheal (I.T.) administration of
100 ug/mL HDM. Twenty-one days later a second administration of alexafluor 405-labelled
HDM was administered I.T. 72-hours after the last HDM challenge mice were anesthetized with sodium pentathol and airway measurements (outlined below) were taken. The mice were then sacrificed, and broncho-alvelar lavage was performed using 1 mL Hanks balanced salt solution without calcium or magnesium. BAL fluid was then centrifuged to remove cells.
Lungs were removed and fixed in formaldehyde, then sent to core personnel at CCHMC who embedded them in paraffin, sectioned them, and mounted them on slides.
Airway measurements: 72 hours after the final allergen exposure, mice were anesthetized,
intubated, and ventilated at 120 breaths/min. After airway pressure stabilized, mice were given
25 ug/kg weight of acetylcholine intra venous and airway pressure changes were followed for
five minutes. Airway measurements were quantified as airway pressure time index (APTI),
which is a quantitative measure of airway constriction, such that APTI increases with increasing
airway resistance.
Isolation and culture of mouse tracheal epithelial cells (mTECs): Mice were anesthetized with sodium
pentathol and the inferior vena cava was severed to prevent recovery. Trachea were then
removed and incubated overnight at 4 OC in 1% pronase (Roche) in DMEM/F12 (50/50) mix
(Fisher). Cells were then removed from trachea via sheer force in the presence of 10% FBS and
102 DNase. Cells were then incubated on Primaria plates for four hours to allow fibroblasts to adhere. Non-adherent cells were then removed and grown on rat-tail collagen-coated trans- well plates in DMEM/F12 supplemented with penicillin/streptomycin, L-glutamine, retinoic acid, and 5% FBS. After reaching 1000 ohms trans-epithelial resistance, media was replaced with 0.1% FBS media overnight prior to experimental treatments.
Statistical analysis: Statistical analysis was performed using GraphPad Prism 5 software. As the data were normally distributed, statistical significance was determined using an unpaired student’s t-test for comparisons between two treatments, or two-way ANOVA combined with
Bonferroni post-test analysis for comparison between multiple groups. Values of p<0.05 were considered statistically significant.
103 Results
A chloride cotransporter is required for rapid, HDM-induced CCL20 release
We sought to identify the cellular signaling mechanisms that lead to the rapid release of intracellular stores of CCL20 after HDM stimulation. Previously, we showed that CCL20 is released from secretory lysosomes in a V-ATPase and chloride-proton co-transporter— dependent manner. We therefore hypothesized that the intracellular signaling pathways leading to this secretory lysosome release may also be ion transporter-dependent. To test this, we treated HBE cells with a variety of transporter inhibitors prior to HDM stimulation and assayed for early CCL20 release into the supernatant. While common cation transport inhibitors, such as amiloride (epithelial sodium channel) and bumetanide (NKCC1) had no effect on CCL20 release (data not shown), broad chloride transporter inhibitors such as DIDS and niflumic acid dramatically reduced CCL20 release (Fig 1A). This suggests a vital role for anion transport in CCL20 release, as no detectable CCL20 was released in cells where anion transport was inhibited.
To further confirm the role of chloride in CCL20 release, we depleted cells of chloride by treating them in a modified Ringer solution. We replaced 100 mM of sodium chloride with sodium gluconate, thereby creating a reduced chloride solution containing 25 mM chloride (Fig
1B). This solution was applied to the cells one hour before HDM stimulation. Cells under reduced chloride conditions released substantially less CCL20 than did cells treated under normal chloride conditions. Although we retained about 25 mM chloride in reduced chloride conditions (to minimize shock to the cells) we wanted to confirm that the cells were still viable
104 after the two-hour stimulation with HDM. Therefore, subsequent to collecting the supernatant from the cells in figure 1 B (2 hr), we restored normal chloride solution to these cells, while continuing to stimulate with HDM for an additional two hours. In cells that were initially chloride deficient, restoration of chloride was sufficient to fully restore CCL20 release in response to HDM. Interestingly, in cells that were initially chloride sufficient, and had therefore showed strong release of CCL20 after 2 hours, we noted that CCL20 release after the second stimulation was significantly reduced. As the initial burst of CCL20 from these cells had been removed prior to restoring chloride-sufficient solution plus HDM, this low secondary release represents HDM-induced CCL20 release from 2 to 4 hours after exposure. This low release from 2 to 4 hours is consistent with our previous observation that preformed stores of CCL20 are not replenished within the first few hours after HDM stimulation.
To further define the halogen anion selectivity of this CCL20 release mechanism, we replaced chloride in our modified Ringer solution with anions known to exhibit increased permeability through most chloride transporters: namely nitrate, iodide, and fluoride (data not shown). We found that when chloride was replaced with nitrate or iodide CCL20 release increased both at baseline and under HDM-stimulated conditions. (Fluoride-containing solution was highly cytotoxic.) This increase was also susceptible to suppression by DIDS (Fig
1C), strongly supporting our hypothesis that the effect of chloride on CCL20 release is through anion transporter activity. From this we concluded that the anion transporter required for
CCL20 release has an anion selectivity of gluconate< baseline and HDM-induced rises in CCL20 release in the nitrate-treated cells. This suggests that 105 the impact of nitrate on CCL20 release is independent of allergen exposure. Additionally, the difference between PBS and HDM in cells treated in chloride was about 75% less than the difference between PBS and HDM in cells treated in nitrate. However, the percentage increase of HDM over PBS treatment (about 66%) was the same for both treatment groups. This suggests that the impact of nitrate is on the overall rate of CCL20 release. To determine whether chloride dependence requires active or passive chloride transport, we eliminated any existing or potential chloride gradients by treating cells with TPPMn(III), which is a chloride ionophore. TPPMn(III) had no effect on either HDM-induced CCL20 release or DIDS inhibition (Fig 1D). This suggests the chloride transporter functions via an active mechanism, possibly a co-transporter. To confirm the ionophore was sufficient to equalize chloride concentrations across the epithelium, we measured intracellular chloride concentrations prior to and after HDM treatment in bronchial epithelial cells in the presence or absence of DIDS and TPPMn(III) (Fig 1E). We found that in cells treated with PBS or HDM alone, intracellular chloride decreased after treatment with HDM from baseline levels. Interestingly, cells treated with either DIDS or TPPMn(III) displayed intracellular levels of chloride at baseline that were equivalent to the HDM-treated cells. This evidence suggests that homeostatic levels of chloride are elevated above equilibrium in bronchial epithelial cells, and that they return to equilibrium in response to HDM. We next asked whether this chloride dependence of CCL20 release affects mobilization of chemokine from lysosomal stores. Cells transfected with CCL20-mCherry and loaded with LysoTracker were visualized in situ over the course of one hour of HDM exposure in various 106 anion replacement conditions (Fig 1F). Cells in media alone demonstrated CCL20 release from intracellular vesicles, while cells treated in the presence of DIDS did not mobilize intracellular CCL20 in response to HDM. This supports our hypothesis that the observed chloride- dependence of CCL20 release affects lysosomal CCL20 stores. Ion depletion modulates CCL20 release In addition to chloride, many other ions are physiologically important in bronchial epithelial cells. To determine which of these might play a role in modulating CCL20 release, we created media deficient in, or supplemented with, major ions. When we removed either magnesium or sulfate from bronchial epithelial cells, we found that intracellular chloride concentrations no longer decreased after HDM exposure (Fig 2A). Additionally, adding twice the normal level of magnesium sulfate induced intracellular chloride depletion even in the absence of HDM (Fig 2B). Removal of sulfate only marginally reduced HDM-induced CCL20 release, and removal of magnesium dramatically increased CCL20 release, while excess magnesium sulfate paradoxically had a similar effect as removal of magnesium (Fig 2C). The impact of magnesium appears to be restricted to baseline release, as the increased CCL20 release above baseline (HDM treatment minus PBS treatment) in control cells compared to magnesium deficient cells was identical, suggesting that the increase can be entirely attributed to increased constitutive CCL20 release. These effects persisted after 24 hours (Fig 2D). These data suggest that magnesium and sulfate have opposing roles in modulating CCL20 release. A major ion transporter family responsible for passage of both chloride and sulfate across the cell membrane is the SLC26A family of solute carrier proteins. We chose two 107 SLC26A family members that are expressed in bronchial epithelial cells, namely Pendrin (SLC26A4) and SLC26A9, as targets for siRNA knockdown (Fig 3 A, B). This knockdown had no effect on CCL20 release for either Pendrin or SLC26A9, suggesting magnesium and sulfate elicit their effects on intracellular chloride concentrations and CCL20 release through some other mechanism. Potassium is sufficient but not necessary to drive rapid CCL20 release In addition to reduction of magnesium, sulfate, and chloride, we tested whether removal of potassium from cell media would affect HDM-induced CCL20 release. We replaced the normal 5 mM potassium with sodium, and measured levels of intracellular chloride after HDM treatment. Potassium removal decreased intracellular chloride under homeostatic conditions (Fig 4C). However, this had no effect on CCL20 release (Fig 4B). This suggests that the effect of potassium on intracellular chloride concentration is independent of chemokine release, and that the presence of potassium in solution is not required for CCL20 release. We next tested whether addition of excess potassium would affect HDM-induced CCL20 release. Addition of potassium chloride also causes cell shrinkage due to osmotic changes. To control for this, we compared the effect of added potassium with excess sodium, a similar cation (Fig 4A). We observed that hyperosmolarity, from either potassium chloride or sodium chloride, increased CCL20 release, and that it particularly affected baseline release (PBS treatment). However, for baseline release, excess potassium allowed further increase of CCL20 release in comparison to excess sodium. Additionally, we treated cells with mannitol to induce cell shrinkage without changing the sodium or potassium concentration (data not shown). We observed no change in cells treated with mannitol compared with controls. Taken together, we found that while the 108 presence of extracellular potassium was not necessary for HDM-induced CCL20 release, the presence of additional potassium was sufficient to induce CCL20 release. The surprising absence of an effect from potassium removal may be explained by the fact that extracellular potassium concentrations are low relative to intracellular potassium concentrations. In our defined solutions, potassium concentrations were about 5 mM, compared to an estimated 120-140 mM intracellular potassium concentration for most cells. Thus, any gradient that was established under low potassium conditions may not be disrupted under null potassium conditions if the cell were able to retain high intracellular potassium. To further understand the mechanism of potassium-induced CCL20 release, we treated cells with the potassium ionophore, valinomycin. Valinomycin inserts into the cell membrane and selectively facilitates free diffusion of potassium ions, thereby eliminating any established potassium gradients. Valinomycin treatment alone had little or no effect on HDM-induced CCL20 release. However, when we added excess potassium to cells treated with valinomycin, we observed increased CCL20 release. This release was greater even when compared to the increased release seen with excess potassium alone (Fig 4D). Indeed, although addition of potassium alone increased baseline CCL20 release (with no apparent further increase in HDM- induced CCL20 release), after addition of valinomycin potassium addition increased both baseline release (PBS-treated cells) and HDM-induced CCL20 release. Thus the impact of potassium on release appears to differ depending on which pool of CCL20 is targeted. To determine whether there was an interaction between the effect of chloride and the effect of potassium on CCL20 release, we treated cells in gluconate replacement media with the 109 potassium ionophore, valinomycin. Strikingly, when we treated cells with chloride-to- gluconate replacement media concurrently with valinomycin, we saw that CCL20 release returned to normal levels for both baseline release, and for HDM-induced release (Fig 4E). Indeed, this effect was mirrored somewhat when we paired valinomycin with the ion transporter DIDS. However, DIDS plus valinomycin induced additional CCL20 release under control conditions as well as after HDM exposure. These data suggest a strong interaction between chloride and potassium in HDM-induced CCL20 release. Gap junctions are necessary for rapid CCL20 release Many ions and other small solutes are exchanged between cells via gap junctions. Changes in ion concentration that begin in one cell of the epithelium may affect the ion concentration of adjacent cells, thereby enhancing the initial signal. To test whether gap junctions affect the signaling that leads to rapid CCL20 release, we treated 16HBE14o- cells with different inhibitors of gap junctions, including 1-octanol (Fig 5A), phloretin (Fig 5B), or carbenoxolone (Fig 5C) for one hour prior to HDM treatment for two hours. We then measured CCL20 release into the supernatant. At this concentration, we observed that 1-octanol was highly cytotoxic, but that neither phloretin nor carbenoxolone displayed any appreciable cytotoxicity at the concentrations used (data not shown). We found that each inhibitor partially reduced HDM-induced CCL20 release, suggesting that gap junctions play a role in enhancing HDM-induced CCL20 release. 110 The JNK and Raf1 signaling pathways contribute to rapid CCL20 release We next sought to determine whether common signaling pathways implicated in initiating gene transcription and other downstream signaling pathways were required for rapid CCL20 release. To do so, we tested a panel of inhibitors of downstream signaling pathways. Table 1 outlines each of these inhibitors and their targets. Each inhibitor was compared with the appropriate solvent concentration as control [Ctrl (1)=1:500; Ctrl (2)=1:2,000; Ctrl (3)=1:25,000]. HDM was added one hour after incubation with inhibitor. Cells were allowed to incubate for a further 2 hours after HDM addition, after which supernatants were harvested for CCL20 ELISA. We found that most downstream signaling pathway inhibitors we tested had no effect on HDM-induced CCL20 release. However, the JNK inhibitor SP600126, and the Raf1 kinase inhibitor both significantly inhibited rapid CCL20 release, suggesting these pathways may be involved in HDM-induced CCL20 release. Interestingly, an activator of Raf1, ZM 336372, had no effect on HDM-induced CCL20 release, suggesting Raf1 is necessary but not sufficient to drive HDM-induced CCL20 release. We previously reported our finding that Raf1inhibition reduces HDM-induced CCL20 release. Other groups have demonstrated phosphorylation of ERK within 15 minutes of agonist exposure, concurrent with CCL20 release 50 Interestingly, we saw no decrease in HDM-induced CCL20 release after treatment with an inhibitor of MEK, a protein downstream of Raf1, and upstream of ERK.,51 suggesting that MEK (and potentially its downstream target ERK) is not 111 necessary for HDM-induced CCL20 release, although it may still be activated in response to HDM exposure. To individually test these panel data, we repeated this experiment with the inhibitors R031-8820, SB216763, (data not shown) and SP600126 (Fig 6A, B). SP600126 remained the only inhibitor to significantly reduce rapid CCL20 release. In addition, SP600126 treatment in the absence of HDM was sufficient to reduce intracellular chloride levels (Fig 6B), similar to DIDS treatment. These data suggest JNK signaling may play a role in HDM-induced CCL20 release upstream of chloride transporter involvement. Actin remodeling suppresses rapid CCL20 release The actin cytoskeleton is active in the priming and docking of vesicular stores. 52 In addition, many extracellular structural proteins associate with intracellular actin either directly or indirectly. We tested a panel of actin inhibitors to determine their effect on HDM-induced CCL20 release after 2 hours (Fig 7). The inhibitors we used, and their targets/functions, are listed in table 2. Although we hypothesized that inhibition of actin polymerization would decrease HDM-induced CCL20 release, due to the presence of intracellular stores of CCL20 within secretory lysosomes, we observed that inhibition of actin polymerization by any one of a variety of mechanisms was sufficient to increase rapid CCL20 release in response to HDM. These data suggest that under homeostatic conditions, functional actin polymerization contributes to the suppression of HDM-induced CCL20 release. 112 Mixed effects of reactive oxygen species on rapid CCL20 release To determine the role of reactive oxygen species (ROS) in rapid, HDM-induced release of CCL20, we treated cells with inhibitors targeting different pathways for generating ROS. Diphenylene iodinium (DPI) inhibits nitric oxide synthase and NADPH oxidase, while apocynin is a selective inhibitor of NADPH oxidase alone. When we treated cells with DPI and measured rapid CCL20 release, we observed that HDM-induced CCL20 release was diminished compared to control cells (Fig 8A). However, when we treated cells with apocynin we observed a dose-dependent increase in CCL20 release (Fig 8B). This suggests that nitric oxide synthase contributes to HDM-induced CCL20 release, while NADPH oxidase may antagonize HDM- induced CCL20 release. We next looked at the effect of inhibiting cyclooxygenase (COX) enzymes. There are two major COX enzymes, designated COX-1, and COX-2 – with COX-3 also having been described in recent years. Celecoxib selectively inhibits COX-2, 53 while aspirin inhibits COX-1 and modifies COX-2 away from production of prostanoids and toward lipoxins. 54 Treatment with either celecoxib or aspirin significantly reduced HDM-induced CCL20 release, suggesting that cyclooxygenase function is required for CCL20 release (Fig 9A, B). We next supplied the cells directly with the ROS precursor peroxide to determine if excess availability to this reactant would directly affect CCL20 release (Fig 9C). Peroxide (H 2O2) is a precursor to the generation of superoxide free radicals. Direct addition of peroxide to cell culture had no effect on CCL20 release. Thus, the effect of artificially increased peroxide in 113 solution may not reflect the function of increased generation of free radicals within the cells, which may have additional signaling roles. CCL20 release is not inhibited by corticosteroids Inhaled corticosteroids are an established treatment for long-term control of asthma symptoms. We asked whether corticosteroid treatment is capable of reducing CCL20 release from bronchial epithelial cells after exposure to HDM. We measured rapid CCL20, IL-6, IL-8, and GM-CSF release after exposure to HDM in cells that were treated with dexamethasone, a potent corticosteroid (Fig 10A). We found that dexamethasone treatment had no effect on rapid HDM-induced CCL20 release, even as it reduced release of IL-6, IL-8, and GM-CSF. When we looked at longer-term CCL20 release, after 24 hours HDM exposure (Fig 10B), we observed a small increase in CCL20 release, and again saw decreases in IL-6, IL-8, and GM-CSF. We saw this same trend in CCL20 release in vivo . We treated mice with HDM intra trachea , and looked at chemokine released into the broncho-alveolar lavage fluid (BAL) after four hours. We selected a four hour time point because this was when we saw maximal early release of CCL20. Similar to in vitro cultured cells, CCL20 was increased in the BAL of mice treated with HDM, as compared to PBS control (Fig 10C). Additionally, we observed a trend toward increased CCL20 release in mice concurrently treated with both HDM and dexamethasone, although this increase was not considered significant. Taken together, these data suggest that CCL20 release is controlled via a mechanism that is not subject to corticosteroid inhibition, as is the release of other chemokines and cytokines from bronchial epithelial cells. 114 Discussion In this chapter we identified a number of signaling pathways that affect CCL20 release, both under homeostatic conditions and after treatment with HDM. Pathways that are necessary for CCL20 release include raf1, cyclooxygenase, and gap junctions. These pathways were often necessary, but not sufficient, to drive CCL20 release; as in each case the presence of HDM was necessary to induce CCL20 release. For example, inhibition of Raf1 reduced CCL20 release, and decreased intracellular chloride. However, the Raf1 activator ZM 336372 had no effect on CCL20 release even after treatment with HDM, suggesting Raf1 is necessary, but not sufficient for CCL20 release. In contrast, excess potassium was sufficient, but not necessary to drive increased CCL20 release, even under homeostatic conditions. This effect was further enhanced in cells treated with valinomycin. Although the mechanisms driving this effect remain unclear, a role for potassium in countering the effect of chloride removal was suggested when we treated cells with valinomycin and either DIDS or gluconate-replacement media. Previously, we have seen no CCL20 release in the absence of chloride (gluconate replacement media). However, valinomycin overcame this effect and restored normal levels of CCL20 release. Free diffusion of potassium (valinomycin treatment) restored not only HDM-induced CCL20 release, but also baseline CCL20 release – which was almost entirely eliminated in gluconate-treated cells. This suggests that the interaction between potassium and chloride fundamentally affects all mechanisms of CCL20 release (intracellular and extracellular) and not just HDM-induced mechanisms. These processes remain to be further elucidated. 115 Although lysosome acidification has been shown to be aided by Cl -/H + co-transporters, some evidence also suggests potassium transport may play a similar role in lysosome acidification. 22 This may explain why altering potassium transport with valinomycin is able to compensate for the absence of chloride, and why removal of potassium had no effect on CCL20 release in the presence of chloride. We observed that baseline release of CCL20 was also eliminated in the absence of the anion chloride, but was enhanced by the cations potassium and magnesium. Previously we demonstrated that lysosome inhibition had no impact on baseline release of CCL20 (chapter 2), suggesting that these changes in ion concentration mainly impact release of extracellular CCL20. Other data from our lab (not shown) suggest that CCL20 binds to the heparin sulfate proteoglycan syndecan-1 on the cell surface. The syndecan binding motif is dependent on electrostatic interactions between heparin-sulfate and CCL20. Our observation that the addition of cations – such as potassium, magnesium, and sodium – into solution has the general effect of increasing CCL20 release may suggest the increased cation concentration interferes with these electrostatic interactions, and passively induces CCL20 dissociation from these heparin-sulfate stores at the cell surface. Additional experiments are required to confirm whether the increased CCL20 release observed in the presence of excess cations is derived from extracellular stores in general, and syndecan-1 in particular. Given that cation addition to the extracellular environment potentiated increased baseline release while leaving additional HDM-induced CCL20 release largely unaffected, our results suggest a model in which constitutive (not 116 allergen-induced) CCL20 release is derived from extracellular stores attached to syndecan-1, while the main source for HDM-induced CCL20 release is derived from intracellular stores. Although we observed a number of different treatments that were sufficient to induce reduction of intracellular chloride levels, this reduction alone was insufficient to induce CCL20 release in some instances. However, our observations suggest inhibitors that reduce CCL20 release correspondingly reduce intracellular chloride levels even in the absence of HDM. For example, both DIDS and SP600125 reduced intracellular chloride and CCL20 release. However, potassium depletion and TPPMn(III) also reduced intracellular chloride (potentially by disrupting homeostatic mechanisms driving increased intracellular chloride in these cells) but had no effect on CCL20 release. Thus a reduction in intracellular chloride appears to be necessary but not sufficient to induce release of CCL20. One likely mechanism driving this dependence is the membrane depolarization needed to induce secretory vesicle fusion with the cell membrane, similar to release of synaptic vesicles in neurons. 55 In neurons, synaptic vesicles are first docked to the cell membrane prior to fusion in a ‘priming’ step. 56 After this docking takes place, a depolarization event is sufficient to induce fusion with the cell membrane. A similar mechanism in bronchial epithelial cells could explain why reduced intracellular chloride does not always lead to CCL20 release. If CCL20-containing vesicles are not first primed, or if modulation of intracellular chloride does not represent a depolarization event, we would not expect reduced intracellular chloride to potentiate CCL20 release. Further experiments are necessary to determine whether changes in intracellular chloride concentration contribute to CCL20 release through depolarization or through another mechanism. 117 Interestingly, although we previously showed that impacting chloride transport into the lysosome (by knocking down Clcn7 ) partially reduced HDM-induced CCL20 release, when we inhibited chloride transport using a more global method (DIDS and NFA) we were unable to detect any CCL20 release. We previously hypothesized that knocking down Clcn7 partially inhibits CCL20 release because it does not affect extracellular chemokine stores. Given that general inhibition of chloride completely eliminates CCL20 release, it is likely that one or more additional chloride-dependent mechanisms impact other aspects of HDM-induced CCL20 release. Additional studies will be needed to outline these mechanisms in more detail. Increased levels of CCL20 are often observed in the BAL of severe asthmatics, a group that typically displays a weak response to inhaled corticosteroids. 57 Our data suggest a possible explanation for this, as even a potent corticosteroid like dexamethasone had no effect on HDM- induced CCL20 release. Indeed, at higher doses we saw a trend toward increased CCL20 release. These observations were mirrored in vivo . This is in contrast to other signaling molecules like GM-CSF, IL-6, and IL-8; which were all reduced after exposure to dexamethasone. It is therefore not surprising that steroid-resistant asthmatics have elevated levels of CCL20 in their BAL. Inhibition of reactive oxygen species generation via DPI reduced CCL20 release, but apocynin did not. Other groups have reported a link between the Raf1 pathway and ROS generation. Thus, CCL20 release may be downstream of a pathway that includes Raf1 signaling that promotes ROS generation. Given that increased ROS generation has been observed in 118 asthmatic airways, this suggests the production of ROS may serve as a danger signal to the cells to enhance release of preformed inflammatory mediators. 58 Contrary to our expectations, inhibition of actin polymerization resulted in increased CCL20 release, not decreased release. Additional experiments need to be done to further explain this observation, however based on our current understanding of vesicular release some hypotheses present themselves. For example, actin polymerization is required for endocytic reuptake of vesicles from the plasma membrane, including clathrin-coated endocytosis. 59 This reuptake could cycle syndecan-bound CCL20 from the cell surface under normal conditions. In this instance, inhibition of actin could result in increased cell-surface CCL20 and increased release due to HDM-induced syndecan shedding. Further experiments would be needed to demonstrate whether this is the case, or increased CCL20 release due to actin inhibition is due to another mechanism. We demonstrated here that gap junctions promote HDM-induced CCL20 release. Although the specific mechanism for this remains unclear, one attractive possibility is that the movement of ions through gap junctions allows signals that originate in one cell to propagate to adjacent cells. In one study, epithelial cells infected with intracellular bacteria induced intracellular signaling in uninfected neighboring cells, which responded to the infection of juxtaposed cells by releasing the chemokine IL-8. This mechanism was shown to be gap junction-dependent. A similar mechanism may work to enhance HDM-induced CCL20, leading to our observation of reduced release after gap junction inhibition. 29 In addition to this specific mechanism, gap junctions have been shown to allow the passage of small molecules that often 119 function as second messengers in intracellular signaling pathways. 60,61 In this way, gap junctions could also enhance intracellular signaling to nearby cells. We showed that a number of diverse signaling pathways contribute to the release of CCL20 from bronchial epithelial cells. This is unsurprising, as we observed changes in CCL20 release while handling and passaging the cells. Thus, it is likely that the cell responds to many different danger signals by releasing CCL20, thereby helping to recruit iDCs to the site of danger. Additional research will be necessary to show what other chemokines and cytokines released by the bronchial epithelium are similarly affected by these signaling pathways, and whether each pathway is active after exposure to different allergens. As discussed previously, many of the mechanisms we have explored were previously known to promote AHR via separate mechanisms. In addition to these known mechanisms, we demonstrated that they may also promote the release of CCL20 from the bronchial epithelium, suggesting that targeting these pathways could beneficial outcomes in regards to reduction of airway inflammation, in addition to these previously established effects. 120 References 1. Tapper H. The Secretion of Preformed Granules by Macrophages and Neutrophils. J Leukoc Biol . 1996;59(5):613–622. 2. Øynebråten I, Barois N, Hagelsteen K, et al. Characterization of a Novel Chemokine- Containing Storage Granule in Endothelial Cells: Evidence for Preferential Exocytosis Mediated by Protein Kinase A and Diacylglycerol. J Immunol . 2005;175(8):5358–5369. 3. Kunkel SL, Standiford T, Kasahara K, Strieter RM. Interleukin-8 (IL-8): the major neutrophil chemotactic factor in the lung. Exp. Lung Res. 1991;17(1):17–23. 4. Chintakuntlawar AV, Chodosh J. Chemokine CXCL1/KC and its Receptor CXCR2 Are Responsible for Neutrophil Chemotaxis in Adenoviral Keratitis. Journal of Interferon & Cytokine Research . 2009;29(10):657–666. 5. Kopf M, Gros GL, Bachmann M, et al. Disruption of the murine IL-4 gene blocks Th2 cytokine responses. , Published online: 18 March 1993; | doi:10.1038/362245a0 . 1993;362(6417):245–248. 6. Ather JL, Ckless K, Martin R, et al. Serum amyloid A activates the NLRP3 inflammasome and promotes Th17 allergic asthma in mice. J Immunol . 2011;187:64–73. 7. Besnard A-G, Guillou N, Tschopp J, et al. NLRP3 inflammasome is required in murine asthma in the absence of aluminum adjuvant. Allergy . 2011;66(8):1047–1057. 8. Blitz M, Blitz S, Hughes R, et al. Aerosolized Magnesium Sulfate for Acute Asthma*A Systematic Review. CHEST . 2005;128(1):337–344. 9. Cairns CB, Kraft M. Magnesium attenuates the neutrophil respiratory burst in adult asthmatic patients. Acad Emerg Med . 1996;3(12):1093–1097. 10. Skobeloff EM SW. INtravenous magnesium sulfate for the treatment of acute asthma in the emergency department. JAMA . 1989;262(9):1210–1213. 11. Fischer BM, Voynow JA. Neutrophil Elastase Induces MUC5AC Gene Expression in Airway Epithelium via a Pathway Involving Reactive Oxygen Species. Am. J. Respir. Cell Mol. Biol. 2002;26(4):447–452. 12. Seltzer J, Bigby BG, Stulbarg M, et al. O3-induced change in bronchial reactivity to methacholine and airway inflammation in humans. J Appl Physiol . 1986;60(4):1321–1326. 13. Murray JJ, Tonnel AB, Brash AR, et al. Release of Prostaglandin D 2 into Human Airways during Acute Antigen Challenge. New England Journal of Medicine . 1986;315(13):800–804. 14. Knight DA, Stewart GA, Lai ML, Thompson PJ. Epithelium-derived inhibitory prostaglandins modulate human bronchial smooth muscle responses to histamine. European Journal of Pharmacology . 1995;272(1):1–11. 15. Schmidt LM, Belvisi MG, Bode KA, et al. Bronchial epithelial cell-derived prostaglandin E2 dampens the reactivity of dendritic cells. J. Immunol. 2011;186(4):2095–2105. 16. Luster AD. Chemokines — Chemotactic Cytokines That Mediate Inflammation. New England Journal of Medicine . 1998;338(7):436–445. 17. Martinon F, Burns K, Tschopp J. The Inflammasome. Molecular Cell . 2002;10(2):417–426. 121 18. Agostini L, Martinon F, Burns K, et al. NALP3 Forms an IL-1β-Processing Inflammasome with Increased Activity in Muckle-Wells Autoinflammatory Disorder. Immunity . 2004;20(3):319–325. 19. Martinon F, Pétrilli V, Mayor A, Tardivel A, Tschopp J. Gout-associated uric acid crystals activate the NALP3 inflammasome. Nature . 2006;440(7081):237–241. 20. Mariathasan S, Weiss DS, Newton K, et al. Cryopyrin activates the inflammasome in response to toxins and ATP. Nature . 2006;440(7081):228–232. 21. Kanneganti T-D, Lamkanfi M, Kim Y-G, et al. Pannexin-1-Mediated Recognition of Bacterial Molecules Activates the Cryopyrin Inflammasome Independent of Toll-like Receptor Signaling. Immunity . 2007;26(4):433–443. 22. Qiu MR, Campbell TJ, Breit SN. A potassium ion channel is involved in cytokine production by activated human macrophages. Clinical & Experimental Immunology . 2002;130(1):67–74. 23. Malpuech-Brugère C, Nowacki W, Daveau M, et al. Inflammatory response following acute magnesium deficiency in the rat. Biochimica et Biophysica Acta (BBA) - Molecular Basis of Disease . 2000;1501(2–3):91–98. 24. Weglicki WB, Phillips TM, Freedman AM, Cassidy MM, Dickens BF. Magnesium- deficiency elevates circulating levels of inflammatory cytokines and endothelin. Mol Cell Biochem . 1992;110(2):169–173. 25. Soyombo AA, Tjon-Kon-Sang S, Rbaibi Y, et al. TRP-ML1 Regulates Lysosomal pH and Acidic Lysosomal Lipid Hydrolytic Activity. J. Biol. Chem. 2006;281(11):7294–7301. 26. Kominkova V, Malekova L, Tomaskova Z, et al. Modulation of intracellular chloride channels by ATP and Mg^2^. BBA - Bioenergetics . 2010;1797(6-7):1300–1312. 27. Kumar NM, Gilula NB. The Gap Junction Communication Channel. Cell . 1996;84(3):381– 388. 28. Bao X, Lee SC, Reuss L, Altenberg GA. Change in permeant size selectivity by phosphorylation of connexin 43 gap-junctional hemichannels by PKC. PNAS . 2007;104(12):4919–4924. 29. Kasper CA, Sorg I, Schmutz C, et al. Cell-Cell Propagation of NF-κB Transcription Factor and MAP Kinase Activation Amplifies Innate Immunity against Bacterial Infection. Immunity . 2010;33(5):804–816. 30. Adler V, Yin Z, Tew KD, Ronai Z. Role of redox potential and reactive oxygen species in stress signaling. Oncogene . 1999;18(45):6104–6111. 31. Thannickal VJ, Fanburg BL. Reactive oxygen species in cell signaling. Am J Physiol Lung Cell Mol Physiol . 2000;279(6):L1005–L1028. 32. Segal AW, Abo A. The biochemical basis of the NADPH oxidase of phagocytes. Trends in Biochemical Sciences . 1993;18(2):43–47. 33. Janssen-Heininger YMW, Mossman BT, Heintz NH, et al. Redox-based regulation of signal transduction: Principles, pitfalls, and promises. Free Radical Biology and Medicine . 2008;45(1):1–17. 34. Eu JP, Sun J, Xu L, Stamler JS, Meissner G. The Skeletal Muscle Calcium Release Channel: Coupled O2 Sensor and NO Signaling Functions. Cell . 2000;102(4):499–509. 122 35. Bolotina VM, Najibi S, Palacino JJ, Pagano PJ, Cohen RA. Nitric oxide directly activates calcium-dependent potassium channels in vascular smooth muscle. Nature . 1994;368(6474):850–853. 36. Gu Z, Kaul M, Yan B, et al. S-Nitrosylation of Matrix Metalloproteinases: Signaling Pathway to Neuronal Cell Death. Science . 2002;297(5584):1186–1190. 37. Levinthal DJ, DeFranco DB. Reversible Oxidation of ERK-directed Protein Phosphatases Drives Oxidative Toxicity in Neurons. J. Biol. Chem. 2005;280(7):5875–5883. 38. Kamata H, Honda S, Maeda S, et al. Reactive Oxygen Species Promote TNFα-Induced Death and Sustained JNK Activation by Inhibiting MAP Kinase Phosphatases. Cell . 2005;120(5):649–661. 39. Seth D, Rudolph J. Redox Regulation of MAP Kinase Phosphatase 3†. Biochemistry . 2006;45(28):8476–8487. 40. Cruz CM, Rinna A, Forman HJ, et al. ATP Activates a Reactive Oxygen Species- dependent Oxidative Stress Response and Secretion of Proinflammatory Cytokines in Macrophages. J. Biol. Chem. 2007;282(5):2871–2879. 41. Kuehl FA Jr, Egan RW. Prostaglandins, arachidonic acid, and inflammation. Science . 1980;210(4473):978–984. 42. Fu JY, Masferrer JL, Seibert K, Raz A, Needleman P. The induction and suppression of prostaglandin H2 synthase (cyclooxygenase) in human monocytes. J. Biol. Chem. 1990;265(28):16737–16740. 43. Feng L, Sun WQ, Xia YY, et al. Cloning Two Isoforms of Rat Cyclooxygenase: Differential Regulation of Their Expression. Archives of Biochemistry and Biophysics . 1993;307(2):361–368. 44. Chandrasekharan NV, Dai H, Roos KLT, et al. COX-3, a cyclooxygenase-1 variant inhibited by acetaminophen and other analgesic/antipyretic drugs: cloning, structure, and expression. Proc. Natl. Acad. Sci. U.S.A. 2002;99(21):13926–13931. 45. Loll PJ, Picot D, Garavito RM. The structural basis of aspirin activity inferred from the crystal structure of inactivated prostaglandin H2 synthase. Nature Structural & Molecular Biology . 1995;2(8):637–643. 46. Szczeklik A. The cyclooxygenase theory of aspirin-induced asthma. Eur Respir J . 1990;3(5):588–593. 47. Stevenson DD, Simon RA. Lack of cross-reactivity between rofecoxib and aspirin in aspirin-sensitive patients with asthma. Journal of Allergy and Clinical Immunology . 2001;108(1):47–51. 48. Snijdewint FG, Kaliński P, Wierenga EA, Bos JD, Kapsenberg ML. Prostaglandin E2 differentially modulates cytokine secretion profiles of human T helper lymphocytes. J Immunol . 1993;150(12):5321–5329. 49. Scherle PA, Jones EA, Favata MF, et al. Inhibition of MAP Kinase Kinase Prevents Cytokine and Prostaglandin E2 Production in Lipopolysaccharide-Stimulated Monocytes. J Immunol . 1998;161(10):5681–5686. 50. Reibman J, Hsu Y, Chen LC, Bleck B, Gordon T. Airway Epithelial Cells Release MIP- 3α/CCL20 in Response to Cytokines and Ambient Particulate Matter. Am. J. Respir. Cell Mol. Biol. 2003;28(6):648–654. 123 51. Macdonald SG, Crews CM, Wu L, et al. Reconstitution of the Raf-1-MEK-ERK signal transduction pathway in vitro. Mol. Cell. Biol. 1993;13(11):6615–6620. 52. Wit H de, Cornelisse LN, Toonen RFG, Verhage M. Docking of Secretory Vesicles Is Syntaxin Dependent. PLoS ONE . 2006;1(1):e126. 53. Goldstein JL, Silverstein FE, Agrawal NM, et al. Reduced risk of upper gastrointestinal ulcer complications with celecoxib, a novel COX-2 inhibitor. The American Journal of Gastroenterology . 2000;95(7):1681–1690. 54. Catella-Lawson F, Crofford LJ. Cyclooxygenase inhibition and thrombogenicity. American Journal of Medicine, The . 2001;110(3):28–32. 55. Rizo J, Rosenmund C. Synaptic vesicle fusion. Nature Structural & Molecular Biology . 2008;15(n7):665–674. 56. Pevsner J, Hsu S-C, Braun JEA, et al. Specificity and regulation of a synaptic vesicle docking complex. Neuron . 1994;13(2):353–361. 57. Hastie AT, Moore WC, Meyers DA, et al. Analyses of asthma severity phenotypes and inflammatory proteins in subjects stratified by sputum granulocytes. Journal of Allergy and Clinical Immunology . 2010;125(5):1028–1036.e13. 58. Henricks PAJ, Nijkamp FP. Reactive Oxygen Species as Mediators in Asthma. Pulmonary Pharmacology & Therapeutics . 2001;14(6):409–420. 59. Saheki Y, Camilli PD. Synaptic Vesicle Endocytosis. Cold Spring Harb Perspect Biol . 2012;4(9): 60. Sáez JC, Connor JA, Spray DC, Bennett MV. Hepatocyte gap junctions are permeable to the second messenger, inositol 1,4,5-trisphosphate, and to calcium ions. PNAS . 1989;86(8):2708–2712. 61. Anselmi F, Hernandez VH, Crispino G, et al. ATP release through connexin hemichannels and gap junction transfer of second messengers propagate Ca2+ signals across the inner ear. PNAS . 2008;105(48):18770–18775. 124 Downstream signaling inhibitors used in figure 2 Inhibitor Target Concentration SP600126 JNK (inhibitor) 10 uM R031 -8220 PKC (inhibitor) 10 uM Ly294002 PI3Kinase (inhibitor) 10 uM Raf 1 kinase inhibitor Raf1 kinase (inhibitor) 10 uM Bafilomycin V-ATPase (inhibitor) 0.5 uM Pertussus toxin Increases cAMP 100 ng/mL ZM 336372 Raf1 (activator) 10 uM U0126 MEK1/2 (inhibitor) 10 uM SB216763 GSK -3 (inhibitor) 10 uM CPG 57380 MNK1 (inhibitor) 10 uM Wortmannin PI3Kinase (inhibitor) 1 uM Pimelic diphenylamide 106 HDAC 1, 2, 3, 8 (inhibitor) 2 uM PP242 mTOR (inhibitor) 1 uM Rapamycin mTOR (inhibitor) 0.1 uM MS -275 HDAC 1>3 (inhibitor) 0.5 uM Trichostatin HDAC 1, 3, 4, 6, 10 (inhibitor) 0.1 uM SAHA HDAC (inhibitor) 1 uM EX -527 SIRT1 (inhibitor) 0.1 uM CAY10603 HDAC 6 (inhibitor) 0.1 uM IKK2 inhibitor NFkappaB (inhibitor) 1 uM GSIxxx Gamma secretase/Notch (inhibitor) 1, 10, 100 nM Table 1 125 Actin inhibitors and their functions Inhibitor Function Cytochalasin D Binds G -actin and prevents incorporation into filaments Jasplakinolide Prevents F -actin extension and polymerization Latrunculin B Prevents G -actin polymerization Swinholide A Inhibits actin polymerization, and severs F -actin Table 2 126 - Chloride nitrate-containing media (NO3 ) with or without DIDS were treated with HDM for 2 hours prior 300 150 Gluconate A *** B *** *** to harvesting supernatants for ELISA. (D, E) Cells treated with and without TPPMn(III) and F 200 PBS 100 HDM (min) DIDS were exposed to HDM for 2 hours prior to harvesting supernatants for ELISA (D) or for 30 HDM 0 30 60 minutes prior to measurement of intracellular chloride. (F) Cells transfected with CCL20- CL20 (pg/mL) CL20 (pg/mL) 100 50 ** ** C C acker r mCherry (red) and loaded with LysoTracker dye (green) were treated with DIDS or vehicle T ND ND ND s o y 0 0 L control. Cells were then imaged for 60 minutes after treatment with HDM. Data are repr- Ctrl DIDS NFA 2 h 2 h + Cl- add back ol esented as mean +/- SEM. *p<0.05 compared to control PBS treatment t r o n 300 150 CL20-mCh C C *** D C *** *** *** PBS y 200 100 l a * PBS e r HDM v HDM O CL20 (pg/mL) 100 CL20 (pg/mL) 50 C C *** ND ND ND acker 0 0 r - - - - T Cl NO3 Cl NO3 DMSO TPP DMSO TPP s o y DIDS DIDS L DIDS CL20-mCh E 0.08 * C * ** PBS 0.06 y ] (mM) HDM - l a e r Cl v 0.04 O ellular [ a c 0.02 t r n I 0.00 DMSO DIDS TPP Figure 1 – A chloride co-transporter is necessary for HDM-induced CCL20 release. (A) After incubation with DIDS and ni umic acid (NFA) cells were treated with HDM for 2 hours prior to harvesting supernatants for ELISA. (B) Cells incubated in media containing either normal chloride or gluconate nitrate were treated with HDM for 2 hrs. Supernatants were then harvested and cells were further incubated with normal chloride for another two hours prior to harvesting supernatants again (add back). (C) Cells treated in either normal chloride (Cl-) or - nitrate-containing media (NO3 ) with or without DIDS were treated with HDM for 2 hours prior to harvesting supernatants for ELISA. (D, E) Cells treated with and without TPPMn(III) and DIDS were exposed to HDM for 2 hours prior to harvesting supernatants for ELISA (D) or for 30 minutes prior to measurement of intracellular chloride. (F) Cells transfected with CCL20- mCherry (red) and loaded with LysoTracker dye (green) were treated with DIDS or vehicle control. Cells were then imaged for 60 minutes after treatment with HDM. Data are repr- esented as mean +/- SEM. *p<0.05 compared to control PBS treatment A-D n=3, E n=6 0.05 A PBS ** HDM ** ] - 0.04 0.03 0.02 Intracellular [Cl Intracellular 0.01 0.00 2+ 2- Ctrl Mg SO4 Ion removed *** *** *** *** *** *** Figure 2 – Magnesium and sulfate are required for chloride depletion after HDM treatment. (A) HBE cells were incubated in media containing (ctrl) or def cient in magnesium (Mg2+) or 2- sulfate (SO4 ) ions prior to HDM treatment for 30 minutes and measured for intracellular chloride concentration. (B) Cells were incubated with balanced ions (ctrl) or in the presence of excess magnesium sulfate (+3.6mM MgSO4) prior to HDM treatment for 30 minutes and measured for intracellular chloride concentration. (C and D) Cells were treated as above with either suf cient ion concentrations (ctrl), sulfate def cient media (no SO4), magnesium def - cient media (no Mg), or excess magnesium sulfate (Ctrl + MgSO4); cells were then treated with HDM for either 2 hours (C) or 24 hours (D) prior to harvest of supernatants. Data are repre- sented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001 compared to control HDM treatment. A-B n=6, C-D n=3. ns ns A 80 ns B 50 *** *** PBS PBS HDM HDM 40 60 30 40 20 20 10 0 0 Ctrl Pdr A9 Ctrl Pdrn siRNA (100pM) Pendrin siRNA 500 pM Figure 3 – siRNA against the Cl-/SO42- exchangers pendrin and SLC26A9 have no ef ect on HDM-induced CCL20 release. HBE cells were electroporated with 100 pM (A) or 500 pM (B) of siRNA targeted against pendrin (Pdr) or SLC26A9 (A9). 72 hours later, cells were treated with HDM for 2 hours and supernatants were harvested for ELISA. n=6. B A 500 250 PBS C 0.08 *** PBS ** HDM ns HDM *** 400 *** 200 - *** *** 0.06 PBS HDM 300 150 ** 0.04 200 *** 100 0.02 100 50 Cl Intracellular 0 0 0 Ctrl NaCl KCl Ctrl -K+ Ctrl -K+ D 100 E 40 PBS *** PBS HDM HDM 80 *** 30 *** 60 20 ††† * 40 ††† 10 20 0 0 Ctrl K+ Ctrl K+ Cl- Cl- Gl- DIDS Valinomycin Valinomycin Figure 4 – Potassium is suf cient, but not necessary for HDM-induced CCL20 release. CCL20 release from HBE cells treated with HDM for 2 hours (A) in the presence of excess sodium (NaCl) or potassium (KCl), (B) in the absence of potassium (-K+), (D) in the presence of excess potassium (K+) and/or valinomycin, or (E) in anion inhibition conditions (gluconate or DIDS- treated) with valinomycin. (C) Intracellular chloride concentration after HDM treatment in HBE cells treated in potassium-free conditions. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001 compared to control HDM treatment. tp<0.05, ttp<0.01, tttp<0.001. A-B and D-E n=3, C n=6. A B 150 PBS 200 PBS HDM HDM 150 100 ** *** *** 100 * ** 50 50 0 0 Ctrl 0.1 0.5 1.0 Ctrl Low HighMed 1-octanol Phloretin C 150 PBS HDM 100 ** 50 0 Ctrl Carb Figure 5 – Gap junction inhibition reduces HDM-induced CCL20 release. Cells were treated with gap junction inhibitors 1-octanol (A), phloretin (B), or carbenoxolone (C) one hour prior to HDM treatment. Two hours after HDM treatment, CCL20 release into the supernatant was measured by ELISA. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001. n=3. 80 B A PBS 0.08 PBS HDM ** * HDM 60 * - 0.06 40 0.04 20 0.02 Intracellular Cl Intracellular 0 0 Ctrl SP6 Ctrl SP6 Figure 6 – SP600126 inhibits CCL20 release and baseline chloride import. HDM-induced CCL20 release after 2 hours in HBE cells treated with selected inhibitors from f gure 1, (A) SP600126. (B) Intracellular chloride concentration after HDM treatment in HBE cells treated with SP600126. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, compared to control HDM treatment. A n=3, B n=6. *** *** ** 250 *** * 200 150 PBS * 100 HDM 50 0 Ctrl Cyto Jasp Lat Swin Figure 7 – Inhibition of structural proteins alters HDM-induced CCL20 release. HDM-induced CCL20 release after 2 hours in HBE cells treated with various inhibitors. Ctrl=control, Cyto=cytochalasin D, Jasp=jasplakinolide, Lat=latrunculin, Swin=swinholide A, Carb=carbenoxolone. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001 compared to control HDM treatment. n=3. A B 80 PBS 150 PBS HDM HDM 60 ** 100 *** *** 40 *** 50 *** 20 0 0 Ctrl DPI Ctrl Low HighMed Apocynin Figure 8 – Inhibition of dif erent ROS generation pathways dif erentially modulated CCL20 release. HBE cells were treated with the ROS inhibitors DPI (A) or apocynin (B) one hour prior to HDM treatment. Two hours after HDM treatment, CCL20 release into the supernatant was measured by ELISA. Data are represented as mean +/- SEM. **p<0.01, ***p<0.001 compared to control HDM treatment. n=3. PBS A 250 * B 250 *** C 80 HDM * ** 200 PBS 200 PBS 60 HDM HDM 150 150 40 100 100 20 50 50 0 0 0 Ctrl 0.5 5 Ctrl 3 10 Ctrl H2O2 Aspirin (uM) Celecoxib (uM) Figure 9 – Inhibition of dif erent cyclooxygenase pathways dif erentially modulated CCL20 release. HBE cells were treated with inhibitors of dif erent COX proteins; including celecoxib (A), and aspirin (B); one hour prior to HDM treatment. (C) Alternately, peroxide was added to the culture one hour prior to HDM treatment. Two hours after HDM treatment, CCL20 release into the supernatant was measured by ELISA. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001. n=3. PBS A 80 *** PBS B 600 PBS C 1500 *** *** HDM HDM HDM *** ) ) L 60 L *** ns m *** m / / 400 1000 g g p p ( ( 40 * 0 0 2 2 L L 200 C 500 C C 20 C CCL20 in BAL (pg/mL) 0 0 0 Ctrl Dex *** 50 *** PBS 150 PBS * *** HDM *** HDM 40 ** *** 100 *** *** 30 ** ** 20 50 10 GM-CSF (pg/mL) GM-CSF (pg/mL) 0 0 *** PBS *** HDM *** 100 600 *** *** ** PBS *** *** HDM 80 2 hour HDM treatment ** 24 hour HDM treatment * 400 60 *** ** 40 200 IL-6 (pg/mL) 20 IL-6 (pg/mL) 0 0 *** 150 PBS 300 *** PBS *** HDM *** *** HDM * * 100 200 50 100 * IL-8 (pg/mL) IL-8 (pg/mL) 0 0 Ctrl 1 uM 10 uM Ctrl 1 uM 10 uM Ctrl + DEX Ctrl + DEX A-B n=3, C n=6. Chapter 4. Allergen-induced release of many preformed chemokines and cytokines is through secretory lysosomes Mark Webb*, Marsha Wills-Karp** * Division of Allergy and Immunology, Cincinnati Children’s Hospital Medical Center, 3333 Burnet Ave, Cincinnati, OH, 45229 ** Department of Environmental Health Sciences, Johns Hopkins School of Public Health, Baltimore, MD, USA, 21205 139 Abstract A variety of chemokines and cytokines have been shown to be released into the broncho-alveolar lavage fluid in asthmatics. These act as inflammatory mediators that provide many of the signals that drive asthma pathogenesis. However, the mechanisms controlling their release are poorly characterized. Using a multiplex assay, we measured levels of preformed stores of a broad panel of chemokines and cytokines in bronchial epithelial cells, and found preformed stores of IL-6, IL-8, gro-α, TNF-α, IL-1α, GM-CSF, VEGF, and CCL20 among others. Of these, a subset including IL-6, IL-8, gro- α, TNF-α, IL-1α and CCL20 were released in response to HDM exposure. Using multi- color confocal microscopy and flow cytometry of subcellular organelles, we investigated whether multiple mediators were stored in the same subcellular location. Our data suggest that some mediators, such as gro-α and CCL20, localize to the same subcellular compartments. We also found that release of some, but not all, of these subcellular stores was induced by HDM. We then measured rapid release of various chemokines and cytokines in response to a panel of allergens, including particulate matter, cockroach frass, LPS, Fel d 1, and various pollens; we then compared these allergens to control and HDM stimulation. We found that HDM induced greater overall chemokine and cytokine release. We also found that different allergens induced release of different sets of chemokines from bronchial epithelial cells. These findings identify a mechanism of preformed lysosomal stores for multiple chemokines and 140 cytokines. These are rapidly released from bronchial epithelial cells in response to different allergens. 141 Introduction Multiple inflammatory mediators are released into asthmatic airways in response to allergen exposure. 1 These include CCL20, Gro-a, IL-1a, IL-6, and IL-8, among others. 2 As discussed previously, these chemokines and cytokines mediate many of the phenotypic responses observed in asthmatics, including inflammation and airway hyperresponsiveness. 3 The mechanisms for release of each of these various chemokines and cytokines have previously been poorly defined. Although they are often thought to be released through direct vesicular export from the golgi, 4 storage and rapid release have been observed for other chemokines and cytokines in addition to CCL20, 5–7 suggesting their release may be mediated by a similar mechanism of secretory lysosome release to what we have seen previously for CCL20. In some cell types, a variety of different secretory lysosomes may reside within an individual cell. 8 Cells from the hematopoietic lineage are among the most studied examples of this granular release, where cytotoxic T lymphocytes, neutrophils, and natural killer cells all contain these granules. Three major types of granules have been identified in neutrophils. 9 Primary granules, or azurophils, contain antimicrobial peptides called defensins, neutrophil elastase, and cathepsin G;10 secondary granules, also called specific granules, contain lactoferrin, collagenase, and lysozyme;11,12 and tertiary granules, or gelatinase granules, contain gelatinase, lysozyme, and leukolysin.13,14 Thus, the cell can respond to different stimuli by selective secretion of 142 different types of subcellular granules. 15–18 Although secretory granules are normally thought to reside in hematopoietic cells, recent studies have shown similar granules in other cell types as well. 19 Previously, we identified CCL20 within secretory lysosomes in bronchial epithelial cells, noting that the signal peptide for CCL20 is most closely related to Cathepsin C. This CCL20 was released after HDM exposure in a similar manner observed in release of secretory granules of other cell types. This suggests that cells of the bronchial epithelium may also contain secretory granules that may be selectively released. We hypothesized that multiple chemokines and cytokines that are released in response to HDM exposure are contained in the same lysosomal granules. A major challenge in subcellular tracking of multiple chemokines and cytokines simultaneously lies in the limitations of techniques currently developed to identify subcellular localization. Currently, analysis of the contents of subcellular organelles is accomplished by a number of methods, including immuno-transmission electron microscopy (TEM), confocal microscopy, density-dependent centrifugation, and single organelle fluorescence analysis (SOFA). These methods each present practical limitations. 20 In immuno-TEM, samples are stained with antibodies containing gold- conjugated particles of different sizes. 21 The antibodies appear as dark (electron-dense) 143 dots in the TEM image. Visual distinction between antibodies is accomplished through subjective size comparison. Because of this, effectively distinguishing between more than two antibodies becomes increasingly subjective. 22 Additionally, colocalization studies using TEM remain qualitative analyses, as images with sufficient resolution to detect immune-stained particles represent a smaller area of the cell, making quantification difficult. With density-dependent centrifugation, cell lysates are centrifuged in a density gradient to separate specific subcellular components. 23 This method is useful for quantifying protein differences between fractions of different densities, but it is incapable of distinguishing the contents of subcellular organelles of similar densities. 24 Confocal microscopy has sufficient resolving power to identify individual lysosomes, and many proteins may be targeted with fluorescently labeled antibodies within a cell. 25 However, superposition of fluorophores observed by confocal microscopy does not necessarily imply colocalization, as two proteins may appear together in a 2D image, but be in different cell compartments when viewed on the z- axis. 26 This effect can be minimized with confocal microscopy, but when visualizing smaller vesicles, apparent colocalization should still be verified using alternate methods. 144 A relatively newer method that has shown some promise in identification of subcellular organelles has been named SOFA for single organelle fluorescence analysis. However, a major limitation to SOFA is the necessity of transfecting the target cells with fluorescently-tagged proteins for visualization. Previously, fluorescent labeling of subcellular components for SOFA has been limited to cellular dyes, which preferentially insert into specific organelles, or to transfection of the target cells with a plasmid containing a fluorescently-tagged protein of interest. 20,27 In the present study, we use density-dependent centrifugation, confocal microscopy, and a modified application of SOFA to assess whether multiple chemokines and cytokines reside in the same subcellular organelles. As discussed previously, atopic asthma is associated with sensitivity to a number of different allergens. These allergens have many attributes in common that contribute to their allergenicity, such as enzyme activity, LPS, beta-glucans, and chitins.28–32 In addition to these individual factors, many allergens are a complex mix of multiple factors. Previously, we have focused on HDM for our model allergen, as it is a very potent allergen, and a majority of atopic asthmatics display sensitivity toward HDM. 33 Given that many of these allergens share similar features that have been linked to their allergenicity, we hypothesized that exposure to each allergen may induce release of many of the same chemokines as HDM. 145 Although we are interested in the effect of various allergens on secretory granule release, allergen challenge induces the release of a broad array of chemokines and cytokines. This release may be mediated by a variety of mechanisms. We previously showed that CCL20 release is governed by secretory lysosome release. This suggests a mechanism of granular storage that is found in other cell types, and is a major mechanism for the simultaneous release of a number of inflammatory mediators. Therefore we hypothesized that multiple chemokines and cytokines are stored in the same granules within bronchial epithelial cells; that these granules may be selectively released by allergen exposure; and that different allergens induce release of different subsets of chemokines and cytokines. To determine whether multiple types of secretory lysosome are secreted from 16HBE14o- cells in response to HDM, we combined confocal microscopy with SOFA. In order to stain multiple targets with SOFA, we developed a direct antibody staining technique similar to traditional flow cytometry. We found multiple chemokines localized within individual subcellular organelles, with some evidence that multiple combinations of chemokines were contained within different granules. Additionally, we found that chemokine and cytokine release was specific to allergen exposure. 146 Materials and Methods Cell culture and treatment: The 16HBE14o- human bronchial epithelial cell line was cultured in 10-15 cm round culture dishes that had been collagen coated. Cells were passaged by applying trypsin (0.05% with EDTA) for 10 min. Treatments were performed in 24-well collagen-coated culture plates in DMEM including 10% FBS, penicillin/streptomycin, and L-glutamine. Upon reaching 70-80% confluence, media was replaced and FBS reduced to 0.1% overnight. 34 Where listed, the inhibitor DIDS (10 uM) was added one hour prior to allergen addition. Inhibitors: Reagents used as inhibitors were obtained from SigmaAldrich unless specified and were used at the following concentrations: DIDS (10 uM). HDM treatment: Whole HDM extract was obtained from Greer (B70). 100 ug/mL HDM was added to cell supernatant for two hours prior to harvesting cells. ELISA : Mouse and human ELISAs were performed in half-well plates using DuoSet ELISA kits from R&D Systems according to their specifications. Multiplex quantification was performed via Luminex using proprietary kits and 50 uL sample volume. Samples were from BAL or cell supernatants that had been frozen at -80 oC. Mouse asthma model: Sensitization to HDM was achieved by intra tracheal (I.T.) administration of 100 ug/mL HDM. Twenty-one days later a second administration of alexafluor 405-labelled HDM was administered I.T. 72-hours after the last HDM 147 challenge mice were anesthetized with sodium pentathol and airway measurements were taken. The mice were then sacrificed, and broncho-alvelar lavage was performed using 1 mL Hanks balanced salt solution without calcium or magnesium. BAL fluid was the centrifuged to remove cells. Lungs were removed and fixed in formaldehyde, then sent to core personnel at CCHMC who embedded in paraffin, sectioned, and mounted them on slides. Airway measurements: 72 hours after the final allergen exposure, mice were anesthetized, intubated, and ventilated at 120 breaths/min. After airway pressure stabilized, mice given 25 ug/kg weight of acetylcholine intra venous and airway pressure changes were follow for five minutes. Airway measurements were quantified as airway pressure time index (APTI). Isolation and culture of mouse tracheal epithelial cells (mTECs): Mice were anesthetized with sodium pentathol and the inferior vena cava was severed to prevent recovery. Trachea were then removed and incubated overnight at 4 OC in 1% pronase (Roche) in DMEM/F12 (50/50) mix (Fisher). Cells were then removed from trachea via sheer force in the presence of 10% FBS and DNase. Cells were then incubated on Primaria plates for four hours to allow fibroblasts to adhere. Non-adherent cells were then removed and grown on rat-tail collagen-coated trans-well plates in DMEM/F12 supplemented with penicillin/streptomycin, L-glutamine, retinoic acid, and 5% FBS. After reaching 148 1000 ohms trans-epithelial resistance, media was replaced with 0.1% FBS media overnight prior to experimental treatments. Allergen treatments: Allergens were added to cell supernatant at the following concentrations: HDM (100 ug/mL), cockroach frass, LPS, ragweed pollen (100 ug/mL), ambient urban Baltimore particulate matter (AUB) (50 ug/mL), Fel D 1, alder pollen (50 ug/mL), birch pollen (50 ug/mL), maple pollen (50 ug/mL). Degranulation assay: Lamp-1 antibody conjugated to Af647 was added to cell culture concurrently with allergen. After 30 minutes, cells were rinsed with PBS, and removed from culture plate with trypsin. Antibody uptake was then analyzed on a BD LSR Fortessa flow cytometer. Immunofluorescence staining: Cells were grown and treated in 8-well chamber slides. After treatment, slides were fixed in 4% PFA for 15 minutes and rinsed with PBS. Cells were then permeablized with 0.1% triton-x 100, and blocked in 10% rabbit serum. Primary antibody was incubated overnight, followed by rinse with PBS-tween 0.1%. Secondary antibodies obtained from Invitrogen were incubated for one hour, then cells were rinsed with PBS-tween and mounted with VectaShield (Vector Labs) in preparation for imaging. Immunofluorescence staining on fixed lungs: Paraffin was removed from tissues on slides by immersing them in xylene. Slides were subsequently immersed in 100% Ethanol and 149 rehydrated by transferring them to 90%, 80%, 70% ethanol, and finally water. Antigen retrieval was accomplished by incubating slides in sodium citrate buffer. Slides were blocked in 10% serum matching the host species of the secondary antibody diluted in PBS-tween 0.1%, followed by application of primary and secondary antibodies to the slides. Slides were then mounted with VectaShield in preparation for imaging. Sucrose gradient fractionation: Cells were cultured as described above, removed from culture via trypsin, and transferred to 370 mM sucrose solution to prevent aggregation of subcellular organelles. Cells were then lysed via Misonix sonicator for 15 cycles. Nuclei were removed by spinning down cells at 3000 x g for 10 minutes. Supernatants were then run on a Percoll gradient of 1.05 g/mL. Fractions were removed serially from the top of the tube, and lysed with RIPA buffer that contained protease inhibitors. Final lysed fractions were then analyzed by ELISA. Single organelle fluorescence analysis: Cells were cultured and treated as described above, and placed in suspension by trypsin removal. Cells were then stained with antibodies as described above under flow cytometry . Cells were rinsed in 370 mM sucrose that had been filtered through a 0.2 uM membrane, and lysed using a Misonix sonicator for 10 cycles. Finally, cells were centrifuged at 5000 x g for 10 minutes to remove nuclei, and analyzed via flow cytometry. Voltages for photo-multiplier tubes were set by 150 comparison of 0.2 to 1.2 uM beads, and threshold values were reduced to accommodate the smaller event sizes. Statistical analysis: Statistical analysis was performed using GraphPad Prism 5 software. Statistical significance was determined using an unpaired student’s t-test for comparisons between two treatments, or two-way ANOVA combined with Bonferroni post-test analysis for comparison between multiple groups. Values of p<0.05 were considered statistically significant. 151 Results Blockade of CCL20 in vivo does not reduce airway responsiveness As mentioned previously, knockout mice for the only known receptor of CCL20, CCR6, displayed decreased airway hyperactivity in vivo , as demonstrated by other groups. 35 We wanted to determine whether directly blocking CCL20 itself in vivo would also lead to decreased airway responsiveness, and a decrease in airway inflammation. To do this, we treated mice concurrently with HDM and CCL20 neutralizing antibodies to suppress the signal of CCL20 after HDM release. We did this at either sensitization (Fig 1A,C the first day of HDM treatment), or at challenge (Fig 1B,D two weeks after the first HDM treatment); then measured airway responses, and BAL cellularity 72 hours after the last challenge. Eight mice were used in the treatment groups, and four were used in the PBS control groups. We observed a small decrease in airway responses under both conditions; however this decrease was statistically significant. A small, though insignificant, decrease was also observed in BAL cellularity. A high degree of variability in the treatment groups suggests this experiment does not have sufficient power to resolve whether blocking CCL20 has a substantive effect on airway inflammation and AHR. Although increasing the power of the experiment may allow us to resolve whether there is a statistically significant decrease in AHR and airway inflammation after blocking CCL20, given our current results this decrease is likely to be small compared to the overall effect of HDM on the mouse airways. As such, it is 152 unlikely that elimination of CCL20 signaling itself is sufficient to eliminate HDM- induced AHR and inflammation. Sub-cellular stores of other mediators are found in bronchial epithelial cells The observation that CCL20 blockade only partially decreases airway responsiveness and inflammation suggests that additional chemokines and cytokines also contribute to the early recruitment and activation of DCs and the initiation of inflammation and AHR in response to HDM. Indeed, DCs have been shown to reside in the lungs under homeostatic conditions, although not in the numbers seen after allergen challenge. 36 Additionally, many other cytokines and chemokines released into the BAL of asthmatics have been shown to induce inflammation of other cell types in addition to DCs. 2,37 In chapter 2, we outlined an intracellular storage and release mechanism for CCL20. We hypothesized that other chemokines and cytokines may also be stored within the bronchial epithelium, and that they may be released under a similar mechanism. To determine which chemokines and cytokines may share a common release mechanism, we first asked which mediators are present in preformed stores within the bronchial epithelium. To do this, we analyzed cell lysates via the multiplex Luminex assay, which measures numerous analytes simultaneously. We measured additional targets by ELISA. Table 1 includes a list of the targets we measured. We found a number of chemokines and cytokines that were stored in bronchial epithelial cells 153 under homeostatic conditions (Fig 2a). Some targets that were pre-formed in the cell lysate at the highest levels include gro-α, IL-8, IL-6, and IL-1α. CCL20 stores were orders of magnitude lower than these higher concentration proteins. It was not the lowest measured chemokine stored, however. Additionally, a background level of CCL20 is released by these cells, similar to what has been observed previously, along with background release of other chemokines such as IL-8 and FGF-2. Low-level, constitutive IL-8 release from endothelial cells has been reported by other groups. 38 We next set out to determine which chemokines and cytokines that are present in the lysate are also released after HDM exposure. We treated 16HBE14o- cells with HDM for 2 hours and collected the supernatants to measure chemokine and cytokine release via multiplex assay (Fig 2b). We found that many, but not all, chemokines and cytokines that were pre-formed were released by HDM. Also, release was not directly proportional to initial levels of preformed stores. For example, preformed stores of CCL20 were much lower than those of FGF-2, but after HDM treatment we observed more CCL20 had been released into the supernatant than FGF-2. We saw this same phenomenon with IL-8 and IL-6. While slightly more IL-8 was measured in the cell lysate than IL-6, HDM-induced release of IL-6 was almost twice that of IL-8. These data suggest that significant amounts of pre-formed stores are not released in response to HDM exposure. 154 To test whether chemokines other than CCL20 are stored in similar intracellular compartments as CCL20, we used density-dependent fractionation in a Percoll gradient to separate the subcellular components of cell lysates. Fractions removed from the top of the gradient were tested for the presence of CCL20, gro-α, IL-17f, and TSLP via ELISA (Fig 3). We found that each of these chemokines and cytokines were stored within bronchial epithelial cells. Gro-α and CCL20 localized to the same cell fractions, while IL-17f and TSLP localized to unique fractions, and therefore presumably different subcellular locations. This experiment is insufficient to determine whether gro-α and CCL20 are colocalized, but only measures whether they reside in vesicles of similar specific gravity. Also, measurement of lysosomal markers is required to further determine which of these cell fractions represent lysosomal vesicles. However, lysosomes are known to have multiple specific gravities and hence to separate to multiple fractions during density-dependent centrifugation. 39 To further determine the subcellular localization of chemokines and cytokines stored in bronchial epithelial cells, we visualized cells grown on slides using confocal microscopy. Cells were stained with antibodies against GM-CSF, gro-α, IL-8, and CCL20 (Fig 4). We saw robust staining of GM-CSF, gro-α, IL-1α, and IL-8; but very little staining of CCL20. This may reflect the absolute abundance of these proteins within the cells (indeed, this matches our observations in whole-cell lysates from figure 2A) or the antibodies used to stain. While subcellular localization appeared in punctate 155 bodies in GM-CSF, gro-a, and IL-8; staining of IL-1α appeared diffuse, with no apparent vesicular localization. Additionally, IL-1α and IL-8 staining localized primarily within the nucleus, with some staining outside the nucleus as well. This may explain why high levels of stored IL-8 in cell lysates did not translate to correspondingly high levels of IL- 8 release after treatment with HDM (Fig 2). In contrast, GM-CSF appeared to have some nuclear staining, and gro-α showed no nuclear staining. Establishing a protocol for single organelle fluorescence analysis To further distinguish the movement and release of individual chemokine and cytokine-containing vesicles, we sought to analyze single organelles via flow cytometry. Previous groups have reported performing single organelle fluorescence analysis (SOFA) by transfecting cells with one or more fluorescent probes. 40 We sought to modify this technique using antibody staining. First, we used beads of specific sizes to set flow gates within which we expect to find lysosomes and other vesicles. As lysosomes are reported to be found within a 0.2-1.2 µM range, we used beads of 0.2-1.25 µM size (Fig 5A) to set voltages for forward and side scatter photo multiplier tubes (PMTs). At this size, we were forced to reduce the exclusion threshold on the cytometer; therefore, we wanted to determine whether our measurements are affected by debris that were previously screened out under the previous threshold. We tested unfiltered sucrose (Fig 5B), 0.22 µM filtered sucrose (Fig 5C), and sucrose that was spun down by ultracentrifugation for 2 hours at 40,000 x g (Fig 5D) for a fixed period and 156 measured background events over a defined time period. We found that most debris was removed via filtration, and centrifugation removed an even greater amount of debris. Each of these measurements were taken over a five minute interval – about five times longer than individual samples were subsequently run for – and therefore represent what would be a much smaller representation of the background observed in any specific sample. Additionally, none of these background events displayed auto- fluorescence in any of the channels we measured. As such, this population represents a minimal number of aberrant events that can only serve to dilute the target vesicles of interest. We expect that once lamp-1+ vesicles are sorted out of the population this effect will disappear. We next sought to determine whether we could remove larger components, such as nuclei and mitochondria, from cell lysates by centrifugation at 5,000 x g for 10 minutes, as has been reported by others. 20 Using the same size exclusion reported earlier (Fig 6A,B), we compared size and complexity (forward scatter and side scatter) of: 1) vesicles that were directly lysed (Fig 6C), 2) the supernatant of lysates spun down at 5,000 x g for 10 minutes (Fig 6D), 3) and the pellets of this centrifugation step (Fig 6E). Direct measurement of the lysate revealed both large and small populations of organelles. We found that the larger cellular components were successfully separated by centrifugation, as evidenced by the concentration of large vesicles in the nuclear pellet, and their relative absence in the post-nuclear supernatant (PNS). In subsequent 157 experiments, we used the PNS to measure lysosome localization. By doing so, we ensured cells that were not lysed during the sonication step were removed from the analysis. We next sought to determine whether we could measure lysosomes via SOFA by staining with antibodies against lamp-1. When we directly stained the cells with lamp-1 and measured intact cells via flow cytometry, we found that all cells stain positively for the lysosomal marker, as expected (Fig 7A). Staining for SOFA was performed on intact cells. After staining, cells were lysed via sonication. To determine what amount of sonication is necessary to liberate subcellular organelles, we performed a series of increasing rounds of sonication on cells to determine the optimum number of cycles that would lyse the cells without compromising the antibodies themselves (Fig 7B). We found that we could successfully stain subcellular organelles with antibodies and visualize them via flow cytometry. We also observed an increased concentration of lamp-1 positive vesicles after removal of the nuclear pellet. Finally, we observed that increasing the number of sonication cycles did not improve the yield of lysosomes, but instead reduced the number of recovered lysosomes, presumably due to the sonication disrupting the lysosomes themselves over extended cycles. Having established a protocol for antibody staining and measurement of subcellular organelles, we sought to determine whether we could replicate these results 158 with antibodies against various chemokines and cytokines. To do this, we stained cells with antibodies against gro-α (Fig 8A), lamp-1 (Fig 8B), CCL20 (Fig 8C), and TNF-α (Fig 8D). We found that we could detect positive antibody staining for all antibodies tested. The strongest staining was exhibited by lamp-1 and TNF-α, with significant staining of gro-α manifesting at higher concentrations. While we did observe a small amount of CCL20 staining, this small increase was insufficient, even at higher concentrations, to allow us to quantify CCL20 in vesicles using multiple antibody stains. The isotype controls displayed for each target correspond to the medium antibody concentration used. Higher isotype concentrations did not increase background staining within the concentrations we measured. Many chemokines and cytokines colocalize in vivo We next sought to determine whether chemokines and cytokines that are stored in the bronchial epithelium are stored in the same vesicles with CCL20. Because CCL20 staining in human cells was too dim to measure colocalization, we stained mouse lungs with antibodies against multiple chemokines/cytokines to determine whether they colocalize. We found that in mouse lungs CCL20 colocalized with lamp-1 (Fig 9A-C), similar to our observations in human cells. Additionally, we observed significant, though not complete, colocalization of CCL20 with IL-6 (Fig 9D-F) and gro-α (Fig 9G-I), suggesting that many, but not all, intracellular lysosomes containing CCL20 also contain IL-6 and gro-α. 159 Degranulation of bronchial epithelial cells is induced by a variety of allergens Having observed that multiple chemokines and cytokines localize to the same secretory lysosomes, we hypothesized that this mechanism for chemokine/cytokine release may be common to allergens other than HDM. To test whether other allergens are able to induce secretory lysosome release, we performed a degranulation assay in the presence of a panel of allergens, including cockroach frass, LPS, ragweed pollen, and maple pollen. We found that the number of cells that displayed significant lamp-1 uptake increased with each of the allergens we tested (Fig 10A). In addition, MFI for each of the allergen treatments was increased in comparison to PBS control (Fig 10B). These data indicate that increased degranulation is occurring in these cells. We also found that different allergens produced different levels of degranulation; in particular, cockroach frass induced much more degranulation than other allergens we tested. When we treated the cells with the chloride inhibitor DIDS, we saw a dramatic decrease in degranulation for all treatments. DIDS treatment further reduced cockroach-specific degranulation in addition to baseline decreases (Fig 10C). Additional tests are needed to determine whether DIDS, bafilomycin A, and other signaling pathway inhibitors also reduce this measure of degranulation, and whether this same mechanism is induced by each of the specific allergens we tested. 160 Degranulation of specific vesicles is differentially induced by specific allergens We next sought to determine whether stimulation with various allergens was sufficient to induce release of specific chemokine and cytokine-containing secretory lysosomes using the SOFA technique outlined previously. After treating human bronchial epithelial cells with various allergens, we stained them with to identify lysosomes and specific chemokines. Our earlier observations showed sufficient subcellular staining to distinguish vesicles containing gro-α, TNF-α, and Lamp-1; but not CCL20. Therefore, we selected gro-α and TNF-α as a candidate chemokine/cytokine pair to determine whether they are stored in the same or separate vesicles, with lamp-1 used to identify lysosomes. We then lysed the cells and performed SOFA on the PNS. We observed that levels of gro-α and TNF-α were generally enriched in lamp-1 containing vesicles when compared with lamp-1 deficient vesicles. This lends support to our hypothesis that these chemokines and cytokines are stored in secretory lysosomes (Fig 11). Additionally, we observed increased numbers of gro-α positive, TNF-α negative vesicles when we treated cells with frass, LPS, or maple; suggesting treatment with these allergens induces accumulation of gro-α in these lysosomes (Fig 11A). Additionally, gro-α and TNF-α double-positive vesicles accumulated in the presence of LPS, suggesting stimulation with LPS induces accumulation of both gro-α and TNF-α in lysosomes (Fig 11B). No accumulation of TNF-α was observed in lysosomes that did not contain gro-α (Fig 11D). Finally, we observed depletion of 161 vesicles that contain neither gro-α nor TNF-α when treated with any allergen, suggesting accumulation of these mediators into lysosomes is driven by allergen exposure (Fig 11C). Although this experiment has been repeated, the magnitude of the error bars in A and B suggest a larger n should be collected to further verify these results. Additionally, collection of an increased number of events could also help reduce the uncertainty in the data. There are, however, practical limits to the number of events that can be obtained when collecting a large number of samples. Some co-loading of lysosomes is indicated by the increase in the number of Lamp-1+ vesicles containing both gro-a and TNF-a after treatment with LPS. However in each case we observed an increase in the number of vesicles containing chemokine and cytokine in response to allergen. This may indicate the target proteins are being loaded into vesicles after allergen stimulation. Since there is no change in the Lamp-1- population, this increase is likely to be derived from a non-vesicular source, such as the cytosol. Release of many mediators is induced by a broad spectrum of allergens We treated human bronchial epithelial cells with a panel of allergens, including ambient particulate matter collected from urban Baltimore (AUB); lipopolysaccharide (LPS); which is found on many bacterial cell walls; HDM; cockroach frass; fel d 1, the major cat allergen; ragweed pollen; birch pollen; alder pollen; and maple pollen (Fig 12, summarized in table 2). We found a number of interesting patterns governing release 162 of mediators. For example, HDM induced the greatest release of any allergen we treated cells with for any chemokine or cytokine. Additionally, stimulation with the other insect allergen, cockroach frass, often mirrored HDM stimulation, suggesting they share common pattern recognition receptor stimulation. Also, LPS induced a number of mediators, such as IL-1α, IL-6, IP-10 and TNF-α, suggesting that allergens containing significant amounts of LPS, such as HDM and frass, should induce release of these mediators. Interestingly, the pollen allergens induced less chemokine and cytokine release overall, and in the case of Alder even decreased MIP-1β release. This suggests a bias of early epithelial cells against reactivity to pollen allergens. 163 Discussion Although when we blocked CCL20 alone we saw a small trend of reduced AHR and BAL cellularity, this did not appear to be effective in substantially reducing the airway response to HDM. One explanation for this muted response could be that the anti-CCL20 neutralizing antibody was delivered concurrently with HDM, and the CCL20 was released to the baso-lateral side of the epithelium before the neutralizing antibodies could travel across the epithelium. This co-delivery method was necessary to ensure the mice would not receive multiple i.t. treatments within a short period of time, with insufficient time to recover from the previous anesthetization. Pretreatment of the airways with CCL20 would allow time for penetration of anti-CCL20 prior to HDM-induced CCL20 release, given that we expect baso-lateral release of CCL20. Another option would be to directly measure the effect of blocking CCL20 by measuring recruitment of iDCs into the airways after treating with HDM, with and without CCL20-blocking antibodies. Although this method would more directly test the effect of blocking CCL20, measuring downstream endpoints, such as AHR and airway inflammation, is a better measure for whether blocking CCL20 alone is sufficient to diminish the development of asthma overall. A number of therapies based on inhibition of individual chemokines or cytokines have been pursued recently.41,42 Although promising, many of these therapies have proved inadequate for reducing asthma symptoms. 43 It is likely that targeting multiple 164 chemokine and cytokine pathways simultaneously will be necessary to dramatically impact AHR for some asthmatics. The prospect that multiple mediators might be released under similar cellular mechanisms led us to investigate whether multiple chemokines and cytokines are stored and released from bronchial epithelial cells, and whether chemokines and cytokines other than CCL20 are also stored and released from secretory lysosomes in response to allergen exposure. Using a combination of subcellular flow cytometry, and confocal microscopy, we showed that multiple chemokines and cytokines may reside in distinct vesicles within bronchial epithelial cells. In lung sections visualized by confocal microscopy, a number of vesicles appear to be stained for both CCL20 and gro-a, or CCL20 and IL-6. This staining is not exclusive, suggesting multiple types of these vesicles may to reside within a single cell. This is similar to secretory granules in other cell types, such as in neutrophils that have azurophilic, specific, and gelatinase granules; with individual subsets of granules containing distinct patterns of protein and enzyme storage. We also showed that different allergens could induce release of different combinations of chemokines and cytokines. Our data also suggest that this difference may be due to release of different combinations of vesicles; with some containing gro-α, but not TNF-α induced by HDM; and some containing both gro-α and TNF-α induced by LPS. Thus, release from these vesicles was triggered by allergen exposure, and individual allergens triggered release of specific subsets of lysosomes. As mentioned previously, gro-α and 165 TNF-α were both found at high concentrations within the cell lysates, and these proteins were also both released after HDM treatment. Future studies will determine whether this differential sorting is true across other chemokines and cytokines. Similar measurements of proteins that were found in lower abundance in the cell lysate, such as CCL20, were difficult due to the sensitivity constraints of the assay. Further improvements to the SOFA assay to improve sensitivity will be required to make comparisons between CCL20 and other chemokines and cytokines. Our attempts to improve sensitivity included increasing CCL20 antibody concentration, decreasing the number of sonication cycles to reduce dissociative stress, and employing a primary- secondary antibody system to enhance staining. A future approach may include creating a stabile-transfected cell line expressing CCL20. Despite the sensitivity limitations of SOFA, we demonstrated that staining and measurement of subcellular organelles, such as lysosomes, could be accomplished by flow cytometry. This is significant, because previous efforts at measurement of subcellular organelles by flow cytometry required either transfection of the cells with fluorescently labeled tags for the protein of interest, or relied on staining with less- specific dyes, such as MitoTracker. 44,45 Here, we demonstrate a method for accomplishing subcellular flow by antibody staining, similar to traditional flow cytometry. This approach offers several advantages to using transfected plasmids. 166 Antibody staining allows for greater flexibility and a broader array of targets, as plasmids need not be prepared for each target and transfected into the cell line. Also, cells from existing animal models can be interrogated directly. Additionally, as we showed here, staining with multiple antibodies allows us to identify colocalization within a subcellular organelle. This colocalization is superior to superposition observed in confocal microscopy. Superposition only suggests colocalization and must be validated to ensure the staining does not represent protein in the cytosol. In flow analysis, the vesicles are separated from their surrounding cytosol, eliminating this confounding factor. Indeed, SOFA could be useful as a general tool to validate colocalization observed by microscopy. Another benefit of this method is that it is quantitative. While methods such as immuno-TEM are sufficient to show colocalization of two proteins within the same organelle, we demonstrate that SOFA has the added benefit of measuring populations of organelles, and changes in these populations after stimulation. We also noted some significant limitations to this approach. First, we found that measurement of subcellular organelles requires increased sensitivity, and our efforts at SOFA were unsuccessful on older, less sensitive machines. This is similar to what was reported by researchers attempting SOFA with fluorescently tagged proteins. Second, we noted that increased sensitivity (and a decreased threshold for events on the 167 instrument) required increased purity of the sucrose buffer in which the cell lysates were suspended. Most fine debris could be removed by filtering the sucrose buffer through a 0.22 um filter. We found that additional debris could be eliminated by centrifuging the buffer at 40,000 x g for 2+ hours. This could be useful for identifying organelles of low abundance. Since lysosomes are plentiful within the cell, we only required filtration for debris removal. The increased sensitivity requirement for SOFA is due to the smaller size of the target. When measuring an intact cell, the signal returns the sum fluorescence across a larger volume than when measuring subcellular features. Therefore, SOFA necessitates fluorescent probes with greater sensitivity, or a target molecule that is in high abundance. In microscopy, increased sensitivity can be easily accomplished by increasing the dwell time of the laser for each pixel, but for flow cytometry this is impossible, as the lysosome passes through the flow cell rapidly and does not allow for increased dwell time. Thus, we found a practical limit for SOFA that impeded our attempts to measure less abundant molecules, such as CCL20. As such, we were unable to determine whether vesicles containing CCL20 also contain other mediators. Another approach to this problem is addressed through the Amnis ImageStream instrument, which takes a continuous exposure of the event as it passes through the flow cell. This has the practical effect of increasing exposure time, similar to that seen on a confocal microscope. Unfortunately, one of the main limiting factors of the ImageStream is the 168 number of events that can be acquires, which is orders of magnitude lower than traditional flow cytometry. Given that we were already running somewhat high numbers of events for each sample, this practical limitation reduces the ImageStream’s effectiveness due to a smaller sample size, even though it may allow detection of otherwise low-abundance proteins such as CCL20. A major strength of using a flow cytometry-based technique is that it was not necessary to separate the cell lysate by the time-consuming process of density- dependent centrifugation, because we had a specific marker to identify our organelle of interest. In this case we used lamp-1 to identify lysosomes, although this method could be applied to any subcellular organelle. A drawback of lysing the cell is that fragments of the cell membrane are also likely to form micelles and appear as a small number of events during analysis. These micelles are likely to be small, but may still affect our analysis. Our data suggest that after allergen exposure there is an increase in Lamp-1 that localizes to the cell surface. Specific cell surface stains could help remove this population in future experiments. As previously discussed, multiple chemokines and cytokines are released into the BAL of asthmatics after exposure to allergen. Our finding that many allergens could induce the release of multiple chemokines and cytokines from bronchial epithelial cells supports this previous observation, and suggests that a primary source for many 169 mediators of inflammation is the bronchial epithelium. However, we also observed that while some mediators are stored within the same cell compartment, not all the mediators released upon allergen stimulation can be traced back to a single vesicle, or even vesicular storage at all in the case of IL-1α. Thus, the release of mediators governed by many different release mechanisms suggests that multiple signaling pathways contribute to the diverse set of mediators released after allergen challenge. For IL-8, we observed significant levels of nuclear staining in addition to cytoplasmic staining. At present, it is unclear what role IL-8 plays in the nucleus. Additionally, other groups have reported IL-8 storage on the cell surface. We did not observe significant staining of surface IL-8 or CCL20. In contrast, surface staining of CCL20 by flow cytometry reveals some CCL20 localizes to the cell surface, although at significantly lower levels than intracellular staining. It is likely that surface concentrations of both CCL20 and IL-8 are significantly lower and more diffuse than intracellular vesicular concentrations, such that they are not visible by immunofluorescence microscopy. As a potential treatment option, there are several advantages to targeting the mechanism for chemokine and cytokine release outlined above, as opposed to targeting a single signaling molecule. We found that targeting CCL20 alone had limited biological activity, possibly due to the many other inflammatory mediators induced by 170 HDM. Our studies suggest an alternate strategy of targeting the mechanism for release of multiple mediators. Broadly speaking, this would reduce the overall signal produced by the bronchial epithelium in response to HDM. Blocking the release pathway initiated by specific allergens, instead of the chemokine itself, may allow us to target activation in response to HDM (or other allergens) specifically, as opposed to more general immune components. This would allow attenuation of allergen-specific immunity, without the unwanted effect of compromising beneficial immune responses to unwelcome pathogens. For example, long-term treatment with glucocorticoids leads to osteoporosis, and increases susceptibility to infections as a result of its immunosuppressive effects.46,47 Finally, our finding that this pathway of secretory lysosome release is common to multiple allergens suggests targeting their release may be potent against multiple types of allergens. 171 References: 1. Broide DH, Lotz M, Cuomo AJ, et al. Cytokines in symptomatic asthma airways. Journal of Allergy and Clinical Immunology . 1992;89(5):958–967. 2. Fitzpatrick AM, Higgins M, Holguin F, Brown LAS, Teague WG. The molecular phenotype of severe asthma in children. Journal of Allergy and Clinical Immunology . 2010;125(4):851–857.e18. 3. Chung KF, Barnes PJ. Cytokines in Asthma. Thorax . 1999;54(9):825–857. 4. Stanley AC, Lacy P. Pathways for Cytokine Secretion. Physiology . 2010;25(4):218–229. 5. Kouzaki H, Iijima K, Kobayashi T, O’Grady SM, Kita H. The Danger Signal, Extracellular ATP, Is a Sensor for an Airborne Allergen and Triggers IL-33 Release and Innate Th2-Type Responses. J Immunol . 2011;186(7):4375–4387. 6. Gibbs BF, Wierecky J, Welker P, et al. Human skin mast cells rapidly release preformed and newly generated TNF-α and IL-8 following stimulation with anti-IgE and other secretagogues. Experimental Dermatology . 2001;10(5):312–320. 7. Øynebråten I, Bakke O, Brandtzaeg P, Johansen F-E, Haraldsen G. Rapid chemokine secretion from endothelial cells originates from 2 distinct compartments. Blood . 2004;104(2):314–320. 8. Wright DG, Gallin JI. Secretory Responses of Human Neutrophils: Exocytosis of Specific (Secondary) Granules by Human Neutrophils During Adherence in Vitro and During Exudation in Vivo. J Immunol . 1979;123(1):285–294. 9. Fournier BM, Parkos CA. The role of neutrophils during intestinal inflammation. Mucosal Immunology . 2012;5(4):354–366. 10. Egesten A, Breton-Gorius J, Guichard J, Gullberg U, Olsson I. The heterogeneity of azurophil granules in neutrophil promyelocytes: immunogold localization of myeloperoxidase, cathepsin G, elastase, proteinase 3, and bactericidal/permeability increasing protein. Blood . 1994;83(10):2985–2994. 11. Baggiolini M, de Duve C, Masson PL, Heremans JF. ASSOCIATION OF LACTOFERRIN WITH SPECIFIC GRANULES IN RABBIT HETEROPHIL LEUKOCYTES. J Exp Med . 1970;131(3):559–570. 12. Murphy G, Reynolds JJ, Bretz U, Baggiolini M. Collagenase is a component of the specific granules of human neutrophil leucocytes. Biochem J . 1977;162(1):195–197. 13. Kjeldsen L, Bainton DF, Sengelov H, Borregaard N. Structural and functional heterogeneity among peroxidase-negative granules in human neutrophils: identification of a distinct gelatinase- containing granule subset by combined immunocytochemistry and subcellular fractionation. Blood . 1993;82(10):3183–3191. 14. Faurschou M, Borregaard N. Neutrophil granules and secretory vesicles in inflammation. Microbes and Infection . 2003;5(14):1317–1327. 15. Bentwood BJ, Henson PM. The sequential release of granule constitutents from human neutrophils. J Immunol . 1980;124(2):855–862. 172 16. Goldstein IM, Hoffstein ST, Weissmann G. Mechanisms of lysosomal enzyme release from human polymorphonuclear leukocytes. Effects of phorbol myristate acetate. J Cell Biol . 1975;66(3):647–652. 17. Klempner MS, Dinarello CA, Gallin JI. Human leukocytic pyrogen induces release of specific granule contents from human neutrophils. J Clin Invest . 1978;61(5):1330–1336. 18. Ganz T. Extracellular release of antimicrobial defensins by human polymorphonuclear leukocytes. Infect. Immun. 1987;55(3):568–571. 19. Øynebråten I, Barois N, Hagelsteen K, et al. Characterization of a Novel Chemokine-Containing Storage Granule in Endothelial Cells: Evidence for Preferential Exocytosis Mediated by Protein Kinase A and Diacylglycerol. J Immunol . 2005;175(8):5358–5369. 20. Pasquali C, Fialka I, Huber LA. Subcellular fractionation, electromigration analysis and mapping of organelles. Journal of Chromatography B: Biomedical Sciences and Applications . 1999;722(1–2):89–102. 21. Mey JD, Lambert AM, Bajer AS, Moeremans M, Brabander MD. Visualization of microtubules in interphase and mitotic plant cells of Haemanthus endosperm with the immuno-gold staining method. PNAS . 1982;79(6):1898–1902. 22. Stirling JW. Immuno- and affinity probes for electron microscopy: a review of labeling and preparation techniques. J Histochem Cytochem . 1990;38(2):145–157. 23. Brakke MK. Density Gradient Centrifugation: A New Separation Technique1. J. Am. Chem. Soc. 1951;73(4):1847–1848. 24. Zahler S, Kowalski C, Brosig A, et al. The function of neutrophils isolated by a magnetic antibody cell separation technique is not altered in comparison to a density gradient centrifugation method. Journal of Immunological Methods . 1997;200(1–2):173– 179. 25. Shotton DM. Confocal scanning optical microscopy and its applications for biological specimens. J Cell Sci . 1989;94(2):175–206. 26. Bolte S, Cordelières FP. A guided tour into subcellular colocalization analysis in light microscopy. Journal of Microscopy . 2006;224(3):213–232. 27. Brabec M, Schober D, Wagner E, et al. Opening of Size-Selective Pores in Endosomes during Human Rhinovirus Serotype 2 In Vivo Uncoating Monitored by Single-Organelle Flow Analysis. J. Virol. 2005;79(2):1008–1016. 28. Platts-Mills TAE, Woodfolk JA. Allergens and their role in the allergic immune response. Immunological Reviews . 2011;242(1):51–68. 29. Page K, Strunk VS, Hershenson MB. Cockroach proteases increase IL-8 expression in human bronchial epithelial cells via activation of protease-activated receptor (PAR)–2 and extracellular-signal-regulated kinase. Journal of Allergy and Clinical Immunology . 2003;112(6):1112–1118. 173 30. Elias JA, Homer RJ, Hamid Q, Lee CG. Chitinases and chitinase-like proteins in TH2 inflammation and asthma. Journal of Allergy and Clinical Immunology . 2005;116(3):497–500. 31. Valerio C, Murray P, Arlian L, Slater J. Bacterial 16S ribosomal DNA in house dust mite cultures. Journal of Allergy and Clinical Immunology . 2005;116(6):1296–1300. 32. Paton JB. Pollen and Pollen Enzymes. American Journal of Botany . 1921;8(10):471– 501. 33. Huss K, Adkinson Jr NF, Eggleston PA, et al. House dust mite and cockroach exposure are strong risk factors for positive allergy skin test responses in the Childhood Asthma Management Program*. Journal of Allergy and Clinical Immunology . 2001;107(1):48–54. 34. Nathan AT, Peterson EA, Chakir J, Wills-Karp M. Innate immune responses of airway epithelium to house dust mite are mediated through beta-glucan-dependent pathways. J. Allergy Clin. Immunol. 2009;123(3):612–618. 35. Lundy SK, Lira SA, Smit JJ, et al. Attenuation of Allergen-Induced Responses in CCR6–/– Mice Is Dependent Upon Altered Pulmonary T Lymphocyte Activation. J Immunol . 2005;174(4):2054–2060. 36. Schon-Hegrad MA, Oliver J, McMenamin PG, Holt PG. Studies on the Density, Distribution, and Surface Phenotype of Intraepithelial Class II Major Histocompatibility Complex Antigen (Ia)-Bearing Dendritic Cells (DC) in the Conducting Airways. J Exp Med . 1991;173(6):1345–1356. 37. Kelly C, Ward C, Stenton CS, et al. Number and activity of inflammatory cells in bronchoalveolar lavage fluid in asthma and their relation to airway responsiveness. Thorax . 1988;43(9):684–692. 38. Marshall LJ, Ramdin LSP, Brooks T, DPhil PC, Shute JK. Plasminogen Activator Inhibitor-1 Supports IL-8-Mediated Neutrophil Transendothelial Migration by Inhibition of the Constitutive Shedding of Endothelial IL-8/Heparan Sulfate/Syndecan-1 Complexes. J Immunol . 2003;171(4):2057–2065. 39. Bitensky L. Lysosomes and the connective tissue diseases. J Clin Pathol Suppl (R Coll Pathol) . 1978;12:105–116. 40. Böck G, Steinlein P, Huber LA. Cell biologists sort things out: Analysis and purification of intracellular organelles by flow cytometry. Trends in Cell Biology . 1997;7(12):499–503. 41. Desai D, Brightling C. Cytokine and anti-cytokine therapy in asthma: ready for the clinic? Clinical & Experimental Immunology . 2009;158(1):10–19. 42. Hansbro PM, Kaiko GE, Foster PS. Cytokine/anti-cytokine therapy – novel treatments for asthma? British Journal of Pharmacology . 2011;163(1):81–95. 43. Walsh GM. An update on emerging drugs for asthma. Expert Opin Emerg Drugs . 2012;17(1):37–42. 174 44. Dhandayuthapani S, Via L e., Thomas C a., et al. Green fluorescent protein as a marker for gene expression and cell biology of mycobacterial interactions with macrophages. Molecular Microbiology . 1995;17(5):901–912. 45. Salvioli S, Dobrucki J, Moretti L, et al. Mitochondrial heterogeneity during staurosporine-induced apoptosis in HL60 cells: Analysis at the single cell and single organelle level. Cytometry . 2000;40(3):189–197. 46. Goldstein MF, Fallon J, Harning R. Chronic Glucocorticoid Therapy-Induced Osteoporosis in Patients With Obstructive Lung Disease*. CHEST . 1999;116(6):1733– 1749. 47. Klein NC, Go CH, Cunha BA. Infections associated with steroid use. Infect. Dis. Clin. North Am. 2001;15(2):423–432, viii. 175 Tables Table 1 – Chemokines and cytokines measured in bronchial epithelial cell lysate s. Mediator Lysate Stores HDM release Mediator Lysate Stores HDM release CCL20 ++ ++ IL-8 +++ +++ EGF + IL-9 Eotaxin IL-10 FGF-2 ++ ++ IL-12p40 FLT-3L IL-12p70 Fractalkine ++ IL-13 G-CSF ++ IL-15 GM-CSF ++ IP-10 ++ + GRO ++++ ++++ MCP-1 + IFN-a 2 MCP-3 IFN-g MDC IL-1Ra MIP-1a ++ IL-1a +++ ++ MIP-1b ++ + IL-1b SCD40L IL-2 sIL-2Ra IL-3 TGF-a IL-4 TNF-a ++ + IL-5 TNF-b IL-6 +++ ++++ VEGF ++ IL-7 (Number of “+” equal to log stores or half-log released chemokine.) 176 Table 2 – Chemokine/cytokine release by various allergens AUB LPS HDM Frass Fel D 1 Ragweed Birch Alder Maple FGF-2 +++ + GM-CSF + + + + Gro-a +++ - IL-1a + ++ + IL-6 + + ++++ + + IL-8 ++ + + IP-10 + ++ + MIP-1α + + +++ ++ MIP-1β + + - Rantes + TNF-α ++ + + Increased release compared to PBS treatment: + 25% increase ++ 50% increase +++ 75% increase ++++ 100% increase - 25% decrease 177 A Day: 0 21 24 challenge (C, E). Airway hyper-responsiveness measured by APTI 72 hours after nal HDM Sacri ce exposure are given in (B) and (C), and BAL cellularity for the same time point is given in (D) Sensitization Challenge and (E). Data are represented as mean +/- SEM. *p<0.05 compared to control PBS treatment. HDM i.t. HDM i.t. APTI BAL B 1500 C 2500 * * 2000 1000 O x sec.) O x sec.) 1500 2 2 1000 500 500 APTI (cm H APTI (cm H 0 0 PBS HDM aCCL20 PBS HDM aCCL20 +HDM +HDM Ab given at sensitization Ab given at challenge D 200000 * E 200000 150000 150000 * 100000 100000 Total BAL cells 50000 50000 Total BAL cells 0 0 PBS HDM aCCL20 PBS HDM aCCL20 +HDM +HDM Ab given at sensitization Ab given at challenge Figure 1 – CCL20 neutralization has little e ect on airway responsiveness, and airway in amma- tion after HDM sensitization and challenge. (A) Experimental outline; mice were treated with HDM or PBS control intra trachea on days 0 and 21. Mice were sacri ced 72 hours after nal treatment. Neutralizing antibody to CCL20 was given at either sensitization (B, D) or n=8. *p<0.05, **p<0.01, ***p<0.001 compared to control HDM treatment. A Gro-a IL-8 IL-6 IL-1a FGF-2 IP-10 TNF-a G-CSF GM-CSF VEGF Fractalkine MIP-1a CCL20 MIP-1b MCP-1 EGF 0 1 2 3 4 5 Log protein in lysate (pg/mL) B IL-6 *** Gro-a *** IL-8 *** IL-1a *** CCL20 * FGF-2 ** TNF-a *** IP-10 *** PBS MIP-1b * HDM 0 200 400 600 800 Protein in supernatant (pg/mL) Figure 2 – Many chemokines and cytokines that are stored in bronchial epithelial cells are released in response to HDM. (A) Untreated cell lysates were measured by multiplex protein quanti cation assay for the presence of a panel chemokines and cytokines. (B) Supernatants of cells treated with HDM or PBS control for 2 hours were measured by multiplex assay for the presence of a panel of chemokines and cytokines. Proteins measured are listed in table 1; proteins below the limit of detection are not depicted. Data are represented as mean +/- SEM. n=3. 750 500 250 200 CCL20 150 Gro-a IL-17f 100 pg/mL TSLP 50 0 1 2 3 4 5 6 7 8 9 10 11 12 Fraction number Figure 3 – Chemokines and cytokines are stored in multiple cellular compartments. HBE cells were separated on a Percoll gradient. After centrifugation, fractions were removed from the top of the gradient and measured for various chemokines and cytokines. GM-CSF IL-8 DAPI DAPI Gro-α CCL20 DAPI DAPI Figure 4 – Chemokines and cytokines in lung epithelial cells localize to distinct cellular compart- ments. Unstimulated HBE cells stained with di erent antibodies, and imaged via confocal microscopy. Antibody staining appears green, and Nuclei are stained with DAPI and appear blue. A B Un ltered sucrose 100 Bead size 10 5 0.2 μM 1.2 μM 80 10 4 60 SSC 10 3 % of Max % of Ma x 40 2 20 10 0 0 0 10 2 10 3 10 4 10 5 0 10 2 10 3 10 4 10 5 FSC FSC C 0.22 μM ltered sucrose D Centrifuged sucrose 10 5 10 5 10 4 10 4 SSC 10 3 SSC 10 3 10 2 10 2 0 0 0 10 2 10 3 10 4 10 5 0 10 2 10 3 10 4 10 5 FSC FSC Figure 5 – Sucrose puri cation is necessary to remove debris prior to single organelle ow analy- sis. (A) Size beads were used to set appropriate gates for forward and side scatter on a log scale. Sucrose debris was then assessed by measuring events for 60 seconds at a constant ow rate. Debris is represented for direct, un ltered sucrose (B), sucrose that was ltered through 0.22 µM pores (C), and sucrose centrifuged at 40,000 x g for 2 hours (D). A 10 5 B 10 5 10 4 10 4 SSC 3 SSC 3 SSC-A10 SSC-A 10 10 2 10 2 0 0 0 10 2 10 3 10 4 10 5 0 10 2 10 3 10 4 10 5 FSC FSC C 10 5 10 4 SSC 10 3 Whole lysate Whole 10 2 0 0 10 2 10 3 10 4 10 5 FSC D 10 5 E 10 5 10 4 10 4 SSC 10 3 SSC 10 3 PNS 10 2 Nuclear pellet 10 2 0 0 0 10 2 10 3 10 4 10 5 0 10 2 10 3 10 4 10 5 FSC FSC Figure 6 – Isolated post-nuclear supernatant is distinct from nuclear pellet. Size exclusion gates were set for forward and side scatter using bead sizes of 0.2 µM (A) and 1.25 µM (B). Density plots for forward and side scatter are depicted for HBE cells that were lysed (C) and then centrifuged at 5,000 x g for 10 minutes prior to separation of the post-nuclear supernatant (D) and the nuclear pellet (E). Lysate Nuclear pellet 100 Lamp-1 PNS A Isotype B 15 100 80 60 10 % of Ma x 40 20 5 Lamp-1 (% vesicles) 0 0 10 2 10 3 10 4 10 5 Lamp-1 0 Iso 5 10 15 Cycles Figure 7 – Vesicles containing Lamp-1 are primarily located in the post-nuclear supernatant. After staining whole cells with Lamp-1, cells were lysed via sonication for 5, 10, or 15 cycles, and then centrifuged at 5,000 x g for 10 minutes prior to separation of the post-nuclear supernatant and the nuclear pellet. Whole cells (A) or vesicles (B) were analyzed by ow cytometry for the presence of Lamp-1. A 100 High B 100 High Med Med Low Low 80 80 Isotype Isotype 60 60 % of Max % of Max % of max 40 % of max 40 20 20 0 0 0 10 2 10 3 10 4 10 5 0 10 2 10 3 10 4 10 5 Gro-α Lamp-1 C 100 High D 100 High Med Med Low Low 80 80 Isotype Isotype 60 60 % of max 40 % of max 40 % of Max % of Max 20 20 0 0 0 10 2 10 3 10 4 10 5 0 10 2 10 3 10 4 10 5 CCL20 TNF-α Figure 8 – Vesicles containing chemokines and cytokines can be measured by SOFA. Unstimu- lated HBE cells stained with varying antibody concentrations for di erent chemokines and cytokines were lysed and PNS was recovered after centrifugation. Histogram plots for Gro-α (A), Lamp-1 (B), CCL20 (C), and TNF-α (D) are shown. A B C DAPI CCL20 Lamp-1 CCL20 Lamp-1 D E F DAPI IL-6 CCL20 IL-6 CCL20 G H I DAPI IL-33 CCL20 IL-33 CCL20 Figure 9 – Di erent vesicles in lung epithelial cells contain di erent subsets of chemokines and cytokines. Unstimulated mouse lung sections stained with di erent antibody combinations. (A-C) Lungs stained with Lamp-1 (A) and CCL20 (B) with a merged pseudo-color image (C). (D-F) Lungs stained with CCL20 (D) and IL-6 (E) with a merged pseudo-color image (F). (G-I) Lungs stained with CCL20 (G) and IL-33 (H) with a merged pseudo-color image (I). Nuclei are stained with DAPI and appear blue (C, F, I). *** ** A 6 B 1000 *** ** ** 800 * 4 600 ** * 400 2 Lamp-1 (MFI) 200 Lamp-1 (% live cells) 0 0 PBS Fra LPS Rag Mpl PBS Fra LPS Rag Mpl C 20 PBS HDM 15 Frass 10 5 Lamp-1 (% live cells) 0 Ctrl DIDS Figure 10 – Various allergens induce rapid degranulation of HBE cells. Degranulation measured by lamp-1 uptake after 30 minutes of culture with uorescent lamp-1 and a panel of allergens. Data represented as percentage of positive cells (A), or mean uorescence intensity of all cells (B). (C) Cells treated with chloride transporter inhibitor DIDS or vehicle control for 1 hour were treated with HDM, frass, or PBS control for 30 min. in the presence of uorescently- labeled lamp-1 antibodies. PBS = phosphate bu ered saline control, HDM = house dust mite, Fra = cockroach frass, LPS = lipopolysaccharide, Rag = ragweed pollen, Mpl = maple pollen. Data are represented as mean +/- SEM. *p<0.05, **p<0.01, ***p<0.001 compared to control HDM treatment n=3. ** *** ** * ** * ** * * ** *** ** ** *** ** ** * * * ** *** * *** * * * * * * * *** * * Chapter 5. Summary, Discussion, and Conclusions Summary The aim of this dissertation is to determine the mechanisms by which bronchial epithelial cells respond to HDM and release inflammatory chemokines and cytokines. We have demonstrated that: 1. CCL20 is stored in secretory lysosomes under homeostatic conditions. a. Multiple other mediators are stored in bronchial epithelial cells under homeostatic conditions. Some, such as Gro-α and IL-33, are stored in secretory lysosomes, and may even be stored in the same secretory lysosomes as CCL20. b. Other mediators, such as IL-1α and TSLP, are stored in the cytoplasm and are released by a different mechanism than secretory lysosome release. 2. HDM induces rapid secretory lysosome release, including the release of CCL20. a. Secretory lysosome release requires functional V-ATPase, and CLC-7. b. Multiple other allergens induce rapid secretory lysosome release from bronchial epithelial cells. 3. Regulation of rapid CCL20 release through secretory lysosomes is a complex process driven by Raf1 signaling and the generation of reactive oxygen species. 194 Discussion CCL20 is well-suited to the lysosomal environment Recruitment of immature dendritic cells (iDCs) into the bronchial mucosa has been well documented in animals and humans in response to allergen. 1,2 This iDC recruitment is a rapid process. In patients challenged with HDM, an increase in the number of DCs in lung biopsies was observed within 4-5 hours of allergen exposure. 3 Central to early DC recruitment, CCL20 is the only known ligand for CCR6, a receptor preferentially expressed on immature DCs. 4 In humans, both CCL20 release and increased CCR6 + DCs in the lungs of asthmatics have been documented. 5 Until now, little has been known about the mechanisms by which CCL20 is released from the bronchial epithelium in response to allergen. We have shown that CCL20 is rapidly released from the bronchial epithelium within minutes of allergen exposure, and that an important pathway leading to release includes storage of CCL20 in secretory lysosomes within bronchial epithelial cells. This storage and release mechanism has important implications for the function of CCL20, as well as potential insights into the storage and release of other chemokines and cytokines from the bronchial epithelium. Indeed, our data suggest that nearly half of the CCL20 release by bronchial epithelial cells within the first 24 hours that can be attributed to HDM is released within the first two hours of exposure. 195 Although lysosomes are often studied for their degradative effect on many proteins, some proteins are actively stored in lysosomes. 6 These proteins are often better suited to the caustic environment of the lysosome. 7 Some proteases, such as cathepsins, are resistant to degradation by other lysosomal proteins, allowing them to remain within the lysosomal environment. 8 Other proteins evade lysosomal degradation by binding to the heparin-sulfate proteoglycans (HSPGs) that make up what is known as an electron-dense core in lysosomes (so-named because in electron micrographs these areas within the lysosome appear dark). Proteins residing in this HSPG-rich environment within the electron-dense core are not accessible to degradative lysosomal enzymes, and therefore they escape intact.9 Many chemokines and cytokines may potentially utilize this strategy to evade lysosomal degradation. Binding motifs to heparin-sulfate have been identified in IL-8, CCL20, and many other chemokines and cytokines. 10–12 Several chemokines have been shown to bind to specific HSPGs in vivo, such as IL-8 binding to syndecan-1. 13 This binding interaction is targeted at the heparin- sulfate side chains, not the protein stock, for both IL-8 and many other chemokines.14 Syndecans have generally been shown to localize to the cell surface, however syndecan- 1 has also been identified within the lysosomes of breast carcinoma cells – presenting the possibility that this mechanism of intracellular syndecan storage may be present in other cell types as well. A more thorough investigation into the cellular localization of syndecans may reveal a trafficking pathway that leads through the lysosomal 196 compartment. Taken together, these studies suggest a mechanism by which chemokines and cytokines may evade degradation by binding to HSPGs located within lysosomes. In the absence of intracellular syndecan-1, this binding may still play a role in chemokine release. As chemokines are released from bronchial epithelial cells, they are exposed to syndecans on the cell surface, which may sequester them after release until these HSPGs are cleaved themselves. Previous studies involving CCL20 indicate that it is not only tolerant of the lysosomal conditions, but that it displays greater functionality under conditions normally found only within lysosomes, such as increased acidity and high chloride concentrations. For example, under neutral pH CCL20 exists in a dimeric form, but acidic conditions cause this dimer to dissociate into its component monomers. These monomers display increased antimicrobial activity – potentially because CCL20 antimicrobial activity is dependent on insertion of its C-terminal domain into the target microbial cell membrane. This C-terminal domain is hidden in dimer form, thereby reducing antimicrobial activity. Dissociation of the CCL20 dimer reveals the two C- terminal domains. In this way, CCL20 antimicrobial activity is similar to that of β- defensin. 4,15 Dissociation of the dimer also increases the number of CCL20 molecules available for chemotaxis, which is thought to be mediated by the N-terminal domain of the protein. 16 Although we would not expect CCL20 to induce chemotaxis or attack invading microbes from within the lysosome, the protein itself is clearly well-suited to 197 an environment in which many other proteins denature (due to the low pH) or are otherwise inactivated (due to exposure to proteases). In addition to low pH, lysosomes also contain elevated levels of chloride ions. Elevated chloride concentrations have also been shown to enhance CCL20 antimicrobial activity.17 Some of the same properties that make CCL20 resistant to lysosomal degradation also make it well-suited to the conditions found in asthmatic airways. For example, asthmatic airways display dramatically increased acidity, in the pH range of 5.2. 18 Under these conditions, CCL20 would exist as a monomer and exhibit increased chemotactic activity. 15 As discussed in previous chapters, CCL20 release is induced by a number of mechanisms that are not necessarily linked to airway pH and as such we expect it will induce chemotaxis independently of decreased airway pH. However, as asthma develops this may lead to a positive feedback mechanism in which, as a T H2 response develops CCL20 chemotaxis is enhanced, leading to increased recruitment of CCR6-expressing cells such as iDCs, TH17 cells, and B-cells. Thus, although most proteins would be expected to denature and be readily degraded within lysosomes, CCL20 is highly suited to persistent storage in the acidic lysosomal environment, protected from degradation by binding to HSPGs in the electron dense core. Given the structural similarities between CCL20 and many other chemokines – such as IL-8, MIG, and IP-10 – we expect many of these may also be well- suited to residence within the lysosomal environment. 4 Most of the chemokines and 198 cytokines we saw released from bronchial epithelial cells after HDM exposure display heparin-binding motifs (including IL-1α, IL-1β, IL-6, IL-8, IP-10, FGF-2, and Gro-α; but not TNF-α). 10,19–21 Some of these proteins were not found in lysosomes in our hands, such as IL-1α, but would still be expected to bind extracellular sulfated heparin upon release from the cell, contributing to their role in chemotaxis. Others may reside either within lysosomal stores or in other intracellular vesicles. Future experiments are needed to determine whether multiple types of intracellular vesicles are used to export different chemokines and cytokines, or if secretory lysosomes are the sole intracellular storage vesicle for chemokines and cytokines. As a chemokine, CCL20 functions both as an antimicrobial peptide and as chemotactic factor for iDCs and T H17 T-cells. These are functionally separate events, as antimicrobial activity relies on the C-terminal domain and chemotaxis relies on the N- terminal domain. 16 Thus, our finding that CCL20 is rapidly released via degranulation suggests a mechanism that allows the cell to respond to danger by simultaneously attacking a potential intruding microbe and initiating an adaptive immune response. These dual activities have also been demonstrated for a number of other chemokines in addition to CCL20; many chemokines display strong antimicrobial activity against gram negative bacteria (with an LD 50 in the range of 1 µg/mL), but weaker activity against gram positive bacteria (with an LD 50 in the range of 5 µg/mL). 17 In comparison, DC chemotaxis has been shown to require only 100-300 ng/mL. 22 199 Additional storage and release mechanisms for CCL20 include extracellular stores on syndecan-1 Although much of our work is focused on intracellular mechanisms of chemokine release we also observed evidence of extracellular CCL20 stores. As mentioned earlier, stores of chemokines, such as IL-8, have been shown to be bound to syndecans on the cell surface. It is likely that some of the signaling pathways described above inhibit either or both intracellular and extracellular mechanisms of release. For example, we saw complete inhibition of CCL20 release in the presence of chloride transporter inhibitors or in chloride deficient solution. However, when we specifically targeted lysosome acidification by inhibiting V-ATPase or by knocking down CLC-7, which participates in lysosome acidification, we only saw about a 50% reduction in CCL20 release. This suggests that the presence of chloride affects CCL20 release through interactions other than through the lysosomal release pathway, such as through a separate chloride transporter. Although the specific identity of this chloride transporter remains unknown, chloride and other ions are known to be involved in a number of cell processes that are important for chemokine release. In addition to lysosome formation, release of lysosomes by fusion with the external plasma membrane is also ion-mediated. 23 In neuronal cells, synaptic vesicles have been shown to tether to the plasma membrane in preparation for release. 24 Upon cell depolarization, calcium released into the cytosol 200 induces membrane fusion complexes, called the SNARE complex, to change conformation and fuse the vesicle with the plasma membrane. 25 This change in membrane potential that initiates vesicle fusion is driven by ion changes in the cell. 26 In different cell types, these changes have been shown to be associated with a variety of ions. An example of this with anions is external ATP-mediated vesicle release from medullary adrenal cells, which has been shown to be a chloride dependent process.27 A cation-specific example of this is synaptic vesicle release in frog neurons, which is induced by high external concentrations of potassium – similar to our observation that increased external potassium dramatically increased CCL20 release. 28,29 In bronchial epithelial cells, chloride efflux, or potassium influx, could potentially drive release of docked vesicles by changing the electrical potential within the cell. The specific identity of these transporters remains to be determined; however some interesting candidates have been associated with synaptic vesicle release in neurons. CLC family members, such as CLC-2, have been associated with tethered synaptic vesicles in particular, which may suggest a direct role in vesicle release. 30 Additionally, potassium transporters, such as calcium-gated potassium channels (gKca), have been shown to be in close proximity to the presynaptic membrane where synaptic vesicles are tethered, suggesting they may also play a role in vesicular release. 31 Whichever transporters participate in vesicular release in bronchial epithelial cells, they are likely to do so by inducing rapid changes in voltage or ion concentration to trigger release. 201 In addition to intracellular changes in ion concentration, extracellular changes could also affect release of extracellular chemokine and cytokine stores. As has been noted previously, many chemokines and cytokines are stored on HSPGs via electrostatic interactions. We would therefore expect that changes in ion concentration at the basement membrane would interfere with these electrostatic interactions and cause these proteins to dissociate from the HSPGs to which they are bound. Indeed, we saw increased baseline release of CCL20 in the presence of various elevated cations, suggesting that changing the extracellular ion balance may be sufficient to passively dissociate CCL20 from the cell surface. Given that these cations did not induce an additional increase in CCL20 release after HDM exposure, this suggests the change in cations predominately affected release of extracellular stores. This mechanism requires further study. For example, we would expect that mouse lungs deficient in baseline syndecan release, for example in syndecan-1 knockout mice, resident lung DCs would be reduced in comparison with mice expressing syndecan-1. However, separating intracellular and extracellular impacts of syndecan-1 knockout mice may be difficult to do, as we discuss below. As we have seen, a number of mechanisms involving ion transport may mediate release of chemokines and cytokines by bronchial epithelial cells. This is not surprising, given the role of the airway epithelium in maintaining the ionic composition of the sol in the airway lumen. Invasion of this environment by bacteria or other microbes would 202 dramatically alter airway ionic composition and pH, which would serve as a danger signal to the epithelium to release chemo-attractants to help fight off invaders. In addition to direct release from HSPGs, our data suggest that release of extracellular chemokines and cytokines is potentiated by cleavage of syndecan-1. We observed that tracheal epithelial cells from syndecan-1 knockout mice showed about a 50% reduction in release of both CCL20 and IL-8 (data not shown), suggesting that the extracellular CCL20 pool previously discussed localizes to syndecan-1. Another possible explanation for this reduction is that if stores of CCL20 within the lysosome are bound to syndecan-1, these would no longer be protected from degradation in the lysosomes of knockout mice. Future research will need to determine whether the reduction of CCL20 release in syndecan-1 knockout mice is due the absence of an extracellular storage location, or to the degradation of intracellular stores. For example, if CCL20 is degraded in the lysosomes of syndecan-1 knockout mice, we would expect this to be reflected in an absence of CCL20 intracellular staining in immunofluorescence images of mouse lungs from knockout mice. Syndecan shedding has been shown to be dependent on the PKC and MAP kinase signaling pathways, and potentially JNK signaling as well. 32 Thus many, but not all, of the signaling pathways we found to be involved in CCL20 release from secretory granules have also been associated with syndecan release. From these studies, we 203 hypothesize that although CCL20 may be stored in multiple locations throughout the cell, release from these locations may not be meditated by mutually exclusive pathways but is rather mediated by similar mechanisms. Future studies will lead to a better understanding of the common pathways that lead to chemokine release from multiple cellular sources at once, as well as those pathways that lead to mobilization of only specific cellular sources. An enhanced understanding of this complex intracellular signaling network will help in further understanding why different allergens induce release of different subsets of chemokines and cytokines from the bronchial epithelium. 204 Chemokine and cytokine combinations found in secretory lysosomes work together to promote a T H2 response In addition to antimicrobial activity; CCL20, IP-10, and other chemokines have also been shown to directly kill parasites such as Leishmania mexicana. 33 It is not surprising, then, that we found other antimicrobial chemokines, such as gro-α, are controlled by a similar release mechanism as that of CCL20. However, while CCL20 recruits iDCs and T H17 T-cells; gro-α, IP-10, and IL-8 recruit neutrophils.34 These divergent activities allow rapidly released chemokines to actively respond to danger signals through direct killing, while also contributing in diverse ways to the downstream inflammatory response. 35 Neutrophils, T H17 cells, and dendritic cells have all been documented in asthmatic lungs – and more particularly in the lungs of severe asthmatics. 36–38 Our findings suggest a common mechanism initiating the release of the chemotactic agents responsible for recruiting these cell types, as well as cytokines that will act on the recruited cells to induce an asthmatic T H2/T H17 phenotype. In addition to those we specifically identified, we expect a number of other preformed chemokines may also be subject to this release mechanism, given their common chemotactic functions and observed rapid release. Some examples of chemokines we expect may are rapidly released include CCL20, gro-α, IL-8, and IP-10. Bronchial epithelial cells are not the sole source of chemokines in the lung, and mechanisms of storage and release in other cell types may by similar to that of epithelial cells. 205 Indeed, IL-8 stored in Weibel-Palade bodies in endothelial cells was shown to be rapidly released, which is strikingly similar to the mechanism we have described for chemokine release from bronchial epithelial cells. 39 This may suggest that secretory lysosome release of vesicles containing chemokines and cytokines is common to additional cell types as well. If these vesicles resemble the vesicles we observed in bronchial epithelial cells, they may also contain a mixture of different chemokines and cytokines that are co-released. As outlined previously, a mechanism of secretory granule release, similar to the one described in this study, has also been described for cells of the hematopoietic lineage as well as for endothelial cells. Interestingly, our findings are the first to describe granule release from cells that are not derived from the mesoderm, as epithelial cells are derived from the endoderm. This suggests that this mechanism may be found in other endoderm-derived cell types in other tissues. Particularly, we hypothesize that release of chemokines from secretory granules may also occur in nasal, gut, and skin epithelial cells, as well as other types of epithelial cells. Future studies would need to determine which, if any, other cell types retain persistent vesicular stores of chemokines and cytokines in this manner, as well as what agonists trigger their release. 206 SOFA is a powerful new tool for analyzing subcellular organelles One major experimental difficulty we encountered was in positively identifying that intracellular chemokine stores were co-localized to the same vesicle. We showed that one promising avenue for identifying the contents of individual granules is through single organelle fluorescence analysis (SOFA). We demonstrated that multi- color flow cytometry, could be performed on subcellular components using SOFA. This technique could be useful in further understanding the nature and components of chemokine and cytokine-containing granules. Specifically, by taking advantage of flow- based sorting techniques, such as fluorescence-activated organelle sorting (FAOS), organelles containing different subsets of chemokines and cytokines could be isolated and analyzed. These sorted populations could then be separated by 2D gel electrophoresis in preparation for peptide mass fingerprinting via matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS), which would allow specific identification of individual proteins residing in different types of granules. A major technical difficulty with this approach is that the sample size required to obtain enough secretory granules to form a sortable population could become prohibitively large. Some proteins we expect to find in these secretory vesicles include proteins involved in secretory granule release, such as synaptotagmin and synaptobrevin – two proteins that make up part of the SNARE complex, and facilitate vesicle fusion with the 207 plasma membrane. 40–42 Identifying the specific SNARE proteins localized to these secretory granules would be beneficial in developing new therapies to block their release. Unlike secretory granules found in some hematopoietic cells, we would not expect epithelial cells to contain effector molecules that drive cell death, such as perforin and granzyme. 43–46 These effector molecules mediate cell death in natural killer cells and cytotoxic T-lymphocytes; 47,48 however perforin-mediated cell death is not a function that has been observed in non-migratory airway epithelial cells. 49 CCL20 is sufficient, but not necessary to promote AHR The importance of CCL20/CCR6 in allergic sensitization was demonstrated in a cockroach model of allergic asthma in which CCR6 knockout mice displayed protection against development of asthma. 50,51 Contrary to these findings, in which cockroach frass was the major allergen studied; we observed that in a HDM-exposure model of asthma, blocking CCL20 was insufficient to eliminate AHR. This discrepancy may be due to the presence of a number of signals, other than CCL20, released in response to HDM exposure that have also been demonstrated as important in the development of AHR. AHR that is driven by cockroach allergen may be primarily dependent on signaling through the CCL20/CCR6 axis, whereas AHR initiated by HDM may induce more robust response, including the release of a more broad array of chemokines and cytokines. Our data suggest that a potential reason for this discrepancy may be due in part to the different initial signals provided by HDM versus cockroach frass. In 208 addition to CCL20, HDM induced release of FGF-2, Gro-α, TNF-α, IL-6, IL-8, and IP-10, among others. In contrast, cockroach frass did not induce release of IL-6 and IP-10, and caused release of significantly lower levels of FGF-2, gro-α, TNF-α, and IL-8. In a cockroach model, blockage of CCL20 signaling may result in reduction of downstream signaling and eventual development of AHR because it is a large component of the bronchial epithelial cells’ initial response to the allergen. In contrast, HDM induces release of a much broader array of chemokines and cytokines from the airway epithelium. All these signals function to promote inflammation through recruitment of a number of cell types, such as DCs and TH17 cells for CCL20, T-cells for IP-10, and neutrophils for IL-8 and gro-α. IL-6 has been shown to inhibit the development of regulatory T cells in favor of T H17 cells, 52 which augment a T H2 response in asthmatics to promote more severe AHR. 53 Thus, our data suggest that HDM induces the release of multiple signals that combine to promote the development of a T H2 response. As such, loss of one signal, namely CCL20, appears to be insufficient to block the overall development of AHR. Both cockroach frass and HDM are common, complex allergens. However, a number of factors contribute to the more robust signal provided by HDM in comparison to cockroach frass and many other allergens. HDM is a complex mixture of several different types of agonists including proteases, chitin, and LPS. 54–56 Thus, multiple signals work together to promote a T H2 response. It is a potent allergen, with 209 atopy to HDM more common in asthmatics than any other allergen. This makes targeting pathways for HDM-induced AHR a valuable strategy for developing novel asthma treatments, since many of the mechanisms that are necessary for chemokine and cytokine release in response to HDM are also likely to be activated in response to other, less complex allergens. Cockroach frass is also a complex allergen, although it is not as potent and not as prevalent in atopic individuals. Frass also contains chitin, active proteases, and LPS, which may explain why it induced release of more chemokines and cytokines from the bronchial epithelium than other allergens we tested, such as tree and grass pollens. The mechanism we describe, in which chemokines and cytokines are stored in and released from intracellular vesicles, is a novel mechanism in bronchial epithelial cells, and may help to explain how upstream signaling pathways lead to the release of different subsets of chemokines and cytokines and therefore different outcomes from exposure to a variety allergens. Different allergens induce release of unique subsets of chemokines and cytokines to promote AHR The bronchial epithelium has recently been highlighted as an important source for a number of different chemokines and cytokines that are critical in the induction of atopic asthma. Much recent focus has turned to IL-33, as well as TSLP, as airway epithelial cell-derived mediators that play an important role in the development of type-2 immune responses. IL-33 expression in the bronchial epithelium correlated with 210 asthma severity in an OVA model of AHR,57,58 and it has been shown to induce production of the TH2 cytokines IL-4, IL-5, and IL-13 as well as airway inflammation. 59,60 TSLP expression has also been shown to be increased in human asthmatic airways, and to induce a type-2 immune response. 61,62 When we consider our finding that CCL20 and IL-33 may share a common release mechanism, this suggests a model in which airway epithelial cells co-release CCL20 and IL-33 from lysosomal stores upon stimulation with insect allergens, such as HDM or cockroach frass. Under these circumstances, CCL20 would work to promote recruitment of cells containing CCR6, such as iDCs, innate lymphoid cells, and T H17 cells, while IL-33 would promote the development of a type-2 response through DCs and ultimately the downstream development of a T H2 response.60,63–67 In addition to our finding that IL-33 appears in punctate vesicles within bronchial epithelial cells, we postulate other potential release mechanisms for preformed IL-33. One example of a cytokine belonging to the IL-1 superfamily with multiple demonstrated release mechanisms is IL-1β. This cytokine has long been known to be stored within the cytoplasm, but recent evidence has indicated that it may also be stored in the secretory lysosomes of monocytes.68 IL-33 may be stored in a similar fashion, with both cytoplasmic and lysosomal storage locations. Although the process of protein release from within vesicles has long been established as simple fusion of the vesicle and plasma membranes, it is unclear how IL-1 family members 211 stored in the cytosol are released from the cell, given the necessity of crossing a lipid bilayer in order to exit the membrane-bound area. Two potential mechanisms for cytoplasmic release of IL-1β that have been proposed include cell death and release of micro-vesicles that are the result of membrane blebbing. 69 As a closely related cytokine to IL-1β, IL-33 may also be stored inside the cell by similar mechanisms. Indeed, staining overlays of both CCL20 and IL-33 in our confocal images suggest that these two proteins share many common secretory vesicles, but additional non-punctate staining of IL-33 is prevalent – leaving the possibility that additional cellular storage locations for IL-33 exist. We also found cellular stores of IL-1α that were rapidly released by HDM. However, these stores did not appear to localize to secretory lysosomes or to discreet intracellular pools, indicating a potential cytosolic storage for this cytokine as well. A unique mechanism of pre-release processing for the IL-1 family member IL-1β has been reported previously, in which activation of the inflammasome by a number of factors, such as uric acid or the presence of external ATP, leads to cleavage of pro-IL-1β. 70,71 This cleavage converts pro-IL-1β to IL-1β in preparation for release of the active cytokine. 72 Although this mechanism shows how IL-1β is processed into its active form, little is known about how IL-1β is subsequently exported from the cell. We observed a diffuse distribution of IL-1α, suggesting a cytosolic distribution similar to IL-1β, and although the inflammasome may not be involved in IL-1α processing, this closely 212 related protein may share a similar release mechanism with IL-1β. A separate release mechanism for IL-1α that has previously been reported is unlikely to be involved in HDM-induced IL-1α release. In this mechanism, IL-1α acts as an alarmin; it translocates to the nucleus in the event of cell damage, and is passively released in the event of cell death. 73 Our data suggest IL-1α was released from healthy bronchial epithelial cells, suggesting that cell death is not the mechanism of release for this cytokine. Additionally, our ELISA measurements are unlikely to detect membrane-bound cytokine, suggesting membrane blebbing is also not the mechanism governing release of IL-1α. Other possible mechanisms for release of proteins from the cytoplasm include active transport across the cell membrane or loading into intracellular transport vesicles. Additional experiments are needed to more fully understand the mechanisms governing allergen-induced release of these IL-1 family members. Common storage and release patterns, especially for chemokines and cytokines stored in a common vesicle or released through a common pathway, suggest the signaling molecules are involved in a coordinated function. In bronchial epithelial cells treated with HDM, we saw nearly complete ablation of intracellular stores of CCL20. Stores of IL-33 that reside in the same vesicles as CCL20 would also be expected to be released after HDM exposure. Co-release would allow CCL20 to induce recruitment of iDCs and T H17 T-cells, while IL-33 induces DCs to promote a downstream T H2 response.60 Similar co-storage and release after HDM exposure was observed with gro- 213 α, which induces recruitment of neutrophils, as previously stated, also contributing to the development of an asthma phenotype. In contrast to IL-33 and gro-α, we did not detect any preformed stores of TSLP; although we did see TSLP release after two hours of ragweed exposure (data not shown), similar to what other groups have observed.74 Of the panel of cytokines and chemokines we looked at, only CCL20 and TSLP were released in response to ragweed pollen. Therefore, ragweed exposure suggests an alternate route to the development of a T H2 response, in which CCL20 works to recruit iDCs and TH17 cells, and TSLP acts on these recruited iDCs to promote a T H2 phenotype.75 We have reviewed two major pathways for release of preformed cytokine from bronchial epithelial cells that appear to play important roles in the development of a TH2 response in asthma, although only specific pathways are activated by any individual allergen. Thus, different classes of allergens may initiate a T H2 response in unique ways. In addition to these two pathways leading to protein release, two other release mechanisms are suggested by our findings: de novo synthesis, and cell surface storage. A better understanding of each of these mechanisms, and the signaling pathways that lead to their release, may help in the development of future targeted treatments for asthma. 214 Given that individual allergens appear to induce the release of specific subsets of chemokines and cytokines, the specific pathways triggered by an individual allergen could be blocked in asthmatics based on atopy to specific allergens, thereby providing targeted inhibition and reducing overall immune suppression. For example, in our hands ragweed pollen predominantly induced release of TSLP and CCL20, but not other mediators, from the bronchial epithelium. In patients who display atopy to ragweed, blockade of TSLP – directly through anti-TSLP neutralizing antibodies or indirectly through blocking the signaling pathway leading to TSLP release – may be an effective treatment to reduce airway inflammation. As CCL20 induced recruitment of iDCs to the airway epithelium, TSLP would be blocked from promoting a T H2 response in these recruited iDCs, preventing the development of AHR. This same treatment would likely prove ineffective in patients who are atopic to HDM, as HDM does not induce immediate release of TSLP from bronchial epithelial cells but does induce release of broad range of different chemokines and cytokines that lead to the development of a T H2 response. Targeting cytokine release mechanisms may be a more effective therapy than targeting individual cytokines for some types of atopic asthma Our research has focused on better understanding the mechanisms of allergen- induced chemokine and cytokine release, with the ultimate goal that this may lead to better therapies for asthmatics. Currently, a number of cytokine-based therapies are 215 undergoing clinical trials for the treatment of asthma. These therapies rely on anti- cytokine antibodies to neutralize the effects of the target cytokine. Some targets that are being pursued include IL-4, 76 IL-5,77 and Il-13 78 (key mediators of the type-2 response in asthma)79,80 as well as TNF-α.81 Although anti-cytokine therapy showed great promise in animal models of disease, trials in humans have proven less effective at alleviating AHR and airway inflammation, 82–84 with some authors concluding that blockade of multiple mediators may provide more robust results.85 Additionally, new targets for therapy are being considered, including anti-IL-33 86,87 and anti-TSLP 88 , among others. Our work suggests a number of potential therapeutic approaches for suppression of chemokine and cytokine release by targeting release pathways as opposed to directly targeting individual cytokines. A few examples include targeting epithelial secretory lysosomes directly, targeting pattern recognition receptors, or targeting the downstream signaling pathways they initiate. We will consider each of these targets and their potential issues in using targeting them for therapy. MAPK signaling pathway activation in a number of different cell types has been implicated in allergic airway inflammation. 89 ERK1/2 and p38 phosphorylation have been shown to contribute to IL- 8 release, 90 and activation of Raf1 has been shown to induce gro-a in epithelial cells. 91 Ras-Raf1-Syk signaling mediates secretory lysosome function 92–96 . Additionally, ICAM- 1 expression in the bronchial epithelium was shown to be dependent on signaling through both ERK and JNK. 97 General inhibition of MAPK signaling led to a decrease 216 in airway inflammation and AHR. 98,99 Thus, signaling through the MAPK pathway; including Raf1, ERK, and JNK; is an attractive target to inhibit the initial airway epithelial response to allergens, as well as the treatment of asthma in general. There are a number of potential drawbacks with this broader approach. Directly targeting lysosomal release could potentially lead to impaired neurological function if used as a long-term therapy, similar to individuals suffering from lysosomal storage diseases such as Tay-Sachs or Salla disease. 100,101 Long-term inhibition of lysosomal release causes a build-up of waste products within the cell, as proteins targeted for degradation are loaded into endosomal bodies, but are never degraded. These endosomal bodies swell the size of the cell, and interfere with normal cell function. Neurons are particularly susceptible to inhibition of lysosome function.102 Another promising treatment target is the Raf1 signaling pathway. Inhibitors to Raf-1 have been developed for treatment in a number of different cancers, including hepatocellular carcinoma. 103 The treatment has been well-tolerated, with skin rashes of the hand and foot being the most common adverse effect documented to date. 104 A larger problem with targeting the release of a broad range of chemokines and cytokines is that these proteins play an important protective role against invading pathogens. For example, in response to respiratory syncytial virus (RSV) a number of chemokines and cytokines are released by the bronchial epithelium, including RANTES, 217 IL-8, and IL-6 – all of which are released in response to HDM in our hands. 105 This suggests that blocking mechanisms of HDM-induced chemokine and cytokine release would likely target RSV-induced immune responses, thereby enhancing susceptibility of these patients to infection. However, some evidence suggests that blocking CCL20 release during RSV infection may diminish adverse responses to RSV by blocking conventional, but not plasmacytoid DC recruitment. 106 Thus, it is not immediately clear that blocking chemokine and cytokine release would dramatically exacerbate RSV infection. One way to minimize these adverse effects would be to focus the more broad lysosomal-release-targeted therapy on severe asthmatics. Severe asthmatics display increased neutrophil inflammation in the lungs, as well as increased TH17 T-cells. 38,107 Our data suggest that inhibiting the upstream mechanisms that control release of intracellular CCL20 would also reduce release of IL-8 and gro-α, which have been shown to induce neutrophil recruitment, and IP-10, which has been shown to contribute to recruitment of T-cells and dendritic cells.108–110 A more ideal treatment paradigm would be to identify patients who are susceptible to developing asthma, and expose them to allergen while blocking lysosomal chemokine release. This would have the added benefit, as a short-term therapy, of avoiding many of the long-term adverse effects mentioned above. Unfortunately, this approach would have little benefit for 218 individuals who already have asthma and have developed a T H2 response to an allergen. One of the benefits of targeting common release pathways instead of individual chemokines and cytokines is that it would reduce the release of multiple mediators simultaneously. This has the practical effect of targeting multiple downstream pathways that lead to AHR, such as neutrophil recruitment, DC recruitment, and T-cell recruitment, among others. Our own data suggest that blocking CCL20 alone had a limited effect on HDM-induced AHR and inflammation. Other groups showed reduced inflammation in CCR6 knockout mice stimulated with either cockroach frass 111 or particulate matter.51 In our hands, HDM induced the release of a broader variety of chemokines and cytokines than any other allergen, including cockroach frass. Thus, for some atopic individuals targeting a single chemokine/receptor pair, like CCL20/CCR6, may be sufficient to reduce asthma symptoms. But for atopic asthmatics sensitive to HDM, the effect of singly targeting CCL20/CCR6 may be blunted by the release of a number of other inflammatory mediators. For these individuals, targeting a release mechanism for chemokines and cytokines should be considered, as it would inhibit the release of a broader range of inflammatory mediators. Since a majority of atopic asthmatics respond to HDM, we would not expect studies that target individual chemokines and cytokines released from the bronchial epithelium to have sufficient 219 power to detect the potentially potent effect these therapies might have on minority populations of asthmatics who are not atopic to HDM. Currently, many trials involving anti-cytokine therapies are focused on stratification of patients based on asthma severity. 112,113 Our data suggest a different method for stratification: allergen sensitivity. Although we would not expect to observe a difference in treatment efficacy for downstream cytokines such as IL-4 and IL-5, we would expect that stratification of patients based on allergen sensitivity for anti- chemokine/cytokine therapy targets such as IL-33, TSLP, and CCL20 would reveal groups of patients susceptible to treatment. These groups would potentially include individuals who are atopic to cockroach (CCL20), and ragweed (TSLP). An additional benefit to this approach is that separation based on atopy can be determined fairly easily and inexpensively. Conclusions The work we have presented in this dissertation contributes to our understanding of chemokine and cytokine storage and release mechanisms in bronchial epithelial cells. Release of a number of important chemokines and cytokines – including CCL20, Gro-α, IL-8, IL-33, and TNF-α – previously thought to be dependent on de novo synthesis, have been shown to exist within pre-formed stores in bronchial epithelial cells in both intracellular and extracellular pools. These stores may then be selected for 220 release, depending on a number of stimuli. Intracellular stores of some chemokines, such as CCL20, are found in secretory lysosomes; and multiple mediators may localize to the same secretory granules. A number of diverse signaling pathways lead to secretory granule loading and release, including Raf1 and JNK. Of the vesicles that are stored within the bronchial epithelium, the specific vesicles that are released depend on the particular allergen the cell is exposed to. A diverse array of chemokine- and cytokine-specific responses are mounted to each allergen the bronchial epithelium is exposed to. Of all the allergens we tested, HDM was the most potent at inducing release of preformed stores of chemokines and cytokines. Although intracellular storage and release is not the sole means by which these mediators are delivered to the extracellular environment, our findings illustrate that they play an important role in inflammation. These studies expand our understanding of how chemokines and cytokines are released from the bronchial epithelium and provide a number of potential targets for treatment of allergic airway diseases. 221 References 1. Kuipers H, Lambrecht BN. The interplay of dendritic cells, Th2 cells and regulatory T cells in asthma. Current Opinion in Immunology . 2004;16(6):702–708. 2. Pichavant M, Charbonnier A-S, Taront S, et al. Asthmatic bronchial epithelium activated by the proteolytic allergen Der p 1 increases selective dendritic cell recruitment. J. Allergy Clin. Immunol. 2005;115(4):771–778. 3. Jahnsen FL, Moloney ED, Hogan T, et al. Rapid Dendritic Cell Recruitment to the Bronchial Mucosa of Patients with Atopic Asthma in Response to Local Allergen Challenge. Thorax . 2001;56(11):823–826. 4. Hoover DM, Boulègue C, Yang D, et al. The Structure of Human Macrophage Inflammatory Protein-3α/CCL20 LINKING ANTIMICROBIAL AND CC CHEMOKINE RECEPTOR-6-BINDING ACTIVITIES WITH HUMAN Β-DEFENSINS. J. Biol. Chem. 2002;277(40):37647–37654. 5. Francis JN, Sabroe I, Lloyd CM, Durham SR, Till SJ. Elevated CCR6+ CD4+ T lymphocytes in tissue compared with blood and induction of CCL20 during the asthmatic late response. Clin Exp Immunol . 2008;152(3):440–447. 6. Braulke T, Bonifacino JS. Sorting of lysosomal proteins. Biochimica et Biophysica Acta (BBA) - Molecular Cell Research . 2009;1793(4):605–614. 7. Kolodny EH, Mumford RA. Human leukocyte acid hydrolases: Characterization of eleven lysosomal enzymes and study of reaction conditions for their automated analysis. Clinica Chimica Acta . 1976;70(2):247–257. 8. Van Dongen JM, Willemsen R, Ginns EI, et al. The subcellular localization of soluble and membrane-bound lysosomal enzymes in I-cell fibroblasts: a comparative immunocytochemical study. Eur. J. Cell Biol. 1985;39(1):179–189. 9. Serafin WE, Katz HR, Austen KF, Stevens RL. Complexes of Heparin Proteoglycans, Chondroitin Sulfate E Proteoglycans, and [3H]diisopropyl Fluorophosphate-Binding Proteins Are Exocytosed from Activated Mouse Bone Marrow-Derived Mast Cells. J. Biol. Chem. 1986;261(32):15017–15021. 10. Webb LM, Ehrengruber MU, Clark-Lewis I, Baggiolini M, Rot A. Binding to Heparan Sulfate or Heparin Enhances Neutrophil Responses to Interleukin 8. PNAS . 1993;90(15):7158–7162. 11. Witt DP, Lander AD. Differential binding of chemokines to glycosaminoglycan subpopulations. Curr. Biol. 1994;4(5):394–400. 12. Gitay-Goren H, Soker S, Vlodavsky I, Neufeld G. The Binding of Vascular Endothelial Growth Factor to Its Receptors Is Dependent on Cell Surface-Associated Heparin-Like Molecules. J. Biol. Chem. 1992;267(9):6093–6098. 13. Marshall LJ, Ramdin LSP, Brooks T, DPhil PC, Shute JK. Plasminogen Activator Inhibitor-1 Supports IL-8-Mediated Neutrophil Transendothelial Migration by 222 Inhibition of the Constitutive Shedding of Endothelial IL-8/Heparan Sulfate/Syndecan-1 Complexes. J Immunol . 2003;171(4):2057–2065. 14. Lortat-Jacob H, Grosdidier A, Imberty A. Structural Diversity of Heparan Sulfate Binding Domains in Chemokines. PNAS . 2002;99(3):1229–1234. 15. Chan DI, Hunter HN, Tack BF, Vogel HJ. Human Macrophage Inflammatory Protein 3α: Protein and Peptide Nuclear Magnetic Resonance Solution Structures, Dimerization, Dynamics, and Anti-Infective Properties. Antimicrob. Agents Chemother. 2008;52(3):883–894. 16. Koopmann W, Krangel MS. Identification of a Glycosaminoglycan-binding Site in Chemokine Macrophage Inflammatory Protein-1α. J. Biol. Chem. 1997;272(15):10103–10109. 17. Yang D, Chen Q, Hoover DM, et al. Many Chemokines Including CCL20/MIP-3α Display Antimicrobial Activity. J Leukoc Biol . 2003;74(3):448–455. 18. Ricciardolo FLM, Gaston B, Hunt J. Acid stress in the pathology of asthma. J. Allergy Clin. Immunol. 2004;113(4):610–619. 19. Ramsden L, Rider CC. Selective and differential binding of interleukin (IL)-1 alpha, IL-1 beta, IL-2 and IL-6 to glycosaminoglycans. Eur. J. Immunol. 1992;22(11):3027–3031. 20. Luster AD, Greenberg SM, Leder P. The IP-10 chemokine binds to a specific cell surface heparan sulfate site shared with platelet factor 4 and inhibits endothelial cell proliferation. J Exp Med . 1995;182(1):219–231. 21. Guimond S, Maccarana M, Olwin BB, Lindahl U, Rapraeger AC. Activating and inhibitory heparin sequences for FGF-2 (basic FGF). Distinct requirements for FGF-1, FGF-2, and FGF-4. J. Biol. Chem. 1993;268(32):23906–23914. 22. Charbonnier A-S, Kohrgruber N, Kriehuber E, et al. Macrophage Inflammatory Protein 3α Is Involved in the Constitutive Trafficking of Epidermal Langerhans Cells. J Exp Med . 1999;190(12):1755–1768. 23. Naccache PH, Showell HJ, Becker EL, Sha’afi RI. Changes in ionic movements across rabbit polymorphonuclear leukocyte membranes during lysosomal enzyme release. Possible ionic basis for lysosomal enzyme release. J Cell Biol . 1977;75(3):635– 649. 24. Pfeffer SR. Transport-vesicle targeting: tethers before SNAREs. Nat Cell Biol . 1999;1(1):E17–E22. 25. Lin RC, Scheller RH. Mechanisms of Synaptic Vesicle Exocytosis. Annual Review of Cell and Developmental Biology . 2000;16(1):19–49. 26. PhD WFBM, MD ELB. Medical Physiology, Updated Edition: With STUDENT CONSULT Online Access, 1e. Saunders; 2004. 27. Pazoles CJ, Pollard HB. Evidence for stimulation of anion transport in ATP- evoked transmitter release from isolated secretory vesicles. J. Biol. Chem. 1978;253(11):3962–3969. 223 28. Ceccarelli B, Grohovaz F, Hurlbut WP. Freeze-fracture studies of frog neuromuscular junctions during intense release of neurotransmitter. II. Effects of electrical stimulation and high potassium. J Cell Biol . 1979;81(1):178–192. 29. Gennaro JF, Nastuk WL, Rutherford DT. Reversible depletion of synaptic vesicles induced by application of high external potassium to the frog neuromuscular junction. J Physiol . 1978;280(1):237–247. 30. Sı́k A, Smith R., Freund T. Distribution of chloride channel-2-immunoreactive neuronal and astrocytic processes in the hippocampus. Neuroscience . 2000;101(1):51– 65. 31. Robitaille R, Garcia ML, Kaczorowski GJ, Chariton MP. Functional colocalization of calcium and calcium-gated potassium channels in control of transmitter release. Neuron . 1993;11(4):645–655. 32. Fitzgerald ML, Wang Z, Park PW, Murphy G, Bernfield M. Shedding of Syndecan-1 and -4 Ectodomains Is Regulated by Multiple Signaling Pathways and Mediated by a Timp-3–Sensitive Metalloproteinase. J Cell Biol . 2000;148(4):811–824. 33. Söbirk SK, Mörgelin M, Egesten A, et al. Human Chemokines as Antimicrobial Peptides with Direct Parasiticidal Effect on Leishmania mexicana In Vitro. PLoS ONE . 2013;8(3):e58129. 34. Geiser T, Dewald B, Ehrengruber MU, Clark-Lewis I, Baggiolini M. The interleukin-8-related chemotactic cytokines GRO alpha, GRO beta, and GRO gamma activate human neutrophil and basophil leukocytes. J. Biol. Chem. 1993;268(21):15419– 15424. 35. Dürr M, Peschel A. Chemokines Meet Defensins: the Merging Concepts of Chemoattractants and Antimicrobial Peptides in Host Defense. Infect. Immun. 2002;70(12):6515–6517. 36. Bradley BL, Azzawi M, Jacobson M, et al. Eosinophils, T-lymphocytes, mast cells, neutrophils, and macrophages in bronchial biopsy specimens from atopic subjects with asthma: Comparison with biopsy specimens from atopic subjects without asthma and normal control subjects and relationship to bronchial hyperresponsiveness. Journal of Allergy and Clinical Immunology . 1991;88(4):661–674. 37. Pène J, Chevalier S, Preisser L, et al. Chronically Inflamed Human Tissues Are Infiltrated by Highly Differentiated Th17 Lymphocytes. J Immunol . 2008;180(11):7423– 7430. 38. Wenzel SE, Schwartz LB, Langmack EL, et al. Evidence That Severe Asthma Can Be Divided Pathologically into Two Inflammatory Subtypes with Distinct Physiologic and Clinical Characteristics. Am. J. Respir. Crit. Care Med. 1999;160(3):1001–1008. 39. Utgaard JO, Jahnsen FL, Bakka A, Brandtzaeg P, Haraldsen G. Rapid Secretion of Prestored Interleukin 8 from Weibel-Palade Bodies of Microvascular Endothelial Cells. J Exp Med . 1998;188(9):1751–1756. 224 40. Söllner T, Bennett MK, Whiteheart SW, Scheller RH, Rothman JE. A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell . 1993;75(3):409–418. 41. Fasshauer D, Eliason WK, Brünger AT, Jahn R. Identification of a Minimal Core of the Synaptic SNARE Complex Sufficient for Reversible Assembly and Disassembly†. Biochemistry . 1998;37(29):10354–10362. 42. Davis AF, Bai J, Fasshauer D, et al. Kinetics of Synaptotagmin Responses to Ca2+ and Assembly with the Core SNARE Complex onto Membranes. Neuron . 1999;24(2):363–376. 43. Jenne DE, Tschopp J. Granzymes, a Family of Serine Proteases Released from Granules of Cytolytic T Lymphocytes upon T Cell Receptor Stimulation. Immunological Reviews . 1988;103(1):53–71. 44. Peters PJ, Borst J, Oorschot V, et al. Cytotoxic T Lymphocyte Granules Are Secretory Lysosomes, Containing Both Perforin and Granzymes. J Exp Med . 1991;173(5):1099–1109. 45. Esser MT, Krishnamurthy B, Braciale VL. Distinct T cell receptor signaling requirements for perforin- or FasL-mediated cytotoxicity. J Exp Med . 1996;183(4):1697–1706. 46. Wood SM, Meeths M, Chiang SCC, et al. Different NK cell–activating receptors preferentially recruit Rab27a or Munc13-4 to perforin-containing granules for cytotoxicity. Blood . 2009;114(19):4117–4127. 47. Kägi D, Ledermann B, Bürki K, et al. Cytotoxicity mediated by T cells and natural killer cells is greatly impaired in perforin-deficient mice. , Published online: 05 May 1994; | doi:10.1038/369031a0 . 1994;369(6475):31–37. 48. Shresta S, MacIvor DM, Heusel JW, Russell JH, Ley TJ. Natural killer and lymphokine-activated killer cells require granzyme B for the rapid induction of apoptosis in susceptible target cells. PNAS . 1995;92(12):5679–5683. 49. Su AI, Cooke MP, Ching KA, et al. Large-scale analysis of the human and mouse transcriptomes. PNAS . 2002;99(7):4465–4470. 50. Lukacs NW, Prosser DM, Wiekowski M, Lira SA, Cook DN. Requirement for the Chemokine Receptor Ccr6 in Allergic Pulmonary Inflammation. J Exp Med . 2001;194(4):551–556. 51. Lundy SK, Lira SA, Smit JJ, et al. Attenuation of Allergen-Induced Responses in CCR6–/– Mice Is Dependent Upon Altered Pulmonary T Lymphocyte Activation. J Immunol . 2005;174(4):2054–2060. 52. Bettelli E, Carrier Y, Gao W, et al. Reciprocal developmental pathways for the generation of pathogenic effector TH17 and regulatory T cells. Nature . 2006;441(7090):235–238. 225 53. Lajoie S, Lewkowich IP, Suzuki Y, et al. Complement-mediated regulation of the IL-17A axis is a central genetic determinant of the severity of experimental allergic asthma. Nat Immunol . 2010;11:928–35. 54. Gough L, Schulz O, Sewell HF, Shakib F. The Cysteine Protease Activity of the Major Dust Mite Allergen Der P 1 Selectively Enhances the Immunoglobulin E Antibody Response. J Exp Med . 1999;190(12):1897–1902. 55. Neville AC, Parry DA, Woodhead-Galloway J. The Chitin Crystallite in Arthropod Cuticle. J Cell Sci . 1976;21(1):73–82. 56. Michel O, Kips J, Duchateau J, et al. Severity of Asthma Is Related to Endotoxin in House Dust. Am. J. Respir. Crit. Care Med. 1996;154(6):1641–1646. 57. Préfontaine D, Lajoie-Kadoch S, Foley S, et al. Increased Expression of IL-33 in Severe Asthma: Evidence of Expression by Airway Smooth Muscle Cells. J Immunol . 2009;183(8):5094–5103. 58. Préfontaine D, Nadigel J, Chouiali F, et al. Increased IL-33 expression by epithelial cells in bronchial asthma. Journal of Allergy and Clinical Immunology . 2010;125(3):752–754. 59. Schmitz J, Owyang A, Oldham E, et al. IL-33, an Interleukin-1-like Cytokine that Signals via the IL-1 Receptor-Related Protein ST2 and Induces T Helper Type 2- Associated Cytokines. Immunity . 2005;23(5):479–490. 60. Rank MA, Kobayashi T, Kozaki H, et al. IL-33–activated dendritic cells induce an atypical TH2-type response. Journal of Allergy and Clinical Immunology . 2009;123(5):1047–1054. 61. Ying S, O’Connor B, Ratoff J, et al. Thymic Stromal Lymphopoietin Expression Is Increased in Asthmatic Airways and Correlates with Expression of Th2-Attracting Chemokines and Disease Severity. J Immunol . 2005;174(12):8183–8190. 62. Rochman I, Watanabe N, Arima K, Liu Y-J, Leonard WJ. Cutting Edge: Direct Action of Thymic Stromal Lymphopoietin on Activated Human CD4+ T Cells. J Immunol. 2007;178(11):6720–6724. 63. Lügering A, Ross M, Sieker M, et al. CCR6 identifies lymphoid tissue inducer cells within cryptopatches. Clinical & Experimental Immunology . 2010;160(3):440–449. 64. Schutyser E, Struyf S, Van Damme J. The CC chemokine CCL20 and its receptor CCR6. Cytokine Growth Factor Rev. 2003;14(5):409–426. 65. Wang C, Kang SG, Lee J, Sun Z, Kim CH. The roles of CCR6 in migration of Th17 cells and regulation of effector T-cell balance in the gut. Mucosal Immunol . 2009;2:173– 83. 66. Kouzaki H, Iijima K, Kobayashi T, O’Grady SM, Kita H. The Danger Signal, Extracellular ATP, Is a Sensor for an Airborne Allergen and Triggers IL-33 Release and Innate Th2-Type Responses. J Immunol . 2011;186(7):4375–4387. 226 67. Barlow JL, Bellosi A, Hardman CS, et al. Innate IL-13–producing nuocytes arise during allergic lung inflammation and contribute to airways hyperreactivity. Journal of Allergy and Clinical Immunology . 2012;129(1):191–198.e4. 68. Andrei C, Margiocco P, Poggi A, et al. Phospholipases C and A2 control lysosome-mediated IL-1β secretion: Implications for inflammatory processes. PNAS . 2004;101(26):9745–9750. 69. MacKenzie A, Wilson HL, Kiss-Toth E, et al. Rapid Secretion of Interleukin-1β by Microvesicle Shedding. Immunity . 2001;15(5):825–835. 70. Martinon F, Pétrilli V, Mayor A, Tardivel A, Tschopp J. Gout-associated uric acid crystals activate the NALP3 inflammasome. Nature . 2006;440(7081):237–241. 71. Pelegrin P, Surprenant A. Pannexin-1 mediates large pore formation and interleukin-1β release by the ATP-gated P2X7 receptor. The EMBO Journal . 2006;25(21):5071–5082. 72. Martinon F, Burns K, Tschopp J. The Inflammasome. Molecular Cell . 2002;10(2):417–426. 73. Lamacchia C, Rodriguez E, Palmer G, Gabay C. Endogenous IL-1α is a chromatin-associated protein in mouse macrophages. Cytokine . 2013;63(2):135–144. 74. Esnault S, Rosenthal LA, Shen Z-J, et al. Thymic stromal lymphopoietin expression in allergic pulmonary inflammation is Pin1-dependent. Journal of Allergy and Clinical Immunology . 2008;121(5):1289–1290. 75. Soumelis V, Reche PA, Kanzler H, et al. Human epithelial cells trigger dendritic cell–mediated allergic inflammation by producing TSLP. Nature Immunology . 2002;3(7):673–680. 76. Hart TK, Blackburn MN, Brigham-Burke M, et al. Preclinical efficacy and safety of pascolizumab (SB 240683): a humanized anti-interleukin-4 antibody with therapeutic potential in asthma. Clinical & Experimental Immunology . 2002;130(1):93– 100. 77. Garlisi CG, Kung TT, Wang P, et al. Effects of Chronic Anti-Interleukin-5 Monoclonal Antibody Treatment in a Murine Model of Pulmonary Inflammation. Am. J. Respir. Cell Mol. Biol. 1999;20(2):248–255. 78. Singh D, Kane B, Molfino NA, et al. A phase 1 study evaluating the pharmacokinetics, safety and tolerability of repeat dosing with a human IL-13 antibody (CAT-354) in subjects with asthma. BMC Pulmonary Medicine . 2010;10(1):3. 79. Wills-Karp M, Luyimbazi J, Xu X, et al. Interleukin-13: Central Mediator of Allergic Asthma. Science . 1998;282(5397):2258–2261. 80. Yamashita M, Ukai-Tadenuma M, Miyamoto T, et al. Essential Role of GATA3 for the Maintenance of Type 2 Helper T (Th2) Cytokine Production and Chromatin Remodeling at the Th2 Cytokine Gene Loci. J. Biol. Chem. 2004;279(26):26983–26990. 81. Desai D, Brightling C. Cytokine and anti-cytokine therapy in asthma: ready for the clinic? Clinical & Experimental Immunology . 2009;158(1):10–19. 227 82. Borish LC, Nelson HS, Corren J, et al. Efficacy of soluble IL-4 receptor for the treatment of adults with asthma. Journal of Allergy and Clinical Immunology . 2001;107(6):963–970. 83. Wenzel S, Wilbraham D, Fuller R, Getz EB, Longphre M. Effect of an interleukin- 4 variant on late phase asthmatic response to allergen challenge in asthmatic patients: results of two phase 2a studies. The Lancet . 20;370(9596):1422–1431. 84. Flood-Page P, Swenson C, Faiferman I, et al. A Study to Evaluate Safety and Efficacy of Mepolizumab in Patients with Moderate Persistent Asthma. Am. J. Respir. Crit. Care Med. 2007;176(11):1062–1071. 85. Hansbro PM, Kaiko GE, Foster PS. Cytokine/anti-cytokine therapy – novel treatments for asthma? British Journal of Pharmacology . 2011;163(1):81–95. 86. Kurowska-Stolarska M, Kewin P, Murphy G, et al. IL-33 Induces Antigen- Specific IL-5+ T Cells and Promotes Allergic-Induced Airway Inflammation Independent of IL-4. J Immunol . 2008;181(7):4780–4790. 87. Kearley J, Buckland KF, Mathie SA, Lloyd CM. Resolution of Allergic Inflammation and Airway Hyperreactivity Is Dependent upon Disruption of the T1/ST2–IL-33 Pathway. Am. J. Respir. Crit. Care Med. 2009;179(9):772–781. 88. Shi L, Leu S-W, Xu F, et al. Local blockade of TSLP receptor alleviated allergic disease by regulating airway dendritic cells. Clinical Immunology . 2008;129(2):202–210. 89. Duan W, Wong WSF. Targeting mitogen-activated protein kinases for asthma. Curr Drug Targets . 2006;7(6):691–698. 90. Cui C-H, Adachi T, Oyamada H, et al. The Role of Mitogen-Activated Protein Kinases in Eotaxin-Induced Cytokine Production from Bronchial Epithelial Cells. Am. J. Respir. Cell Mol. Biol. 2002;27(3):329–335. 91. Kawaguchi M, Kokubu F, Matsukura S, et al. Induction of C-X-C Chemokines, Growth-Related Oncogene α Expression, and Epithelial Cell-Derived Neutrophil- Activating Protein-78 by ML-1 (Interleukin-17F) Involves Activation of Raf1- Mitogen-Activated Protein Kinase Kinase-Extracellular Signal-Regulated Kinase 1/2 Pathway. J Pharmacol Exp Ther . 2003;307(3):1213–1220. 92. Bonnerot C, Briken V, Brachet V, et al. syk protein tyrosine kinase regulates Fc receptor γ-chain-mediated transport to lysosomes. The EMBO Journal . 1998;17(16):4606–4616. 93. Majeed M, Caveggion E, Lowell CA, Berton G. Role of Src kinases and Syk in Fcγ receptor-mediated phagocytosis and phagosome-lysosome fusion. J Leukoc Biol . 2001;70(5):801–811. 94. He J, Tohyama Y, Yamamoto K, et al. Lysosome is a primary organelle in B cell receptor-mediated apoptosis: an indispensable role of Syk in lysosomal function. Genes to Cells . 2005;10(1):23–35. 228 95. Lu A, Tebar F, Alvarez-Moya B, et al. A clathrin-dependent pathway leads to KRas signaling on late endosomes en route to lysosomes. J Cell Biol . 2009;184(6):863– 879. 96. Lu Y, Li Y, Seo C-S, et al. Saucerneol D inhibits eicosanoid generation and degranulation through suppression of Syk kinase in mast cells. Food and Chemical Toxicology . 2012;50(12):4382–4388. 97. Lin F-S, Lin C-C, Chien C-S, Luo S-F, Yang C-M. Involvement of p42/p44 MAPK, JNK, and NF-κB in IL-1β-induced ICAM-1 expression in human pulmonary epithelial cells. Journal of Cellular Physiology . 2005;202(2):464–473. 98. Shibata Y, Kamata T, Kimura M, et al. Ras Activation in T Cells Determines the Development of Antigen-Induced Airway Hyperresponsiveness and Eosinophilic Inflammation. J Immunol . 2002;169(4):2134–2140. 99. Duan W, Chan JHP, Wong CH, Leung BP, Wong WSF. Anti-Inflammatory Effects of Mitogen-Activated Protein Kinase Kinase Inhibitor U0126 in an Asthma Mouse Model. J Immunol . 2004;172(11):7053–7059. 100. Okada S, O’Brien JS. Tay-Sachs Disease: Generalized Absence of a Beta-D-N- Acetylhexosaminidase Component. Science . 1969;165(3894):698–700. 101. Renlund M, Tietze F, Gahl WA. Defective sialic acid egress from isolated fibroblast lysosomes of patients with Salla disease. Science . 1986;232(4751):759–762. 102. Platt FM, Neises GR, Reinkensmeier G, et al. Prevention of Lysosomal Storage in Tay-Sachs Mice Treated with N-Butyldeoxynojirimycin. Science . 1997;276(5311):428– 431. 103. Llovet JM, Ricci S, Mazzaferro V, et al. Sorafenib in Advanced Hepatocellular Carcinoma. New England Journal of Medicine . 2008;359(4):378–390. 104. Cheng A-L, Kang Y-K, Chen Z, et al. Efficacy and safety of sorafenib in patients in the Asia-Pacific region with advanced hepatocellular carcinoma: a phase III randomised, double-blind, placebo-controlled trial. Lancet Oncol. 2009;10(1):25–34. 105. Hornsleth A, Loland L, Larsen LB. Cytokines and chemokines in respiratory secretion and severity of disease in infants with respiratory syncytial virus (RSV) infection. Journal of Clinical Virology . 2001;21(2):163 – 170. 106. Kallal LE, Schaller MA, Lindell DM, Lira SA, Lukacs NW. CCL20/CCR6 blockade enhances immunity to RSV by impairing recruitment of DC. European Journal of Immunology . 2010;40(4):1042–1052. 107. Barczyk A, Pierzchala W, Sozanska E. Interleukin-17 in sputum correlates with airway hyperresponsiveness to methacholine. Respir Med . 2003;97:726–33. 108. Kunkel SL, Standiford T, Kasahara K, Strieter RM. Interleukin-8 (IL-8): the major neutrophil chemotactic factor in the lung. Exp. Lung Res. 1991;17(1):17–23. 109. Chintakuntlawar AV, Chodosh J. Chemokine CXCL1/KC and its Receptor CXCR2 Are Responsible for Neutrophil Chemotaxis in Adenoviral Keratitis. Journal of Interferon & Cytokine Research . 2009;29(10):657–666. 229 110. Dufour JH, Dziejman M, Liu MT, et al. IFN-Γ-Inducible Protein 10 (IP-10; CXCL10)-Deficient Mice Reveal a Role for IP-10 in Effector T Cell Generation and Trafficking. J Immunol . 2002;168(7):3195–3204. 111. Osterholzer JJ, Ames T, Polak T, et al. CCR2 and CCR6, but Not Endothelial Selectins, Mediate the Accumulation of Immature Dendritic Cells Within the Lungs of Mice in Response to Particulate Antigen. J Immunol . 2005;175(2):874–883. 112. Berry MA, Hargadon B, Shelley M, et al. Evidence of a Role of Tumor Necrosis Factor α in Refractory Asthma. New England Journal of Medicine . 2006;354(7):697–708. 113. Morjaria JB, Chauhan AJ, Babu KS, et al. The role of a soluble TNFα receptor fusion protein (etanercept) in corticosteroid refractory asthma: a double blind, randomised, placebo controlled trial. Thorax . 2008;63(7):584–591. 230