SOLITARY CHEMOSENSORY CELLS AND CHEMOSENSORY BRUSH

CELLS: CELL-LIKE AIRWAY CHEMOSENSORS

by

CECIL JAMES SAUNDERS

B.S., Lenoir-Rhyne College, 2004

B.A., Lenoir-Rhyne College, 2005

M.S., Wake Forest University, 2008

A thesis submitted to the

Faculty of the Graduate School of the

University of Colorado in partial fulfillment

of the requirements for the degree of

Doctor of Philosophy

Neuroscience Program

2014 ii

This thesis for the Doctor of Philosophy degree by

Cecil James Saunders

has been approved for the

Neuroscience Program

By

Sukumar Vijayaraghavan, Chair

Thomas E. Finger, Advisor

Linda Barlow

Sue Kinnamon

Diego Restrepo

Susan D Reynolds

Date ___01-06-14______iii

Saunders, Cecil James. (Ph.D., Neuroscience)

Solitary Chemosensory Cells and Chemosensory Brush Cells: Taste Cell-like Airway

Chemosensors.

Thesis directed by Professor Thomas E. Finger.

ABSTRACT

The airway is replete with solitary chemosensory cells (SCCs) and brush cells

(BCs) that can trigger inflammation and chemesthesis. These chemosensors utilize the canonical “bitter” taste transduction cascade (T2Rs, αgustducin, TRPM5) to detect bitter substances and bacterial metabolites. Taste cell-like chemosensors form synapses with peptidergic and, when stimulated, release to trigger nociceptors via acetylcholine neurotransmitter release. While SCCs and BCs share many similarities, they have been considered two different cell types. This dissertation examines the signaling of SCCs and replacement of BCs to help determine what differences might exist between these cells. I present evidence that SCCs, similar to BCs, express the synthetic enzyme for acetylcholine, choline acetyltransferase (ChAT).

Additionally, I show that SCC-induced irritation, like BC-induced irritation, is prevented by a nicotinic acetylcholine receptor (nAChR) inhibitor, mecamylamine. Taken together, these results indicate that SCCs have the capacity to synthesize and release acetylcholine upon activation. Furthermore, I also tested the hypothesis that tracheal BCs, like SCCs and other chemosensory cells, are replaced at the same rate as the surrounding epithelium. To determine the rate of BC replacement, I utilized 5-bromo-2’-deoxyuridine

(BrdU) to birthdate cells in the tracheal epithelium in adult and perinatal mice. Although scattered cells in the tracheal epithelium were labeled by BrdU in adult mice, no labeled

BCs were observed. This result is distinct from similar experiments on nasal epithelium, where SCCs were labeled at the same rate as other epithelial cells and raised questions iv about the origin of BCs. BrdU experiments on perinatal mice determined that BCs arise during the period when the trachea is growing. In summary, SCCs and BCs share a similar mechanism by which they induce irritation, but differences exist in the rate at which these cells are replaced. Whether this difference is sufficient to consider SCCs and

BCs different cell types is largely dependent on the criteria utilized to define a cell type.

However, the close functional similarity between SCCs and BCs is strong evidence for all the taste cell-like chemosensors of the airway to be considered the same cell type.

The form and content of this abstract are approved. I recommend its publication.

Approved: Thomas E. Finger

v

ACKNOWLEDGEMENTS

My research would not be possible without the advice, support and dedication of my PhD mentor, Tom Finger. Science is a collaborative process and I have been fortunate to be surrounded by outstanding lab-mates and many excellent scientists at the Rocky

Mountain Taste and Smell Center and the University of Colorado. In particular, I thank

Marco Tizzano, Sue Kinnamon and Sue Reynolds for their counsel and technical advice.

Finally, I extend my gratitude to Wayne Silver (Wake Forest University) for introducing me to the study of chemical and to my parents for their support over the past five years. vi

TABLE OF CONTENTS

CHAPTER

I. INTRODUCTION ...... 1

From Common Chemical to Chemesthesis ...... 2

Molecular Mechanisms of Noxious Chemesthesis ...... 5

Chemesthesis in the Airway ...... 12

Nerves innervating the conductive airways ...... 13

Protective respiratory reflexes and inflammation ...... 14

Taste Cell-like Chemosensors ...... 16

Solitary chemosensory cells ...... 16

Brush cells ...... 20

Nomenclature of Taste Cell-like Chemosensors ...... 23

Organization of Thesis ...... 24

II. CHOLINGERIC NEUROTRANSMISSION LINKS SOLITARY CHEMOSENSORY CELLS TO NASAL INFLAMMATION ...... 25

Abstract ...... 25

Significance Statement...... 26

Introduction ...... 26

Materials and Methods ...... 29

Animals ...... 29

Immunofluorescence ...... 30

Resinoferatoxin ablation of nerve terminals ...... 31

Plasma extravasation ...... 31

Mast cell degranulation ...... 32

Pharmacology ...... 33

Results ...... 33 vii

SCCs are cholinergic and contact peptidergic nociceptors ...... 33

Gustducin and TRPM5 are required for SCC mediated inflammation ...... 35

Peptidergic nociceptive trigeminal fibers are required for SCC-mediated inflammation ...... 36

SCC-mediated inflammation requires nicotinic ACh receptors (nAChRs) ...... 39

NK1 receptors underlie both plasma extravasation and mast cell degranulation ...... 42

Discussion ...... 43

Parallel pathways for airway irritation ...... 43

Mast cells—a node in the inflammatory signaling cascade ..... 47

SCCs, one component of the airway chemofensor complex ... 48

SCC over-stimulation—a possible pathology for non-allergic (idiopathic) rhinitis? ...... 48

Summary ...... 49

Acknowledgements ...... 50

III. CHEMOSENSORY BRUSH CELLS OF THE TRACHEA: A STABLE POPULATION IN A DYNAMIC EPITHELIUM ...... 51

Abstract ...... 51

Introduction ...... 52

Materials and Methods ...... 53

In vivo experiments ...... 54

Cell counts ...... 57

In vitro experiments ...... 58

Results ...... 60

Identification of chemosensory brush cells...... 60

Chemosensory brush cells do not turnover in adult mice ...... 64 viii

Perinatal generation of chemosensory brush cells ...... 64

New chemosensory brush cells can be generated from adult tracheal epithelial progenitors ...... 69

Discussion ...... 73

Tracheal brush cells are a stable population ...... 73

New brush cells are generated during tracheal growth ...... 75

Reliable markers for chemosensory brush cells...... 77

Conclusions ...... 78

Acknowledgements ...... 78

Grants ...... 79

IV. CONCLUSION ...... 80

Are Sccs And Bcs The Same Thing? ...... 81

Morphology and anatomy ...... 82

Physiology and function ...... 83

Development ...... 83

Molecular character ...... 85

Paracrine Signaling ...... 86

The Airway Chemofensor Complex ...... 88

Concluding Thoughts ...... 90

REFERENCES ...... 92 ix

TABLES

Table

1.1 Several types of nerve fibers innervate epithelial tissues ...... 4

1.2 Agonists of thermal TRPs ...... 6

x

FIGURES

Figure

1.1 A Solitary Chemosensory Cell and associated nerve fibers...... 17

1.2 Brush Cells of the trachea...... 22

2.1 Cross sections of nasal epithelium showing cellular properties and relationships of SCCs...... 34

2.2 Stimulation of SCCs activates a pro-inflammatory pathway that triggers plasma extravasation...... 37

2.3 Stimulation of SCCs activates a pro-inflammatory pathway that triggers mast cell degranulation ...... 40

2.4 Parallel pathways for airway irritation ...... 45

3.1 Diagram of BrdU treatment and culture methods ...... 59

3.2 BCs are a distinct cell type in the tracheal epithelium (Part 1) ...... 61

3.3 BCs are a distinct cell type in the tracheal epithelium (Part 2)...... 63

3.4 BCs are a static population in the adult trachea but are generated by a population of proliferative progenitor cells during perinatal development. .... 66

3.5 Chemosensory brush cells are present in the lower airways of prenatal mice ...... 68 3.6 New BCs can be generated from adult tracheal epithelium in vitro ...... 71

1

CHAPTER I

INTRODUCTION

Epithelial tissues are complex sensory end organs, not just protective coverings.

The primary function of an epithelium is to guard an organism from its external environment (Nestle, Di Meglio et al. 2009). One of the ways epithelial layers accomplish this defensive function is to exclude the external environment from the internal domain (Nestle, Di Meglio et al. 2009). However, organisms are not closed systems, and epithelial layers covering bodily orifices must be highly specialized to monitor the external world as organisms seek out and ingest the various chemicals necessary to maintain life (Boulais and Misery 2008). To accomplish their role as sensory end organs, epithelial layers are highly innervated by nerve fibers (Bryant and Silver

2000, Lumpkin and Caterina 2007). However, these sensory nerve fibers are sensitive to damage and thus also demand protection by epithelial layers.

The sensory function of epithelium is often at odds with its role as a protective barrier (Boulais and Misery 2008). Sensory nerve fibers innervating epithelial tissues generally extend into and between epithelial cells but do not pass through the protective barrier into the external world (Cauna, Hinderer et al. 1969, Mack, Anand et al. 2005,

Lumpkin and Caterina 2007, Boulais and Misery 2008). The integumentary system of and other land animals consists of epithelial cells that are specialized to act as a barrier. To accomplish this goal, epithelial cells in skin often produce protective structures composed of keratins (Fuchs 1995, Mack, Anand et al. 2005), which have the double effect of preventing chemicals and many forms of energy from passing through the barrier to stimulate the underlying nerves (Bryant and Silver 2000, Boulais and

Misery 2008, Silver, Roe et al. 2010). In contrast to the integumentary system, organ systems where absorption or excretion occur cannot be composed of cornified or

2 keratinized epithelial tissues (Bragulla and Homberger 2009). Additionally, a cornified keratinized epithelium would limit the sensory capabilities of these tissues and these mucus membranes must monitor xenobiotics and noxious chemicals (Bragulla and

Homberger 2009).

The epithelial layers covering the mucous membranes are particularly sensitive to

th stimulation by noxious chemicals. Throughout the 20 century, most research on chemical irritation focused on the nerves innervating the mucous membranes (Bryant and

Silver 2000). The assumption among those studying the chemical senses was that only chemicals with certain physical and chemical properties, specifically high lipophilicity and volatility, are capable of diffusing through the muco-ciliary epithelium to activate the

st underlying nerve fibers (Bryant and Silver 2000). However, in the early 21 century,

Finger et al. and Sbarbati et al. identified scattered neuro-sensory epithelial cells throughout the respiratory epithelium of rodents (Finger, Bottger et al. 2003, Sbarbati,

Merigo et al. 2004). Since the initial discovery of these solitary chemosensory cells

(SCCs), further work has firmly established SCCs as sensory end organs capable of activating the nociceptive nerve fibers, which innervate the respiratory epithelium.

This introduction begins with a general sketch of the neuroscience underlying chemical irritation, before exploring the function of solitary chemosensory cells in respiratory epithelium.

From Common Chemical Sense to Chemesthesis

The study of chemical irritation in the modern era begins with experiments that led G.H. Parker to create the term “common chemical sense” (Bryant and Silver

2000). In one experiment, Parker observed that fish were able to detect some noxious chemicals even after the nerves innervating the gustatory and had been severed (Parker 1912). Specifically, these fish writhed in response to acidic, alkaline and

3 high salt solutions (Parker 1922). Parker postulated that free nerve endings which innervated various epithelial tissues were the nociceptors responsible for detecting these chemical irritants (Parker 1922). Since sensations of chemical irritation appeared to be common to all epithelial tissues, he referred to this sensation as a “common chemical sense.” Thus he distinguished the common chemical sense from the chemical senses of gustation and olfaction (Bryant and Silver 2000). Parker’s basic assumption that free nerve fibers in the epithelium are responsible for mediating sensations of chemical irritation has endured. However, Parker was unable to determine the exact characteristics of the nociceptors that mediate the common chemical sense.

Researchers following up on the work of G.H. Parker mistakenly redefined the common chemical sense to be specific to the mucosal epithelium (Keele 1962). This misstep was a result of cornified skin of terrestrial vertebrates protecting peptidergic nociceptors from chemical irritants (Bryant and Silver 2000, Bragulla and Homberger

2009). However, if the protective layer of keratin is breached (Keele 1962, Basbaum,

Bautista et al. 2009) or if the chemical irritant can diffuse through the keratinized layer

(Green and Flammer 1989), then the sensation produced is similar to when the chemical is presented to a mucus membrane (Frasnelli, Heilmann et al. 2004). To be irritating, a chemical must be able to stimulate somatosensory fibers either directly or indirectly through the epithelial layers.

Epithelial tissues are innervated by different types of somatosensory nerve fibers, including nociceptive fibers. Most Somatosensory nerve fibers originate in either the trigeminal, vagal or dorsal root ganglia (Bryant and Silver 2000, Lumpkin and Caterina

2007). These nerve fibers are classified into categories based on their axon diameter, conduction speed and the presence of a myelin sheath (Julius and Basbaum 2001). Each fiber type conducts tactile, temperature and/or sensations. Three different classes of

4 nerve fibers typically innervate epithelial tissues (Table 1.1) but only the thinnest myelinated (Aδ-fibers) and un-myelinated (C-fibers) are responsible for transducing noxious sensations (Bryant and Silver 2000, Julius and Basbaum 2001).

Sensations of chemical irritation are transduced by a subset of these nociceptive fibers and can be identified based on the expression of the inflammatory peptides, Substance P and Calcitonin Gene Related Peptide (CGRP) (Lundblad, Lundberg et al. 1983,

Lundberg, Brodin et al. 1984). These peptidergic nociceptive fibers are found throughout both keratinized and mucosal epithelia tissues but are more easily accessed by chemical irritants in mucosal epithelium (Basbaum, Bautista et al. 2009).

Table 1.1: Several types of nerve fibers innervate epithelial tissues. This table summarizes information from Julius & Basbaum, 2001, and Bryant & Silver, 2000. Conduction Sensation Myelin Diameter Velocity Aβ Mechanoreception Yes 6 to 12 µm 35 to 75 m/sec Mechanoreception, Aδ tempature and/or pain Yes 1 to 5 µm 5 to 30 m/sec C Pain and temperature No 0.2 to 1.5 µm 0.5 to 2 m/sec

As research on chemical irritation progressed in the 20th century, it became clear that chemical irritation can occur anywhere that thermal-sensitive or nociceptive somatosensory nerve fibers are present (Hensel and Zotterman 1951, Armstrong, Dry et al. 1953, Van Hees and Gybels 1972, Szolcsanyi 1977, Green 1985, Schafer, Braun et al.

1986). These nociceptive fibers are traditionally thought to be as part of the , but their stimulation by chemical irritants was generally considered to be different from any accepted definition of somatosensation (Green,

Mason et al. 1990). Similarly, G.H. Parker coined the term “common chemical sense” to refer to a distinct sensory capacity and chemical irritation was a function that was mediated by the somatosensory system―not a separate sense. Thus, a new term was needed that accurately referred to the chemical stimulation of the somatosensory system.

5

At a symposium on irritation in 1988, some researchers suggested that the neologism chemesthesis be used to refer to sensations caused by chemical stimulation of the somatosensory system (Green, Mason et al. 1990). The word chemesthesis is a portmanteau of chemical and somesthesis (Green, Mason et al. 1990). Somesthesis refers to the sensations produced when the somatosensory system is stimulated by kinetic or thermal energy. Thus, chemesthesis refers to sensations produced when the somatosensory system is activated by chemical stimuli (Green, Mason et al. 1990). The term “chemesthesis” is now widely accepted among the chemical senses community as a replacement for the less accurate term “common chemical sense” (Bryant and

Silver 2000, Green 2012).

While chemesthesis is a more specific term for the sensations associated with chemical irritation, the two terms are not synonymous. When Parker coined the term common chemical sense”, he was mainly concerned with an organism’s ability to detect noxious chemicals. However, somatosensory fibers, which conduct sensations other than , can also be stimulated directly by certain chemicals. Specifically, chemicals have been identified that stimulate itch (Wilson, Nelson et al. 2013) and vibration (Bautista, Sigal et al. 2008) transducing nerve fibers. Chemesthesis also includes this activation of all somatosensory fiber types by chemicals―not just the stimulation of peptidergic nociceptors.

Molecular Mechanisms of Noxious Chemesthesis

Noxious chemesthestic stimuli typically activate receptors found on the free peptidergic nociceptive nerve endings innervating epithelial tissues. Our understanding of chemesthesis has greatly increased by identifying what receptors are activated by

6

7 chemicals long known to be irritating (Silver, Roe et al. 2010). Specifically, the receptors responsible for allowing nociceptive nerve fibers to respond to chemicals found in chilies, mustards, mint and numerous other naturally-occurring compounds have all been identified (Gerhold and Bautista 2009) (Table 1.2). Many of these proteins belong to the transient receptor potential (TRP) channel superfamily.

TRP channels are a family of non-specific cation channels involved in transducing several different sensory modalities. The first TRP channel was discovered in the drosophila , where genetic deletion of the channel resulted in a transient response to light (Cosens and Manning 1969). All TRP channels have six transmembrane domains with the fifth and sixth domains forming the cation pore (Zheng 2013). TRP channels often form in hetro-tetramers with other TRP channels from the same sub- family (Song and Yuan 2010). There are seven sub-families of TRP channels identified as follows: canonical (C), polycystic (P), mucolipin (ML), no potential

(N), melastatin (M), ankyrin (A) and vanilloid (V) (Song and Yuan 2010). TRP channels from many of these sub-families play an essential role in a variety of mammalian sensory systems ranging from somatosensation to gustation (Zheng 2013). Several members from the melastatin and vanilloid group also play an essential role in thermal sensation

(Bandell, Macpherson et al. 2007). However, members of the melastatin, ankyrin and vanilloid TRP sub-families are closely associated with the detection of chemical irritants.

TRP vanilloid 1 (TRPV1) is responsible for detecting capsaicin, the irritating chemical found in chilies. For decades, researchers had utilized capsaicin experimentally to activate nociceptors without knowing its exact mechanism of action (Szolcsanyi 2004).

Eventually, TRPV1 was identified as the receptor for capsaicin and shown to mediate sensitivity to a number of other noxious stimuli, including low pH and temperatures above 43°C (Caterina, Schumacher et al. 1997). Previous physiological research

8 established that many peptidergic c-fibers respond to a variety of noxious stimuli and these fibers were labeled polymodal nociceptors for their broad response profiles (Beitel and Dubner 1976) (Tominaga, Caterina et al. 1998). Since TRPV1 is expressed by a subset of c-fibers that innervate most epithelial tissues, it provided a single receptor by which disparate stimuli could activate the nerve fibers that encode for sensations of burning pain (Tominaga, Caterina et al. 1998). However, polymodal nociceptive fibers also express other receptors that could contribute to their polymodal nature.

Shortly after the discovery of TRPV1, another receptor was identified on peptidergic nociceptors, TRP ankyrin 1 (TRPA1). Where TRPV1 responds to a chemical found in chilies, TRPA1 responds to pungent compounds found in a variety of plants including cinnamon, mustards, onions and garlic (Bautista, Movahed et al. 2005).

Additionally, TRPA1 is also activated by many volatile organic compounds and weak

Lewis acids (Wang, Chang et al. 2011). While TRPA1 appears to respond to a diverse collection of chemical stimuli, in reality cinnamaldehyde from cinnamon bark, isothiocyanates from mustards, sulfides from onions and garlic, and volatile organic compounds like acrolein all act on TRPA1 via the same mechanism, electrophilic attack

(Hinman, Chuang et al. 2006, Macpherson, Xiao et al. 2007). Essentially, TRPA1 is a detector of chemical reactivity, binding strongly electrophilic compounds to cysteine residues and warning of the presence of dangerous reactive chemicals (Hinman, Chuang et al. 2006). In contrast to TRPA1, which mainly responds to one class of chemical,

TRPV1 is truly polymodal, responding to thermal energy, low pH and plant metabolites

(Tominaga, Caterina et al. 1998). However, both TRPV1 and TRPA1 contribute to the polymodal nature of some peptidergic nociceptors.

TRPA1 is almost always co-expressed with TRPV1 on peptidergic nociceptive fibers (Kobayashi, Fukuoka et al. 2005). A subset of these TRPV1- and TRPA1-

9 expressing peptidergic fibers also conduct sensations of noxious itching in addition to conducting sensations of pain (Wilson, Nelson et al. 2013). Furthermore, both TRPV1 and TRPA1 are essential members in the signaling cascades responsible for chemesthetic itch, although direct stimulation of either receptor seldom results in these sensations

(Wilson, Nelson et al. 2013). This seeming contradiction, that TRPA1 and TRPV1 are involved in the of both pain and itch but direct activation of either channel only produces the sensation of pain, is resolved by examining the central projections of

“itch-sensitive” and “pain-sensitive” peptidergic nociceptors. The “pain- sensitive” nociceptive neurons activate a that results in the inhibition of inputs from the “itch-sensitive” nociceptive neurons (Ikoma, Steinhoff et al. 2006).

Thus, pain inhibits itch and if TRPV1 or TRPA1 is stimulated in both “pain” and

“itch” nociceptors, then burning pain is the dominate sensation.

The nerve fibers that conduct sensations for burning pain express TRPV1 and sometimes TRPA1, while the nerve fibers responsible for conducting sensations of noxious cold express TRP melastatin 8 (TRPM8) (Bandell, Macpherson et al. 2007).

TRPM8 is activated by temperatures below 28°C, menthol found in mint leaves and the synthetic agonist icilin (McKemy, Neuhausser et al. 2002, Peier, Moqrich et al. 2002). In contrast to TRPV1 and TRPA1, which are activated by strong acids and weak acids, respectively, TRPM8’s activity is inhibited by low pH (McCoy, Knowlton et al. 2011).

TRPM8 is expressed in both C- and Aδ-fibers but is not typically co-expressed in nerve fibers with either TRPV1 or TRPA1 (Abe, Hosokawa et al. 2005, Kobayashi, Fukuoka et al. 2005). However, occasional exceptions have been observed (Dhaka, Earley et al.

2008). Additionally, TRPM8-expressing nerve fibers are not immunoreactive for the inflammatory neuropeptides substance P and CGRP (Dhaka, Earley et al. 2008). TRPM8-

10 expressing nociceptors are primarily responsible for detecting sensations of noxious cold; however, there have also been several reports of TRPA1 being activated by cold temperatures (Bandell, Story et al. 2004, del Camino, Murphy et al. 2010). Currently, most of these reports are considered to be an artifact of studying TRPA1 in heterologous expression systems, as knockout mice for TRPA1 respond normally to noxious cold

(Caspani and Heppenstall 2009). TRPM8, like TRPV1 and TRPA1, is expressed in nerve fibers that conduct sensations of temperature and can be activated by a variety of chemesthestic stimuli; however, not all thermal sensitive TRP channels are expressed in nerve fibers.

TRP vanilloid 3 (TRPV3) and TRP vanilloid 4 (TRPV4) are expressed in keratinocytes in the skin, but not in cutaneous nerve fibers. TRPV3 and TRPV4 are both activated by warm temperatures between 22 and 40°C (Liedtke, Choe et al. 2000, Peier,

Reeve et al. 2002, Smith, Gunthorpe et al. 2002). While TRPV3 and TRPV4 have similar temperature-response profiles, these channels respond to very different chemical stimuli.

TRPV3 can be activated by eugenol in cloves, thymol in thyme and carvacol in oregano

(Xu, Delling et al. 2006). TRPV4, on the other hand, responds to the botanically-derived carcinogens phorbol esters and hypotonic stress (Vriens, Watanabe et al. 2004).

Keratinocytes that express TRPV3 and TRPV4 demonstrate heat-evoked currents, suggesting that they are thermal sensors (Chung, Lee et al. 2004). Additionally, keratinocytes release ATP when exposed to warm temperatures, suggesting that these cells might signal warming temperatures to other cells in the epithelium, including nerve fibers (Souslova, Cesare et al. 2000, Mandadi, Sokabe et al. 2009). In many respects, keratinocytes acting as epithelial sensors signaling to the underlying nerve fibers parallels the role of solitary chemosensory cells in the respiratory epithelium. However, with these

11 two exceptions, most chemesthesitc stimuli act directly on receptors expressed by nerve fibers.

The discovery of TRP channels has been a boon to our understanding of how chemicals stimulate the somatosensory system; however, other receptors can mediate chemesthetic stimulation. Many peptidergic nociceptors also express acid-sensing ion channels (ASIC) that, like TRPV1 and TRPA1, can detect low pH (Olson, Riedl et al.

1998). In a subset of TRPV1- and TRPA1-expressing neurons, the compound chloroquine can induce itch by acting on the G-protein coupled receptor MrgprA3 (Wilson, Nelson et al. 2013). In addition to nerve fibers conducting nociceptive sensations, fibers that detect vibration can also be activated by chemicals like hydroxy-α-sanshool from szechuan peppers (Bautista, Sigal et al. 2008). These vibration-sensitive nerves express potassium channels (KCNK3, KCNK9 and KCNK18) that are forced to open by hydroxy-α- sanshool causing them to be activated (Bautista, Sigal et al. 2008). Finally, numerous neurotransmitter receptors are also found on sensory neurons; specifically, receptors for

ATP, serotonin and acetylcholine are all expressed by peptidergic nociceptors (Xiang, Bo et al. 1998, Alimohammadi and Silver 2000, Ivanusic, Kwok et al. 2011). Like TRPV1,

TRPA1 and TRPM8, some of these neurotransmitter receptors can be activated by botanically-derived compounds. For example, nicotinic acetylcholine receptors expressed by nociceptive nerve fibers are stimulated by nicotine synthesized in plants such as tobacco (Tso 2000). A diverse collection of receptors are involved in detecting different chemesthestic stimuli that produce a variety of sensations.

While chemesthestic chemicals can produce sensations from vibration to itching, these chemicals are most often associated with the sensations of irritation. This association is largely due to the polymodal nature of peptidergic nociceptors, which allows these nerve fibers to respond to multiple classes of chemicals (Bryant and Silver

12

2000, Silver, Roe et al. 2010). However, regardless of the receptors that are activated, the result of stimulating these nociceptors is generally the same―pain and inflammation

(Julius and Basbaum 2001). The pain is a result of the central projection of the nociceptive fibers that extend into brain and spinal cord regions coding for painful sensation (Basbaum, Bautista et al. 2009). Inflammation is primarily a result of the local release of inflammatory mediators, such as substance P and CGRP, from peptidergic nociceptors (O'Connor, O'Connell et al. 2004). While pain and inflammation due to chemesthesis occur anywhere peptidergic nociceptors innervate, chemesthesis is most often studied in mucosal epithelium, as opposed to skin where cornification of the epithelial layer protects the underlying nerve fibers from most noxious chemicals.

Chemesthesis in the airway

The mucus membranes of the airway are irritated by noxious chemicals. While the mucus membranes of the gastrointestinal tract, reproductive tract and urinary system are also very sensitive to chemesthetic stimuli, the respiratory system is arguably the most common site for a terrestrial animal to experience chemical irritation (Bryant and

Silver 2000). Additionally, several reflexive modifications of respiration are associated and intertwined with chemesthetic irritation in the respiratory tract. For these reasons, researchers interested in chemesthesis have often focused upon chemical irritation of the respiratory tract.

The airway can be divided anatomically into the respiratory zone and conducting zone. The respiratory zone is defined as the tissues of the lung where gas exchange occurs. To facilitate gas exchange, the tissues of the respiratory zone are thin, fragile and particularly sensitive to noxious chemicals (Canning and Spina 2009). Conversely, the conducting zone consists of the nasal cavity, nasopharynx, larynx, trachea, bronchi and bronchioles, and is more durable than the tissue of the respiratory zone (Canning and

13

Spina 2009). As such, the main functions of the conducting airways are to condition the air as it is passes into the lower lungs and to monitor the air for dangerous chemicals

(Harkema, Carey et al. 2006).

Nerves innervating the conductive airways

Different sections of the conductive airways are innervated by peptidergic nociceptors that originate from different . The most rostral portion of the conductive airway, the nasal cavity, is innervated by the . The pharynx connects the nasal cavity to the larynx and is innervated by the glossopharyngeal nerve.

Finally, the larynx, trachea, bronchi and bronchioles are innervated by the vagus nerve.

While the trigeminal, glossopharyngeal and vagus nerves all provide somatosensory innervation to the conducting airways, these nerves are not purely sensory nerves.

The trigeminal nerve is responsible for providing somatosensory innervation to the face, head and nasal cavity (Bathla and Hegde 2013). The nasal cavity is innervated by the ophthalmic and maxillary branches of the trigeminal nerve (Silver, Roe et al.

2010). In addition to the nasal cavity, these branches of the trigeminal nerve also provide somatosensory innervation to the cornea, the oral cavity and the upper two-thirds of the head (Bryant and Silver 2000). The ophthalmic and maxillary branches of the trigeminal nerve are composed entirely of sensory nerve fibers (Bathla and Hegde 2013). In contrast, the mandibular branch of the trigeminal nerve contains efferent motor nerve fibers that are responsible for controlling the muscle of mastication, as well as somatosensory afferents that innervate parts of the oral cavity and the bottom third of the face (Bathla and Hegde 2013).

Like the trigeminal nerve, the glossopharyngeal nerve has both a sensory and a motor function. The pharynx and posterior third of the tongue are innervated by somatosensory fibers of the glossopharyngeal nerve. In addition to conducting

14 somatosensory information, branches of the glossopharyngeal nerve conduct chemosensory information from carotid body chemosensors and taste buds on the posterior third of the tongue. Efferent fibers from the glossopharyngeal nerve provide parasympathetic innervation to salivary glands and motor innervation to some muscles in the pharynx.

Like the glosssopharyngeal nerve, the vagus nerve is a mixed nerve consisting of both afferent and efferent nerve fibers. The vagus nerve extends both afferent somatosensory fibers and efferent autonomic fibers throughout much of the viscera including the lower airways (Barratt 1898). Efferent or autonomic motor innervation from the vagus nerve extends to the heart, adrenal glands, gastrointestinal tract and larynx (Ruffoli, Giorgi et al. 2011). Sensory fibers from the vagus project to the gastrointestinal tract and carotid body chemosensors, as well as the lower airways (Muroi and Undem 2011). Peptidergic nociceptors from the trigeminal glossopharyngeal or vagus nerves can be responsible for detecting chemesthetic stimuli along the airway.

Protective respiratory reflexes and inflammation

Stimulation of trigeminal and vagal peptidergic nociceptors also provokes protective respiratory reflexes. These reflexes can range from sneezing to coughing to patterned apnea (Alarie 1966). Coughing and sneezing are mainly mediated by acute stimulation of the vagus nerve and result in a forceful exhalation which clears the airway

(Undem and Carr 2001, Widdicombe 2001, Carr and Undem 2003). In contrast to coughing and sneezing, apnea can occur when either trigeminal or vagal nociceptors are stimulated and is thought to limit the exposure of the lower airway to noxious chemicals by slowing down overall respiration rate (Alarie 1973).

The patterned apnea caused by airway irritation is used as an assay to measure airway irritation. Stimulation of trigeminal and vagal nociceptors results in a reflexive

15 closing of the glottis during the expiratory phase of the respiratory cycle

(Vijayaraghavan, Schaper et al. 1993). Closing of the glottis during exhalation causes a pause between breaths and lowers the respiration rate during periods of trigeminal or vagal irritation (Symanowicz, Gianutsos et al. 2004). The degree of apnea directly correlates to the amount of irritation experienced―more irritating stimuli produce longer periods of apnea, resulting in a lower respiration rate (Alarie 1973, Alarie 1973).

Expiratory pause assays have been used to study respiratory tract irritation for decades

(Alarie 1966, Kane, Barrow et al. 1979, Nielsen 1991) and were essential in determining the role of solitary chemosensory cells in detecting “bitter” irritants (Finger, Bottger et al. 2003, Sbarbati and Osculati 2006, Tizzano, Gulbransen et al. 2010).

Stimulation of trigeminal and vagal peptidergic nociceptors by irritants also results in location inflammation, in addition to protective respiratory reflexes. The signs of inflammation are classically defined as the presence of pain (dolor), the generation of heat (calor), the presence of redness (rubor) and swelling (tumor) of tissues (Rather 1971,

Lawrence, Willoughby et al. 2002). Inflammation is caused by inflammatory peptides released from the nerve terminals of stimulated nociceptors (Rodger, Tousignant et al.

1995). When activated, peptidergic nociceptive fibers release neuropetides bi- directionally, both at the site of stimulation and from axonal projections downstream of the stimulation, through a phenomenon called the axon reflex (Langley 1900, Finger, St

Jeor et al. 1990). The release of inflammatory neuropeptides, such as substance P, sensitizes both neighboring nociceptive fibers and neurons in the resulting in the hypersensitivity to pain and the warming sensation characteristic of inflammation (O'Connor, O'Connell et al. 2004).

Locally, the release of substance P and other neurokinin peptides acts on blood vessels to create the signs of inflammation (Rodger, Tousignant et al. 1995). Specifically,

16 neurokinins cause the endothelial cells, which compose vessel walls, to separate, allowing blood plasma to extravasate into the extracellular space (Rodger, Tousignant et al. 1995). The leakage of blood results in the inflammatory edema and reddening of the tissue (Rather 1971, Lawrence, Willoughby et al. 2002). Neurokinins, like substance P, are important integrators for inflammatory signaling, as they are released from peptidergic nociceptors causing inflammation without regard to the mechanism of stimulation.

Taste Cell-like Chemosensors

Solitary chemosensory cells

The discovery and identification of solitary chemosensory cells (SCCs) in the mammalian airway fundamentally altered the paradigm of chemesthesis and elucidated a new mechanism by which chemical irritation could occur (Finger, Bottger et al. 2003,

Sbarbati, Merigo et al. 2004). SCCs are neuro-epithelial cells in the respiratory epithelium, which are exposed directly to chemicals in the lumen of the airway (Finger,

Bottger et al. 2003) (Figure 1.1). SCCs allow the detection of irritants that cannot normally penetrate the respiratory epithelium to directly stimulate nerve fibers (Finger,

Bottger et al. 2003). These sentinels of the airway resemble taste receptors cells in morphology and molecular character but are found throughout in the respiratory epithelium of the nasal cavity.

SCCs bear a morphological resemblance to cells but exist as scattered cells within the nasal epithelium. SCCs are bottle-shaped cells, with a cell body that typically resides in the bottom half of pseudostratified muco-ciliary epithelium and one or more microvillar processes that reach though to the lumen of the epithelial layer

(Finger, Bottger et al. 2003). SCCs form occasional clusters throughout the respiratory

17 epithelium, but these clusters do not resemble taste buds (Gulbransen and Finger 2005,

Gulbransen 2007), and it is more common for SCCs to exist surrounded by the other cell types of the respiratory epithelium (Gulbransen and Finger 2005, Gulbransen 2007). In contrast to taste receptor cells, which are innervated by non-peptidergic sensory nerve fibers from the facial or glossopharyngeal nerves (Finger, Danilova et al. 2005), SCCs are intimately intertwined with peptidergic nociceptors arising from the trigeminal or vagal ganglia (Tizzano, Gulbransen et al. 2010). Also in contrast to type II taste receptor cells, which do not form traditional synapses, SCCs appear to form conventional synapses with these nociceptors on the ultrastructure level (Finger, Bottger et al. 2003). While there are several anatomical differences between type II taste receptor cells and SCCs, there are also many molecular and physiological similarities.

Figure 1.1: A Solitary Chemosensory Cell and associated nerve fibers. Section of nasal epithelium from a TRPM5-GFP mouse containing a solitary chemosensory cell (orange) intimately associated with CGRP immunoreactive nerve fibers (magenta). Nuclei are shown in green.

18

SCCs utilize molecules associated with the canonical “bitter” taste transduction cascade to detect potentially noxious compounds in the airway. Nasal SCCs produce mRNA for the bitter receptors mT2R8 and mT2R19 (Finger, Bottger et al. 2003).

Additionally, SCCs are immunoreactive for many of the same proteins that type II taste receptor cells utilize to transduce bitter stimuli: G-protein α-gustducin, phospholipase C

β2 (PLCβ2), inositol trisphosphate receptor type 3 (IP3R3) and the cation channel TRP melastatin 5 (TRPM5) (Finger, Bottger et al. 2003, Tizzano, Gulbransen et al. 2010).

Isolated SCCs stimulated with the prototypical bitter compound denatonium respond with an increase in intracellular calcium, which is blocked by inhibition of PLCβ2

(Gulbransen, Clapp et al. 2008, Tizzano, Gulbransen et al. 2010). Presumably activation of the canonical “bitter” taste transduction cascade leads to a release of a previously unidentified neurotransmitter onto peptidergic nociceptors, causing nasal irritation.

Stimulation of nasal SCCs results in irritation. When bitter substances, such as denatonium, quinine and cycloheximide, pass through the nasal cavity of rats, an increase in activity can be recorded from the ethmoid branch of the trigeminal nerve, which innervates the rostral section of the nasal cavity where many SCCs reside (Finger,

Bottger et al. 2003). Additionally, denatonium, quinine or cycloheximide passing through the nose induces an expiratory pause reflex that is indicative of respiratory tract irritation

(Finger, Bottger et al. 2003). However, in mice with a genetic deletion of either α- gustducin or TRPM5, denatonium stimulation does not induce an expiratory pause

(Tizzano, Gulbransen et al. 2010). Taken together, these findings suggest that SCCs utilize the canonical taste transduction signaling cascade to detect a class of noxious compounds that would not stimulate airway peptidergic nociceptors directly.

19

SCCs respond to several noxious compounds; however, many of these compounds are not commonly found in the airway. For example, denatonium is a purely synthetic compound that is manufactured commercially as an aversive tastant (Klein-

Schwartz 1991). While quinine occurs naturally in the bark of several tree species, it is difficult to imagine a scenario in which tree bark would pose a threat to a mammal’s respiratory tract (Flückiger and Hanbury 1874). Similarly, cycloheximide is an antibiotic agent produced by the bacteria Streptomyces griseus and would not typically invade the airway (Antony-Babu and Goodfellow 2008). However, some respiratory tract microorganisms do produce chemicals that are structurally related to these bitter irritants.

The airways are continually under assault from microorganisms, including commensal bacteria which can become pathogenic under some conditions. Pseudomonas aeruginosa is a germ-negative bacterium that is an opportunistic respiratory pathogen

(Folkesson, Jelsbak et al. 2012). Infections of P. aeruginosa can be fatal under certain conditions (Brewer, Wunderink et al. 1996, Van Delden and Iglewski 1998), particularly once the chemical signals that indicate a high density of the bacteria are present (Fuqua,

Winans et al. 1994). P. aeruginosa accomplishes this bacterium-to-bacterium chemical communication, a phenomenon called quorum sensing, by producing acyl-homoserine lactones (AHLs) (Fuqua, Winans et al. 1994, Fuqua and Greenberg 2002).

AHLs bear a structural resemblance to other bitter compounds. The general structure of AHLs consists of cyclic ethers or lactones, bonded to the amino acid homoserine (Eberhard, Burlingame et al. 1981). Many naturally occurring bitter compounds are lactones and lactone derivatives (Vanbeek, Maas et al. 1990, Seo, Yang et al. 2009), including a class of bitter compounds called furanones which directly resemble

AHLs (Sbarbati and Osculati 2006). Thus, AHLs are a potentially bitter compound that

20 would be present naturally in the respiratory tract where the chemical could come in contact with SCCs.

AHLs activate SCCs in a similar fashion to other bitter compounds. When isolated SCCs are exposed to either 3-oxo-C6-homoserine lactone (HSL) or 3-oxo-C12-

HSL, they show an increase in intracellular calcium concentration, similar to stimulation with denatonium (Gulbransen, Clapp et al. 2008, Tizzano, Gulbransen et al. 2010). This increase in intracellular calcium is PLCβ2-dependent, suggesting that these AHLs are activating the canonical taste transduction cascade (Tizzano, Gulbransen et al. 2010).

Furthermore, when these AHLs are passed though the nasal cavity of mice, they induce the expiratory pause reflex, indicating that these compounds are irritating (Tizzano,

Gulbransen et al. 2010). Additionally, knockout mice for α-gustducin or TRPM5 do not show an expiratory pause reflex when exposed to these lactones, demonstrating that

SCCs are responsible for the detection of these compounds (Tizzano, Gulbransen et al.

2010). Thus, AHLs are the naturally-occurring consequence of P. aeruginosa infection of the airway and can be detected by SCCs.

Brush cells

SCCs are not the only taste cell-like chemosensor found along the respiratory tract. Brush cells were first identified in the respiratory epithelium of the trachea by electron microscopy in the 1950s (Rhodin and Dalhamn 1956). Immediately, these cells were ascribed a sensory function on the basis of their contact with nerve fibers (Rhodin and Dalhamn 1956). However, in the half-century between their discovery and the elucidation of their sensory function (Krasteva, Canning et al. 2011), these cells were often maligned as “a cell in search of a function”(Reid, Meyrick et al. 2005), but after

SCCs were discovered it became clear that brush cells had a similar chemosensory role.

21

Brush cells (BCs) share a number of morphological and molecular characteristics with SCCs. Both SCCs and BCs are bottle-shaped cells that send microvillar processes into the lumen of the respiratory tract (Figure 1.2) (Krasteva, Canning et al. 2011,

Krasteva, Hartmann et al. 2012). Like SCCs, BCs are immunoreactive for elements of the canonical bitter taste transduction cascade, including α-gustducin, PLCβ2, IP3R3 and

TRPM5 (Krasteva, Canning et al. 2011, Krasteva, Hartmann et al. 2012). While both

SCCs and BCs produce mRNA for members of several bitter taste receptors, the two cell types express different bitter taste receptors (Finger, Bottger et al. 2003, Krasteva,

Canning et al. 2011). Additionally, while SCCs and BCs are both innervated by peptidergic nociceptive fibers, the nature of the innervation is distinct (Tizzano,

Cristofoletti et al. 2011). SCCs are repeatedly enwrapped by nociceptive fibers originating in the trigeminal ganglia often making repeated contact with the nerve fiber

(Tizzano, Cristofoletti et al. 2011). In contrast, the vagal nociceptors that form synapses with BCs make only glancing contact with the chemosensory cells (Krasteva, Canning et al. 2011).

Despite the difference between BC and SCC nociceptive innervation, both cell types are capable of detecting “bitter” irritants. Cycloheximide placed on the tracheal epithelium induces protective respiratory reflexes typical of airway irritation (Krasteva,

Canning et al. 2011). Additionally, stimulation of the tracheal epithelium with AHLs also results in reflexes associated with irritation (Krasteva, Canning et al. 2012). Interestingly, both cycloheximide and AHLs fail to induce an expiratory pause reflex in mice treated with the nicotinic acetylcholine receptor (nAChRs) antagonists mecamylamine or

Dihydro-β-erythroidine hydrobromide (Krasteva, Canning et al. 2011, Krasteva,

Canning et al. 2012).

22

Figure 1.2: Brush Cells of the Trachea. This image features a surface view of tracheal epithelium, which is composed of diverse cell types. Chemosensory brush cells appear in green and pulmonary neuroendocrine cells in blue. Cilia are stained red, while nerve fibers appear magenta.

When stimulated by “bitter” irritants, tracheal BCs release acetylcholine (ACh) to activate peptidergic nociceptive fibers. BCs are immunoreactive for the enzyme responsible for synthesizing ACh, choline acetyltransferase (ChAT), and vesicular acetylcholine transporter (VAChT), which are prerequisites for the synaptic release of

ACh (Krasteva, Canning et al. 2011). Furthermore, the peptidergic nociceptors that form synapses with BCs express nAChRs, making them sensitive to stimulation by ACh release (Krasteva, Canning et al. 2011). Taken together with the pharmacological data,

23 this morphological evidence offers a strong case for ACh being the neurotransmitter released by BCs. Furthermore, due to the extreme similarities between BCs and SCCs, evidence of ACh neurotransmission by BCs may indicate that SCCs utilize the same mechanism to activate trigeminal nociceptors.

Nomenclature of Taste Cell-like Chemosensors

Respiratory tract solitary chemosensory cells were named after their resemblance to a chemosensory cell in the epidermis of fish. In fish, SCCs are also bottle-shaped and send a microvillar process between the scales of the epidermis to monitor the aqueous environment (Whitear 1965, Whitear 1992). In contrast to mammalian SCCs, piscine

SCCs can be innervated by facial, glossopharyngeal and spinal nerves in addition to the vagal and trigeminal nerves (Kotrschal, Whitear et al. 1993, Finger 1997). The SCCs of fish respond to heterospecific chemical cues (Peters, Kotrschal et al. 1991), a very different type of chemical signal than which activates SCCs in the airway. In addition to airway SCCs and ichtyic SCCs, the term solitary chemosensory cell has also been applied to chemosensory cells along the gastrointestinal tract of mammals (Wu, Rozengurt et al.

2002, Bezencon, le Coutre et al. 2007). Some gastrointestinal SCCs appear to utilize canonical taste transduction signaling, like respiratory SCCs. However, gastrointestinal

SCCs appear to monitor the quality of ingested food, a function more similar to the taste system than the inflammation-inducing respiratory SCCs (Wu, Rozengurt et al. 2002,

Bezencon, le Coutre et al. 2007). Furthermore, the neurosensory cells found in the gut have a more diverse function than respiratory tract SCCs.

To further confuse the nomenclature of taste cell-like respiratory chemosensors, tracheal brush cells have also been referred to as solitary chemosensory cells (Tizzano,

Cristofoletti et al. 2011, Krasteva, Hartmann et al. 2012, Merigo, Benati et al. 2012).

Despite having been referred to by the same name, nasal, gastrointestinal and piscine

24

SCCs appear to have very different functions. In contrast tracheal BCs and nasal SCCs appear to have almost identical functions but the term brush cell has a long precedence in the literature. To allow for precise discussion of these different cell types, this and subsequent chapters abide by the following conventions:

1. The term solitary chemosensory cell (SCC) refers to the taste cell-like

chemosensors found in the nasal respiratory epithelium of mammals.

2. The term brush cell (BC) refers to the taste cell-like chemosensors found

in the tracheal respiratory epithelium of mammals.

3. Anytime chemosensory cells in fish or the gastrointestinal tract are

mentioned, the term SCCs will be modified by the adjective piscine or

gastric (e.g. piscine SCCs or gastric SCCs).

Organization of Thesis

Chapter II describes experiments to determine if ACh is the neurotransmitter released from SCCs to activated peptidergic trigeminal nociceptors. Chapter III describes experiments to determine if BCs are renewed with the surrounding tracheal epithelial cells throughout an organism’s life, as SCC are renewed with the surrounding nasal epithelium.

25

CHAPTER II

CHOLINGERIC NEUROTRANSMISSION LINKS SOLITARY

CHEMOSENSORY CELLS TO NASAL INFLAMMATION.1

Abstract

Solitary chemosensory cells (SCCs) are specialized epithelial chemosensors of the airways, which respond to irritants via the canonical bitter taste transduction cascade

(T2R receptors, Gα-gustducin and TRPM5). When stimulated, SCCs trigger peptidergic nociceptive nerve fibers, causing an alteration of the respiratory rate indicative of trigeminal activation. Direct chemical excitation of trigeminal pain fibers can result in neurogenic inflammation in the surrounding epithelium. In the current study, we test whether activation of nasal SCCs can trigger local inflammation, leading to mast cell degranulation and plasma leakage into the epithelium. The prototypical bitter compound, denatonium caused significant extravasation and degranulation in wild-type mice but not in mice with a genetic deletion of elements of the canonical taste transduction cascade, demonstrating that activation of bitter receptors and their downstream signaling components can lead to local inflammation. Ablation of peptidergic trigeminal fibers by resinoferatoxin prevented the SCC-induced nasal inflammation, indicating that SCCs evoke inflammation only via neural activity and not by release of local inflammatory mediators. Additionally, blocking nicotinic acetylcholine receptors prevents SCC- mediated neurogenic inflammation, showing the necessity for cholinergic transmission from SCC to nerve fiber. Finally, we demonstrate that the neurokinin 1 receptor for

1 Contributions: This chapter is the result of collaboration between myself, Michael Christensen, Tom Finger and Marco Tizzano. Michael Christensen and Marco Tizzano conducted the experiments involving the knockout mice, resinoferatoxin-treated mice, the NK1 receptor antagonist and the mast cell experiments. My contribution to this work includes the nicotinic ACh receptor blocker experiments, immunofluorescence, draw the summarizing cartoon and writing the manuscript.

26 substance P is required for neurogenic inflammation suggesting that release of substance

P from the nerve fibers triggers both the degranulation of mast cells and the changes in endothelial cells leading to plasma leakage. Taken together, these results demonstrate that

SCCs utilize cholinergic neurotransmission to trigger peptidergic trigeminal nociceptors and link SCCs to the traditional neurogenic inflammatory pathway.

Significance Statement

Millions of people worldwide suffer from chronic nasal inflammation involving obstructed airflow and nasal discharge. While nasal inflammation is often considered to be a reaction to allergens, approximately a quarter of all cases are non-allergic rhinitis.

The causes of this disease are unknown, but symptoms may be triggered or exacerbated by a variety of inhaled irritants or even seemingly innocuous odors. We report here that specialized chemosensory cells of the nasal epithelium detect diverse chemicals and transmit this information to pain-sensing nerve terminals in the nose which then release bioactive peptides to trigger an inflammatory response―all without the necessity for activity of the adaptive immune system. This novel pathway may offer therapeutic targets for intervention in non-allergic rhinitis.

Introduction

The respiratory tract is continually assaulted by diverse combinations of irritants and xenobiotics. The nasal cavity serves as the first line of defense against these diverse noxious substances (Richards, Saunders et al. 2010) and houses two parallel chemodetection systems―trigeminal free nerve endings and solitary chemosensory cells

(SCCs)―both of which mediate protective airway reflexes (Silver and Finger 2009). The dual chemodetector systems allow for responses to irritating substances with widely varied physical and chemical properties (Silver and Finger 2009, Richards, Saunders et al. 2010). In the current study, we describe the mechanism and mediators by which the

27 parallel warning systems of free nerve endings and SCCs evoke local inflammation and recruit a pro-inflammatory response in the nasal epithelium.

Free nerve endings of the trigeminal nerve occur throughout the nasal respiratory epithelium and respond directly to many irritants via chemosensitive TRPA1 and TRPV1 ion channels (Bryant and Silver 2000, Kobayashi, Fukuoka et al. 2005). However, these intranasal trigeminal fibers terminate below the level of tight junctions at the surface of the epithelium (Finger, St Jeor et al. 1990), allowing only lipophilic compounds to reach the receptors on the free nerve endings. Accordingly, peptidergic nociceptive trigeminal fibers are responsive to only a subset of potentially dangerous compounds entering the airway.

An alternative means by which irritants can activate the trigeminal system is through solitary chemosensory cells (SCCs), which populate the respiratory epithelium of the nasal cavity (Figure 2.1C) (Finger, Bottger et al. 2003, Tizzano, Gulbransen et al.

2010, Barham, Cooper et al. 2013). These SCCs extend microvillous sensory processes into the lumen of the nasal cavity and, therefore, have access to potential irritants that cannot penetrate the epithelial barrier (Finger, Bottger et al. 2003). SCCs utilize the same chemosensory receptors and transduction cascade as do bitter-responsive taste cells, including the Tas2R (T2Rs) family of G-protein-coupled receptors, Gα-gustducin

(GNAT3), phospholipase Cβ2 (PLCβ2), and the monovalent-selective cation channel

TRPM5 (Finger, Bottger et al. 2003, Gulbransen, Clapp et al. 2008). SCCs respond to both traditional bitter compounds (e.g., denatonium) as well as bacterial metabolites (e.g., the quorum sensing factor acyl-homoserine lactones) (Tizzano, Gulbransen et al. 2010).

Our previous studies have established that the activation of SCCs evokes protective respiratory reflexes via a well-characterized trigeminal mediated brainstem reflex

(Finger, Bottger et al. 2003, Tizzano, Gulbransen et al. 2010) suggesting that SCCs form

28 functional contacts with trigeminal nerve endings. Additionally, SCCs contain vesicles that are positioned near SCC-associated nerve fibers on the ultrastructural level (Finger,

Bottger et al. 2003). Presumably, upon stimulation, SCCs release a hitherto unidentified neurotransmitter onto the nociceptive trigeminal fibers. Experiments in the present study suggest that SCCs release acetylcholine to activate nicotinic cholinergic receptors on the intraepithelial nociceptors to evoke a neurogenic inflammation.

Inflammation is classically defined by pre-Galen physicians of antiquity as the symptoms of sensitivity to pain (dolor), heat (calor), redness (rubor) and swelling (tumor)

(Rather 1971). The swelling or edema, which characterizes inflammation, results when chemical signals trigger changes in the shape of endothelial cells that compose blood vessel walls, opening the junctions between cells and resulting in the leakage of blood plasma into the extracellular space (Rodger, Tousignant et al. 1995).

Inflammation also can be characterized by the subsequent recruitment and activation of the immune system (Qureshi and Jakschik 1988, Wershil, Murakami et al.

1988). Mast cells are components of the innate immune system which reside within epithelial tissues, such as the airways, and react to pro-inflammatory mediators by releasing cytoplasmic granules. These granules contain of a broad array of biologically active substances including histamine, heparin, several proteases, lipid-derived mediators, growth factors, cytokines and chemokines (Metcalfe, Baram et al. 1997, Galli,

Nakae et al. 2005, Gri, Frossi et al. 2012). Previous studies have established that mast cells in other tissues will degranulate in response to a variety of agents, including neuropeptides, such as substance P, released from peptidergic nociceptive fibers

(Lagunoff, Martin et al. 1983, Metcalfe, Baram et al. 1997, Galli, Nakae et al. 2005). In the current study, we utilize degranulated mast cells as an indicator of activation of the immune system after irritant stimulation.

29

We postulated that SCCs can activate peptiergic trigeminal fibers, resulting in local inflammation and activation of the innate immune system. Here, we describe the mechanisms and mediators by which different classes of irritants evoke these responses.

Specifically, we confirm that peptidergic trigeminal fibers play an essential role in generating the SCC-mediated local inflammatory and early immune responses. Further, we present evidence that SCCs release the neurotransmitter acetylcholine to activate trigeminal fibers. Finally, we show that the substance P released from peptidergic trigeminal fibers is the primary signal that induces both the edema characteristic of local inflammation and mast cell degranulation indicative of an early immune response.

Materials and Methods

Animals

The mice utilized in these experiments included animals of both sexes from

ChAT-tauGFP, Gustducin-/-, TRPM5-GFP, TRPM5-/- and C57/Bl6 lines. ChAT-tauGFP mice, which were produced using a construct consisting of tauGFP-3XpolyA-Kan flanked by ChAT homologous arms, have been described previously (Grybko, Hahm et al. 2011). Similarly, Gustducin-/- (Wong, Gannon et al. 1996), TRPM5-GFP (Clapp,

Medler et al. 2006) and TRPM5-/- (Talavera, Yasumatsu et al. 2005) lines produced by R.

Margolskee (Monell Chemical senses center, Philadelphia, PA) are described in detail in

(Tizzano, Gulbransen et al. 2010). All mice were maintained on a 12-hour light/dark cycle and were given ad libitum access to food and water. All experimental procedures were approved by the University of Colorado Anschutz Medical Campus Institutional

Animal Care and Use Committee.

30

Immunofluorescence

For all Immunofluorescence experiments, mice were anesthetized with i.p. 150 mg/kg sodium pentobarbital before transcardial perfusion with 4% paraformaldehyde

(PFA) in 0.1 M phosphate buffer (PB) pH 7.2 - 7.4. To visualize blood vessels in whole mounts of nasal epithelium, mice were perfused with 5mg of lysine fixable dextran conjugated to micro-Ruby (3000 MW, Invitrogen) in 5mL of PBS before fixation. Heads were post-fixed for two to four hours and stored overnight in cryoprotectant wash, 20% sucrose in PB. The nasal epithelium was dissected free and embedded in OCT (Fisher

Scientific, Waltham, MA) medium for sectioning on a cryostat. Sixteen µm cross- sections were cut onto Fisher Plus slides (Fisher Scientific) and stored overnight or longer at -20°C. After being brought to room temperature, slides for immunohistochemistry were rinsed several times in 0.1 M phosphate buffered 0.9% (w/v) saline (PBS), dried on a hot plate at 39°C to improve tissue adherence, and rehydrated in

PBS before being blocked for one hr in 1% Normal Donkey Serum 1% Triton in PBS.

Tissue was incubated overnight with primary antisera diluted in this same blocking solution.

For dual GFP/Gustducin immunofluorescence, tissue was incubated overnight with 1:600 chicken anti-GFP (Cat# AB16901, Millipore) and 1:1000 rabbit anti-α- gustducin (raised against amino acids 93–112 of rat G α-gustducin, Cat# SC-395, Santa

Cruz Biotechnology, Santa Cruz, CA) primary antibodies. For dual CGRP/substance P immunofluorescence, tissue was incubated with 1:500 Rabbit anti-CGRP (Peninsula

Labs) and 1:500 Rat anti-substance P (Accurate Chemical). After several PBS rinses, tissue was incubated with appropriate secondary antisera: 1:500 Alexa Fluor 568 donkey anti-rabbit (Invitrogen), 1:500 Alexa 568 goat anti-rat (1:400; Molecular Probes), 1:500

31

Alexa Fluor 488 donkey anti-chicken (Jackson Immuno Research) secondary antisera diluted in blocking solution.

Resinoferatoxin ablation of TRPV1-expressing nerve terminals

To eliminate nerve endings containing the TRPV1 receptor, we utilized a previously established protocol (Cavanaugh, Chesler et al. 2011) entailing multiple injections of the potent capsaicin analog, resiniferatoxin (RTx). Mice were anesthetized with 5% isoflurane using a Matrx VIP 3000 calibrated vaporizer (Midmark, Versailles,

OH) until unresponsive, then maintained on a dose of 2% isoflurane. The level of isoflurane was then reduced to 1.7% after 10 minutes, 1.5% after 30 minutes. After one hour the mice were taken off the anesthetic. The isoflurane dosage was adjusted to maintain a respiration rate between 60 and 90 breaths per minute. During anesthesia, mice were given 1 mL of saline s.c. to prevent dehydration, treated with protective eye ointment and kept on a heating pad.

Once anesthetized, mice were injected s.c. with either RTx or the vehicle. The

RTx was dissolved in 95% ethanol to the concentration of 1mg/mL, and further diluted in saline. Vehicle-injected mice received ethanol diluted in saline in the same volume as the

RTx-injected mice received. Over a four-day period, mice were injected with 15 µg/kg,

40 µg/kg, 100 µg/kg and 200 µg/kg RTx with a single final injection of 200 µg/kg RTx

11 days after the initial injection.

Plasma extravasation

Based on previous studies (Greiff, Erjefalt et al. 1991, Rodger, Tousignant et al.

1995, Carreno, Domenech et al. 2012), we have developed an assay that utilizes fluorescently-conjugated albumin to measure plasma leakage as an indication of nasal inflammation following application of potential irritants to one nostril. Mice were anesthetized with 1.0 g/kg urethane (Sigma) and placed under a warming lamp for the

32 duration of the experiment. After mice had been fully anesthetized for 5 mins, 20 μL of either 10 mM Denatonium, 2 μM Capsaicin or saline were applied dropwise to the right naris. Five mins after nasal stimulation, mice were injected in the tail vein with 25 mg/kg of fluorescent albumin-A555 in saline. After 5 additional minutes to allow plasma extravasation, mice were perfused with saline and fixative as described above, and the heads bisected in the midsagittal plane. After removal of the nasal septum, photos were taken of the left and right nasal turbinates and fluorescence intensity was quantified using

ImageJ (http://rsb.info.nih.gov/ij/). To generate relative fluorescence values the following formula was used:

RF = (STIM-UNSTIM)/UNSTIM,

where RF=relative fluorescence, STIM = intensity of stimulated nasal cavity;

UNSTIM = intensity of unstimulated nasal cavity. Each experiment was analyzed via a one-way ANOVA and subsequent Tukey’s HSD test.

Mast cell degranulation

To determine the ratio of degranulated to non-degranulated mast cells, we used acidified toluidine blue to stain mast cells in nasal epithelia from mice that had been stimulated retronasally by potential irritants. Mice were anesthetized with 1g/Kg of urethane, and a two-way tracheotomy was performed as described previously (Tizzano,

Gulbransen et al. 2010). In brief, a cannula was inserted into the trachea, and another cannula was inserted retrotracheally into the rhinopharynx to permit chemical stimulation of the respiratory epithelium. Denatonium benzoate (10mM), capsaicin (2µM) or saline were passed through the retronasal cannula to stimulate the nasal cavity. After 10 min, mice were perfused and postfixed as described above. The respiratory epithelium was dissected, stained with acidified toluidine blue and mounted on slides (Blumenkrantz and

Asboe-Hansen 1975). Approximately 500 photos were taken to document the state of

33 over 29,000 mast cells in the nasal tissues. The micrographs were evaluated by an investigator blinded to the treatment conditions for the animal, who counted the number of non-degranulated and degranulated mast cells. A mast cell was considered to be degranulated when more than 5 granules were observed in the immediate vicinity, but outside of the mast cell.

To control for day-to-day variability in baseline mast cell degranulation, some mice were stimulated with saline on each day of the experiment. The baseline percent of degranulated mast cells in these saline-stimulated mice was subtracted from the percent of degranulated mast cells in the other experiment groups. As positive control for mast cell degranulation, some mice were injected i.p. with the classic mast cell secretagogue, compound 48/80 (Sigma-Aldrich) (Rothschild 1970), dissolved in saline. Compound

48/80 acts directly on mast cell to cause degranulation and was used in RTx treated mice as a positive control.

Pharmacology

To test the role of nicotinic cholinergic receptors (nAChR), we utilized the nAChR antagonist mecamylamine hydrochloride (Tocris, Bristol, UK). Mice were injected i.p. with either 3 mg/kg or 6 mg/kg Mecamylamine, or vehicle control, 10 mins prior to urethane injection. To test the role of NK1 receptors, we injected mice i.p. with 5 mg/kg of the NK1 blocker, L73213 (Tocris, Bristol, UK).

Results

Solitary chemosensory cells (SCCs) are cholinergic and contact peptidergic nociceptors

Solitary chemosensory cells in the nasal respiratory epithelium are taste cell-like chemosensors, which express TRPM5 and G-gustducin (Figure 2.1A) (Finger, Bottger et al. 2003, Tizzano, Gulbransen et al. 2010, Barham, Cooper et al. 2013). In the trachea, a related cell type, the chemosensory brush cell, releases ACh to activate vagal pain

34

Figure 2.1: Cross sections of nasal epithelium showing cellular properties and relationships of SCCs. (A) SCCs are immunoreactive for TRPM5 (green) and Gustducin (red), elements of the canonical taste transduction pathway. (B) In addition to gustducin (red), SCCs are also immunoreactive for ChAT-GFP (green), the acetylcholine (ACh) synthesizing enzyme. (C) SCCs expressing TRPM5 (green) are intimately contacted by CGRP immunoreactive peptidergic nociceptive trigeminal fibers. (D) These peptidergic nociceptive fibers (magenta) contact SCCs (TRPM5-GFP immunoreactivity in green) and are immunoreactive for both CGRP (red) and substance P (blue). The nuclear counterstain DRAQ5 is shown in cyan in panels A-C. Scale bars = 10 µm

fibers (Krasteva, Canning et al. 2011, Krasteva, Canning et al. 2012, Saunders, Reynolds et al. 2013) suggesting ACh as a candidate neurotransmitter for SCCs. To determine if

SCCs express choline acetyltransferase (ChAT), the synthetic enzyme of acetylcholine, we examined the nasal epithelium of a transgenic mouse, which expresses tauGFP driven by the ChAT promoter (Grybko, Hahm et al. 2011). In the nose of these mice, cells immunoreactive for gustducin also express GFP (Figure 2.1B) driven by the ChAT promoter. This result suggests that SCCs, like brush cells, are capable of producing ACh

35 for release onto nerve fibers (Krasteva, Canning et al. 2011, Saunders, Reynolds et al.

2013).

The trigeminal nerve fibers that contact the SCCs are immunoreactive for substance P and calcitonin gene related peptide (CGRP) (Figure 2.1CD) both of which are inflammatory mediators typically co-expressed and released by nociceptive (e.g.,

“pain”) fibers (Finger, Bottger et al. 2003). The peptidergic trigeminal nociceptive fibers branch within the epithelium contact SCCs and terminate as free nerve endings in nearby epithelium (Figure 2.1C). The anatomical arrangement of SCCs and trigeminal free nerve endings potentially offers dual pathways for trigeminal activation by irritants, since not only the SCCs, but the free nerve endings themselves, are chemosensitive.

Gustducin and TRPM5 are required for SCC mediated inflammation

Direct chemical activation of trigeminal nasal nociceptors evokes neurogenic inflammation (Lundberg, Brodin et al. 1984). To test the role of SCCs in irritant-induced nasal inflammation, we stimulated mice with either capsaicin or denatonium. Capsaicin directly activates peptidergic trigeminal nociceptors via TRPV1 channels (Lundblad and

Lundberg 1984, Caterina, Schumacher et al. 1997), while denatonium activates “bitter” taste receptors on SCCs in vitro (Gulbransen, Clapp et al. 2008) and is capable of triggering SCC-mediated respiratory reflexes typical of trigeminal simulation in vivo

(Tizzano, Gulbransen et al. 2010). Application of either capsaicin or denatonium to the nasal cavity resulted in a significant increase to both plasma extravasation (Figure

2.2ABD) and the percentage of degranulated mast cells compared to baseline (Figure

2.3ABC). To test whether the inflammation provoked by denatonium, but not by capsaicin requires functional SCCs, we tested both compounds on mice with a disruption of the bitter taste signaling cascade by genetic deletion of either gustducin or TRPM5.

Both gustducin-/- and TRPM5-/- mice showed significantly less plasma extravasation

36 than wild type controls when stimulated with denatonium (Figure 2.2CD), but not with capsaicin (Figure 2.2D). These data are consistent with the hypothesis that functional

SCCs are required for responses to denatonium, but not capsaicin.

Similarly, in TRPM5-/- mice SCC-mediated mast cell degranulation was deficient in response to denatonium, but not to capsaicin, an activator of TRPV1 (Figure 2.3C). To test whether genetic deletion of either gustducin or TRPM5 reduced the capacity for mast cell degranulation, we tested animals of each genotype with the canonical mast cell secretagogue, compound 48/80 (C48/80). Wild type, gustducin-/- and TRPM5-/- mice all showed a similar levels of mast cell degranulation when injected with compound 48/80

(Figure 2.3C). Taken together, these results indicate that SCCs and trigeminal free nerve endings act as twin pro-inflammatory pathways, detecting different chemicals, but ultimately triggering the same local inflammatory response and early immune reactions.

Peptidergic nociceptive trigeminal fibers are required for SCC mediated inflammation

Both SCCs and trigeminal free nerve endings are capable of triggering similar inflammatory and immune reactions (Figure 2.2D and Figure 2.3C). Conceivably, SCCs might trigger this response independent of the nerve via paracrine signals. Alternatively,

SCC-mediated inflammatory and immune reactions might be entirely dependent on the release of neuropeptides from nociceptive trigeminal fibers. To test if peptidergic trigeminal fibers are required for SCC-mediated inflammation and early immune response, we ablated peptidergic nociceptive fibers by treating mice repeatedly with resiniferatoxin (RTx) (Szolcsanyi, Szallasi et al. 1990, Acs, Biro et al. 1997), which destroys TRPV1-expressing peptidergic trigeminal fibers (Szallasi, Joo et al. 1989).

37

Figure 2.2: Stimulation of SCCs activates a pro-inflammatory pathway that triggers plasma extravasation.

38

Figure 2.2: Stimulation of SCCs activates a pro-inflammatory pathway that triggers plasma extravasation. (A) Brightfield image of a bisected head of a mouse showing the two sides of the nasal cavity imaged for the plasma extravasation experiments (scale bar = 1mm). Anterior is up. (B) Flourescence image from a wild type mouse stimulated unilaterally with 10 µM denatonium benzoate, showing increased Alexa555-albumin fluorescence on the stimulated side. (C) Image from a Gustducin-/- mouse stimulated unilaterally with 10 µM denatonium, showing no significant fluorescence on either the stimulated or unstimulated sides. (D - G) Bar graphs illustrating the relative fluorescence of stimulated and unstimulated sides under various conditions and genotypes. Positive values indicate that the stimulated side was brighter than the unstimulated side. (D) Wild type mice stimulated with 10 µM denatonium (green) or 2 μM capsaicin (red) showed significantly more (P<0.001 and P<0.01, respectively) plasma extravasation compared to saline-stimulated mice (blue). Two knockout strains, Gustducin-/- and TRPM5-/-, showed significantly less (P<0.001) extravasation than wild type controls stimulated with denatonium, but normal extravasation with capsaicin. (E) Mice treated with Resinferatoxin (RTx) to eliminate nerve fibers showed significantly less plasma extravasation in response to both denatonium (P<0.001) and capsaicin (P<0.01), than did vehicle treated controls. (F) The nAChR antagonist Mecamylamine significantly reduced denatonium-induced extravasation at both 3mg/kg (P<0.01) and 6 mg/kg (P<0.001) but did not alter capsaicin-induced extravasation. (G) The NK1 antagonist L732138 (i.p. 5 mg/kg), which blocks responses to substance P significantly reduces plasma extravasation in response to both denatonium (P<0.001) and capsaicin (P<0.001) *P<0.05, **P <0.01, ***P<0.001 by one-way ANOVA.

39

In RTx treated mice, capsaicin failed to cause either plasma extravasation or mast cell degranulation, indicating that the peptidergic trigeminal fibers were ablated and necessary for the responses to capsaicin (Figure 2.2E and Figure 2.3D). If SCCs were capable of triggering an inflammatory response through a paracrine pathway, then denatonium would be able to induce plasma extravasation and mast cell degranulation despite the destruction of peptidergic nerve fiber terminals in RTx-treated mice.

However, denatonium fails to cause any significant plasma extravasation in RTx treated mice (Figure 2.2E) or mast cell degranulation (Figure 2.3D), affirming the necessity for polymodal nociceptive nerve fibers in these responses. Taken together, these results indicate that SCC-mediated local inflammation and early immune response requires the presence of peptidergic trigeminal fibers to convey signals from the SCCs to the post- capillary venules and mast cells (McDonald, Thurston et al. 1999).

SCC-mediated inflammation requires nicotinic ACh receptors (nAChRs)

Our results described above demonstrate that SCCs require the presence of nociceptive trigeminal nerve fibers to trigger inflammation (Figure 2.2E) and activation of the innate immune system (Figure 2.3D). The presence of synapses between SCCs and trigeminal fibers (Finger, Bottger et al. 2003) suggests that the SCCs release a neurotransmitter which activates the sensory nerve endings. The expression of choline acetyltransferase suggests that SCCs may be cholinergic (Figure 2.1B), and previous studies have demonstrated the presence of nAChRs on trigeminal fibers (Alimohammadi and Silver 2000). To test whether nAChRs are necessary for SCC-mediated effects, we treated mice with mecamylamine, a blocker of nAChRs, and tested responses to denatonium and capsaicin. Mecamylamine significantly reduced responses to denatonium but had no significant effect on responsiveness to capsaicin, reflecting the necessity for cholinergic neurotransmission for SCC-mediated effects, but not for direct neural effects.

40

Figure 2.3: Stimulation of SCCs activates a pro-inflammatory pathway that triggers mast cell degranulation.

41

Figure 2.3: Stimulation of SCCs activates a pro-inflammatory pathway that triggers mast cell degranulation. Photos of resting (A) and degranulated (B) mast cells stained with acidified toluidine blue. Arrows point to granules forming a halo around the degranulated mast cells. (C) Wild type mice stimulated with 10 µM denatonium showed significantly more mast cell degranulation than TRPM5-/- (P<0.05). Wild type and TRPM5-/- mice responded similarly to 2 μM capsaicin. (D) Mice treated with Resinferatoxin (RTx) to eliminate nerve fibers showed significantly less mast cell degranulation in response to both denatonium (P<0.001) and capsaicin (P<0.01), than did vehicle-treated controls. C48/80, which directly acts on mast cells, caused degranulation in RTx-treated mice. (E) The nAChR antagonist mecamylamine significantly reduced denatonium induced mast cell degranulation at a dose of 6mg/kg

(P<0.01). (F) The NK1 antagonist L732138 (i.p. 5 mg/kg), which blocks responses to substance P significantly reduces mast cell degranulation in response to both denatonium (P<0.05) and capsaicin (P<0.05). *P<0.05, **P <0.01, ***P<0.001 by one-way ANOVA.

42

Mecamylamine treatment significantly reduced plasma extravasation in a dose dependent manner (Figure 2.2F) for denatonium stimulation. Similarly, mecamylamine significantly reduced the proportion of dengranulated mast cells following denatonium stimulation

(Figure 2.3E). However, mecamylamine had no significant effect on the ability of capsaicin to induce plasma extravasation (Figure 2.2F). Taken together, these results indicate that nAChR is required for SCCs to induce local inflammation and early immune responses and most likely reflect nicotinic cholinergic transmission from SCCs to nerve fibers.

NK1 receptors underlie both plasma extravasation and mast cell degranulation

In other systems, neural-induced plasma extravasation depends on the release of substance P or neurokinin A from the nerve fibers, acting respectively on neurokinin 1 receptors (NK1) or neurokinin 2 receptors (NK2) on the endothelial cells (Rodger,

Tousignant et al. 1995). Since the trigeminal polymodal nociceptors of the nasal cavity express substance P, we utilized the NK1 antagonist L732138 to test whether NK1 receptors are required for irritant-induced plasma extravasation. If so, than blocking NK1 receptors should prevent plasma leakage and mast cell degranulation regardless of the avenue of detection (SCC or TRP channels on free nerve endings).

For both denatonium (p<0.001) and capsaicin (p<0.001), an injection of L732138

(i.p. 5 mg/kg) significantly reduced plasma extravasation compared to the vehicle- injected controls (Figure 2.2G). The level of plasma extravasation in mice treated with the NK1 antagonist was not significantly different than control mice stimulated with saline (Figure 2.2G). Additionally, L732138 treatment prevented mast cell degranulation in mice stimulated with either denatonium or capsaicin (Figure 2.3F). Taken together, these results indicate that NK1 activation is necessary for both irritant-induced plasma

43 extravasation and mast cell degranulation regardless of whether the irritant acts via SCCs or directly on the nerve.

Discussion

Our results delineate the mechanism by which chemical activation of nasal epithelial chemosensors, SCCs, can evoke neurogenic inflammation in the absence of an immunoglobulin-mediated response. SCCs utilize the canonical bitter taste transduction cascade to generate responses to diverse noxious and innocuous air-borne or bacterially- derived substances. The SCCs then release the neurotransmitter ACh to activate nAChRs on peptidergic trigeminal nerve fibers within the epithelium (Figure 2.4). Intramucosal collaterals of these nerve fibers in turn release substance P to activate NK1 receptors on the endothelial cells and mast cells resulting in plasma leakage from the vessels and mast cell degranulation, respectively (Figure 2.4). Since plasma extravasation and mast cell degranulation are hallmarks of early inflammatory states, this pathway provides an avenue by which non-allergenic irritants can evoke nasal inflammation.

Parallel pathways for airway irritation

In the airway, multiple pathways exist to trigger inflammatory responses to potentially dangerous substances: immunoglobulin E (IgE)-mediated (Shusterman and

Murphy 2007), nociceptive nerve fibers and SCC-mediated. Each of these pathways can respond to different classes of irritating substances, but all 3 are interconnected through downstream mechanisms. Additionally, while some elements of these pathways are convergent, portions of each are capable of acting independently. The existence of parallel pathways allows for a broadly tuned and redundant warning system for protection of the airway.

IgEs mediate type I immune hypersensitivity, which is responsible for inflammation in allergic rhinitis. In this case, xenobiotics in the airway are bound by IgE,

44 which in turn triggers mast cell activation and degranulation which initiates inflammation

(Metcalfe, Baram et al. 1997, Gilfillan and Tkaczyk 2006). IgEs that are bound to the IgE receptors on mast cells are able to cause degranulation directly (Gilfillan and Tkaczyk

2006), but free IgEs can also trigger mast cell activation through an alternative pathway

(Ishizaka, Sian et al. 1972). Typically, the xenobiotics that activate the IgE pathway are large macromolecules such as the proteins in pollen (MacGlashan 2012). This IgE- mediated pathway is the most independent of the 3 mechanisms and does not require any involvement from other cell types to triggering mast cell activation (Gilfillan and

Tkaczyk 2006).

The second pro-inflammatory pathway involves stimulation of peptidergic nociceptive nerve fibers, which respond directly to diverse irritants due to the presence of several chemosensitive ion channels. Nociceptive fibers cause neurogenic inflammation through an axon reflex mechanism—the release of neuropeptides from both the stimulated terminals and collateral branches of the same axon (Bryant and Silver 2000).

CGRP and substance P released by the axon reflex have multiple effects including the ability to sensitize nociceptors, (Bryant and Silver 2000, Richards, Saunders et al. 2010), cause inflammatory edema (Figure 2.2G) and trigger mast cell degranulation (Figure

2.3F). Substance P, released from peptidergic nociceptors, is capable of causing inflammation in the absence of the IgE mediated pathway via NK1 receptors on mast cells (Figure 2.3F). Thus, mast cell degranulation appears to be an important integrator of pro-inflammatory signals (Gilfillan and Tkaczyk 2006), and substance P release links neurogenic inflammation to the immune system through these cells (Metcalfe, Baram et al. 1997). However, trigeminal nociceptors are limited in the types of noxious compounds to which they can respond. Specifically, lipophobic compounds in the lumen of the nasal cavity are incapable of crossing the epithelial barrier to reach the underlying nerve fibers.

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Figure 2.4: Parallel pathways for airway irritation. Solitary chemosensory cells (SCCs = green) respond to many bitter substances such as denatonium and bacterial metabolites via the canonical taste transduction pathway. SCCs release acetylcholine (ACh), which activates nicotinic acetylcholine receptors on nociceptive trigeminal nerve fibers (white) that innervate the SCCs. These nociceptive trigeminal fibers also express TrpV1 and TrpA1, two chemosensitive ion channels that are responsive to irritants such as capsaicin. Regardless of whether the nociceptive fiber is activated directly via TRP channels, or indirectly via SCCs, the fibers release inflammatory mediators. One of these mediators, substance P (SubP), activates NK1 receptors on blood vessels (red) resulting in plasma extravastion and on mast cells (purple) causing degranulation. Insert: “Bitter” irritants stimulate bitter receptors (T2R) to activate the βγ subunit associated with the G-protein Gustducin which produces inositiol trisphosphate (IP3) via a PLCβ2 mediated cascade. In 2+ turn, IP3 binds to the type 3 IP3 receptor (IP3R3), releasing Ca from the endoplasmic reticulum. Increases in cytosolic Ca2+ activates TRPM5, a non-specific cation channel, and leads to the release of ACh.

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In contrast to nerve fibers that lie beneath the epithelial surface, SCCs extend their apex into the lumen of the nasal cavity. Metabolites and quorum factors produced by potentially virulent bacteria activate SCCs via the canonical “bitter” taste transduction pathway (Tizzano, Gulbransen et al. 2010). When stimulated by these compounds, SCCs release ACh to activate peptidergic nociceptors (Figure 2.2F). Having a specialized epithelial cell mechanism by which compounds in the lumen of the airway can be detected allows for the trigeminal system to detect potentially injurious bacteria before that bacteria has achieved a sufficient concentration to convert to a more pathogenic phenotype and form biofilms (Tizzano, Gulbransen et al. 2010). Additionally, this early warning system may allow for the detection of virulent bacteria before the pathogen has caused tissue damage. Activation of the trigeminal system by ACh results in substance P release (Figure 2.2FG and Figure 2.3EF) which can lead to inflammation and the recruitment of the immune system (Metcalfe, Baram et al. 1997). SCCs are entirely dependent on peptidergic nerve fibers to transduce their pro-inflammatory signal (Figure

2.2E and Figure 2.3D). However, SCCs may release ACh in a paracellular manner to trigger local defenses in the airway, such as increasing ciliary frequency and mucus secretion (Lee, Xiong et al. 2012), which could occur independent of trigeminal nerve fibers.

Although the IgE, neuronal and SCC mediated pro-inflammatory pathways are parallel, all three converge on blood vessels and mast cells to cause inflammation. These convergences allow for interactions between the three pathways that could lead to sensitization. The neuronal-mediated pathways and the IgE pathway appear to be capable of causing inflammatory edema independently. While mast cell degranulation, like plasma extravasation, can also be triggered through multiple mechanisms, mast cell

47 degranulation can also cause many of the symptoms of inflammation. Thus, the mast cell could be viewed as an integrator of signals from all three pathways.

Mast cells—a node in the inflammatory signaling cascade

Mast cells are capable of causing the symptoms of inflammation via the release of signaling molecules during degranulation (Metcalfe, Baram et al. 1997). All three of the parallel inflammatory pathways result in mast cell degranulation, allowing for modulation or sensitization of inflammatory symptoms by mast cells (Metcalfe, Baram et al. 1997). Additionally, many of the mediators released from mast cells have a synergistic effect on inflammation, creating positive feedback, which could easily lead to pathological inflammation (Metcalfe, Baram et al. 1997, MacGlashan 2012, Voedisch,

Rochlitzer et al. 2012).

Mast cells release numerous cytokines during degranulation including TNF-α and interleukin-4 (Breuel and De Ponti 2006, Urb and Sheppard 2012) (Nakae, Suto et al.

2005, Suto, Nakae et al. 2006). One of the primary functions of cytokines is to recruit the cells of the immune system to the site of inflammation (Galli, Nakae et al. 2005, Gilfillan and Tkaczyk 2006, Gri, Frossi et al. 2012). Traditionally, mast cells have been implicated in the recruitment of macrophages to the site of parasite infection (Dawicki and Marshall

2007). However, mast cells have also been implicated in the recruitment of leukocytes involved in adaptive immunity—specifically dendritic cells and T-cells (Galli,

Kalesnikoff et al. 2005, Galli, Nakae et al. 2005, Dawicki and Marshall 2007). By triggering mast cell degranulation in response to bacterial metabolites in the lumen of the airway, SCC stimulation could recruit the innate and adaptive immune system before virulent bacteria have damaged the airway. However, cytokine signaling is complex and could easily over-sensitize the airway to benign stimuli.

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SCCs, one component of the airway chemofensor complex

Since the initial reports about SCCs, many other researchers have suggested that airway chemosensory receptors have a protective role (Deshpande, Wang et al. 2010,

Krasteva, Canning et al. 2011, Krasteva, Canning et al. 2012, Lee, Xiong et al. 2012).

SCCs and other novel chemical senses can be considered part of a chemofensor complex—i.e., that alert an organism to harmful xenobiotics and toxic chemicals (Green 2012). SCCs in the nose and chemosensory brush cells in the trachea are to be responsible for triggering inflammation and recruiting an immune response in reaction to the presence of bacterial metabolites (Tizzano, Gulbransen et al. 2010,

Krasteva, Canning et al. 2012). Additionally, stimulation of chemosensory cilia in the nose and the trachea appears to promote protective mechanisms such as the production of reactive oxygen species and antimicrobial peptides (Lee, Xiong et al. 2012). Taste receptors in tracheal smooth muscle may also have a protective effect (Deshpande, Wang et al. 2010). However, of all the newly discovered components of the airway chemofensor complex, only SCCs in the nose and chemosensory brush cells of the trachea are known to create the neurogenic inflammation that assists in recruiting both the innate and adaptive immune response. The neurogenic inflammation caused by SCC stimulation results in an irritated state that is very similar to the symptoms of non-allergic rhinitis.

SCC over-stimulation—a possible pathology for non-allergic (idiopathic) rhinitis?

In the present study, we demonstrate that stimulation of SCCs triggers a pro- inflammatory pathway resulting in the most common symptoms of idiopathic non- allergic rhinitis (NAR) (Rodger, Tousignant et al. 1995, Shusterman and Murphy 2007,

Gelardi, Maselli del Giudice et al. 2008, Knipping, Holzhausen et al. 2009). NAR, a condition with no known cause and limited treatment options, negatively impacts the

49 quality of life of millions (Venge 1994, Bousquet, Fokkens et al. 2008, Settipane and

Kaliner 2013). Conversely, IgE-mediated allergic rhinitis is well understood and there are many treatment options available for those patients (Gliklich and Metson 1995,

Benninger, Ferguson et al. 2003, Jones 2005, Bousquet, Khaltaev et al. 2008). In addition to patients with NAR, individuals afflicted with asthma or COPD also report hyper- responsiveness to normally benign odorants (Hargreave, Dolovich et al. 1986), including many chemicals that activate SCCs (Lin, Ogura et al. 2008). Essentially, our data elucidate a pathway by which commonly encountered compounds and xenobiotics could lead to a chronic inflammatory state independent of IgE-mediated mechanism—a prerequisite for any proposed cause of non-allergic rhinitis. If SCCs are involved in creating the hyper-inflammatory state of non-allergic rhinitis or exacerbate this condition, then anti-cholinergics or NK1 antagonists may provide relief for individuals suffering from this condition.

Summary

The current study demonstrates that activation of the SCCs in the nasal epithelium of mice can lead to a rapid, neurogenic local pro-inflammatory response. This fast pro- inflammatory response is potentially a defense mechanism to alert the downstream immune and inflammatory systems of a potential danger. SCCs release ACh to activate nAChRs on peptidergic trigeminal nociceptors. When stimulated by ACh, nociceptors release substance P to cause inflammatory edema and mast cell degranulation, consistent with the inflammation observed in individuals with chronic non-allergic rhinitis.

Cholinergic neurotransmission between SCCs and trigeminal fibers represents a potential therapeutic target which could be utilized to block SCC-mediated inflammation for particular classes of irritants.

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Acknowledgements

We thank Sukumar Vijayaraghavan (University of Colorado School of Medicine) for providing the ChAT-tauGFP mice and Robert Margolskee (Monell Chemical Senses

Center) for providing the TRPM5-GFP, TRPM5 knockout and Gustducin knockout mice.

Additionally, we are grateful for Jennifer Strafford’s (University of Colorado School of

Medicine) statistical advice and to Vijay Ramakrishnan (University of Colorado School of Medicine) and J. John Cohen (University of Colorado School of Medicine) for commenting on earlier drafts of this manuscript. This work was supported by National

Institutes of Health grants NIDCD R03 DC012413 (M.T.), R01 DC009820 (T.E.F.), and

P30 DC04657 (to D. Restrepo).

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CHAPTER III

CHEMOSENSORY BRUSH CELLS OF THE TRACHEA: A STABLE

POPULATION IN A DYNAMIC EPITHELIUM2

Abstract

Tracheal brush cells (BCs) are specialized epithelial chemosensors which utilize the canonical taste transduction cascade to detect irritants. To test whether BCs are replaced at the same rate as other cells in the surrounding epithelium of adult mice, we used 5-bromo-2'-deoxyuridine (BrdU) to label dividing cells. Although scattered BrdU- labeled epithelial cells are present 5-20 days post-BrdU, no BCs are labeled. These data indicate that BCs are a relatively static population. To determine how BCs are generated during development, we injected 5-day-old mice with BrdU and found labeled BCs and non-BC epithelial cells 5 days post-BrdU. Over the following 60 days, the percentage of labeled BCs increased, while the percentage of other labeled cell types decreased. These data suggest that BCs are generated from non-BC progenitor cells during postnatal tracheal growth. To test whether the adult epithelium retains the capacity to generate

BCs, tracheal epithelial cells were recovered from adult mice and grown in an air-liquid interface (ALI) culture. After transition to differentiation conditions, BCs are detected and are 1% of the total cell population by day 14. BrdU added to cultures prior to differentiation of BCs was chased into BCs, indicating that the increase in BC density is due to proliferation of a non-BC progenitor. We conclude that: 1) BCs are normally a static population in adult mice; 2) BC progenitors proliferate and differentiate during

2 The work presented in this chapter has been published in the American Journal of Respiratory Cell and Molecular Biology (Saunders, C. J., S. D. Reynolds and T. E. Finger (2013). "Chemosensory Brush Cells of the Trachea: A Stable Population in a Dynamic Epithelium." Am J Respir Cell Mol Biol.)

52 neonatal development; and 3) BCs can be regenerated from a proliferative population resident in the adult epithelium.

Introduction

The tracheal epithelium is a complex tissue containing diverse cell types that include ciliated cells, Clara-like cells, and basal cells. These cells are slowly replaced in the adult but can be regenerated rapidly in the case of cell death or overt epithelial injury

(Hong, Reynolds et al. 2004, Rawlins and Hogan 2008, Rawlins, Okubo et al. 2009,

Cole, Smith et al. 2010, Reynolds and Malkinson 2010, Rock, Randell et al. 2010).

Relatively less studied are brush cells (BCs), which are specialized epithelial chemosensors. BCs are relatively rare compared to other epithelial cell types and are distributed throughout the tracheal epithelium (Krasteva, Canning et al. 2011). Whether these specialized chemosensors are replaced at the same rate as other cells in the adult trachea is unknown (Reid, Meyrick et al. 2005).

Although BCs were long speculated to be sensory end organs (Luciano, Reale et al. 1969), only recently has this chemosensory function been confirmed (Krasteva,

Canning et al. 2011). In responding to many noxious substances, BCs utilize the canonical taste transduction pathway: type 2 Taste receptors (T2Rs), Gα-gustducin,

PLCβ2, IP3R3 and Transient Receptor Potential Melastatin 5 (TRPM5) (Finger, Bottger et al. 2003, Merigo, Benati et al. 2005, Tizzano, Cristofoletti et al. 2011, Krasteva,

Canning et al. 2012). TRPM5 is a non-specific cation channel, which has been used as a marker for several taste cell-like airway chemosensors (Finger, Bottger et al. 2003,

Kaske, Krasteva et al. 2007, Hansen and Finger 2008, Tizzano, Cristofoletti et al. 2011).

BCs respond to “bitter-tasting” irritants by releasing acetylcholine, which activates nearby vagal nerve fibers to elicit protective respiratory reflexes (Krasteva, Canning et al.

2011). The tracheal BCs are similar to nasal solitary chemosensory cells (SCCs), which

53 also utilize the canonical taste transduction cascade (Finger, Bottger et al. 2003, Silver,

Roe et al. 2010).

In other chemosensory systems (e.g., main and accessory olfactory epithelia, taste buds and nasal solitary chemosensory cells), the sensory cells are replaced at a rate similar to that of surrounding epithelium (Beidler and Smallman 1965, Moulton 1974,

Gulbransen and Finger 2005)―e.g. over a span of a few weeks. The lifespan of ciliated and Clara-like cells in tracheal epithelium is longer than in nasal and lingual epithelia

(Basbaum and Jany 1990, Rawlins and Hogan 2008, Rawlins, Okubo et al. 2008,

Reynolds and Malkinson 2010), but the lifespan of BCs has yet to be determined. We predicted that BC renewal and generation would occur at the same rate as the surrounding epithelium and tested this with 5-bromo-2'-deoxyuridine (BrdU) labeling.

Surprisingly, BCs show no evidence of turnover in the adult epithelium. This finding led us to examine the initial generation of BCs during development and the capacity to generate BCs in an in vitro injury model in adult epithelium.

Materials and Methods

The mice utilized in these experiments included C57Bl6, TRPM5-GFP and

ChAT-tauGFP lines for in vivo studies and A/J and TRPM5-GFP mice were used to generate trachea epithelial cultures. TRPM5-GFP mice, which were produced using a construct consisting of 11 kb of untranslated TrpM5 5' flanking sequence through Exon 2 and eGFP, were a gift of R.F. Margolskee of the Monell Chemical Senses Center

(Philadelphia, PA) (Clapp, Medler et al. 2006). ChAT-tauGFP mice, which were produced using a construct consisting of tauGFP-3XpolyA-Kan flanked by ChAT homologous arms, have been previously described (Grybko, Hahm et al. 2011). All mice were maintained on a 12-hour light/dark cycle and were given ad libitum access to food and water. All experimental procedures were approved by the University of Colorado

54

Anshutz Medical Campus and National Jewish Health Institutional Animal Care and Use

Committees.

In vivo experiments

For studies of turnover and proliferation, 5-bromo-2'-deoxyuridine (BrdU, Sigma-

Aldrich, St Louis, MO) dissolved in 0.1 M phosphate buffered 0.9% (w/v) saline (PBS) was injected i.p. to label cells in S-phase. For experiments on adults, mice received a total of three injections of 150 mg/kg BrdU (450 mg/kg BrdU total). The first injection was administered at the onset of the 12-hour light phase and subsequent injections at 6- hour intervals thereafter. Adult mice were killed at 5-day intervals after BrdU injections on days 5 to 20 post BrdU (Figure 3.1A). For experiments on pups, 5-day old mice received a single i.p. injection of 100 mg/kg BrdU. Mouse pups were killed at 5 days, 15 days, 30 days and 60 days post BrdU (Figure 3.1B).

For all BrdU experiments, mice were anesthetized with sodium pentobarbital (150 mg/kg) prior to transcardial perfusion with 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer (PB), pH 7.2 - 7.4. Tracheas were then dissected free and cleaned of connective tissue. Following a 3- to 4-hour post-fix, tracheas were stored in 20% sucrose in PB overnight. For both of these steps, the lumen of the trachea was flushed gently to ensure fluid filled the tracheal lumen. Tracheas were embedded in OCT, and 16 µm cross-sections were cut on a cryostat and thaw-mounted onto Fisher Plus slides (Fisher

Scientific, Waltham, MA). The slides were stored at -20°C at least overnight or until used for immunohistochemistry.

The slides were brought to room temperature, rinsed with 0.1 M phosphate buffer of pH 7.2 - 7.4 (PB) and dried on a slide warmer to enhance section adherence. Slides were rinsed in PBS and processed with appropriate antigen retrieval. For BrdU experiments, antigen retrieval consisted of incubation with 0.1% trypsin (Fisher

55

Scientific) diluted in distilled water at 37°C for 10 minutes, followed by 2N HCl for 20 minutes at 37°C and 20 minutes in 0.1 M Na Borate in distilled water at room temperature. Slides were rinsed in PBS between each solution. For the proliferative marker Ki67, antigen retrieval consisted of incubation with 10mM sodium citrate in distilled water for 1 hour at 95°C. After the slides were rinsed with PBS, non-specific staining was blocked for 1 hour in 1% Normal Donkey Serum 1% Triton in PBS

(blocking solution) and then incubated overnight with primary antibodies diluted in blocking solution.

For BrdU/α-gustducin double label immunofluorescence, primary antibodies were

1:200 biotinylated mouse anti-BrdU (Cat# 03-3940, Zymed Laboratories, South San

Francisco, CA) and 1:1000 rabbit anti-α-gustducin (raised against amino acids 93–112 of rat G α-gustducin, Cat# SC-395, Santa Cruz Biotechnology, Santa Cruz, CA) diluted in blocking solution. Following incubation with primary antibodies, slides were rinsed in

PBS and incubated with secondary antibodies, 1:250 Alexa Fluor568 streptavidin

(Invitrogen, Grand Island, NY) and 1:500 Alexa Fluor 488 donkey anti-rabbit

(Invitrogen), for 2 to 4 hours. No staining was observed in control experiments where rabbit anti-α-gustducin was omitted.

For Ki67 immunofluorescence, rabbit anti-Ki67 (raised against C terminus of Ki67, Cat# MA1-90584, Thermo Scientific) was diluted at 1:200. Following incubation with primary antibody, slides were rinsed in PBS and incubated with secondary 1:500 Alexa Fluor 549 donkey anti-rabbit (ABcam). Any residual rabbit immunoreactivity was then blocked overnight with unlabeled donkey anti-rabbit Fab

(Jackson Immuno Research) diluted 1:50 in blocking solution. Tissue was rinsed in PBS, incubated with rabbit anti-α-gustducin overnight and 1:500 Alexa Fluor 488 donkey anti- rabbit (Invitrogen) for 2 to 4 hours. Endogenous mouse IgG was then blocked overnight

56 with unlabeled donkey anti-mouse Fab (Jackson Immuno Research) diluted 1:50 in blocking solution.

The protocol for acetylated tubulin immunofluorescence was similar to Ki67, only no antigen retrieval was required. Mouse anti-acetylated tubulin (immunogen: acetylated tubulin from the outer arm of Strongylocentrotus purpuratus, Cat# T7451,

Sigma-Aldrich) was incubated at 1:1000 and detected with Alexa 568 donkey anti-mouse

(Cat# A11031, Invitrogen). Tissue from TRPM5-GFP mice was processed as above but with 1:200 rabbit anti-Ki67 and 1:600 chicken anti-GFP (Cat# AB16901, Millipore) primary antisera incubated together and 1:500 Alexa Fluor 568 donkey anti-rabbit

(Invitrogen) and 1:500 Alexa Fluor 488 donkey anti-chicken (Jackson Immuno Research) secondary antisera. Cytokeratin 14 immunofluorescence using mouse anti-Keratin 14

(ThermoFisher, 1:500) was similar to acetylated tubulin immunofluorescence or as previously described (Ghosh, Brechbuhl et al. 2011).

For CGRP immunofluorescence, rabbit anti-CGRP (immunogen SCNTA TCVTH

RLAGL LSRSG GVVKD NFVPT NVGSE AF, Cat# T-4032, Peninsula Laboratories,

San Carlos, CA) was diluted 1:500. For 5HT immunofluorescence, rabbit anti-5HT

(immunogen serotonin coupled to bovine serum albumin, Cat# 20080, Immuno Star,

Hudson, WI) was diluted 1:5000. Both CGRP and 5HT antibodies were detected with

Alexa Fluor 568 donkey anti-rabbit described above. To double-label tissue with α- gustducin and CGRP or 5HT, the rabbit anti-α-gustducin antibody described above was labeled with a Zenon Alexa 488 Rabbit IgG Labeling Kit (Cat# Z-25302, Invitrogen) and the resulting complex co-incubated with the secondary antibodies. For PGP9.5 immunofluorescence, guinea pig anti-PGP9.5 (immunogen GASSEDTLLKDAAKVCR,

Cat# GP14104, Neuromics, Edina, MN) was diluted 1:500 and detected with Alexa 594 goat anti-guinea pig (Cat# A-11076, Invitrogen). For Clara-cell secretory protein (CCSP)

57 immunofluorescence, goat anti-CCSP (immunogen 4x-His-tagged rat-CCSP, produced by

S. Reynolds) was diluted 1:10000 (Stripp, Reynolds et al. 2002) and detected with Alexa

568 donkey anti-goat (Cat# A11057, Invitrogen). This goat anti-CCSP antibody produces no staining in CCSP knockout mice (Stripp, Reynolds et al. 2002) and has been used extensively to label Clara cells in the tracheal epithelium (Ghosh, Brechbuhl et al. 2011,

Ghosh, Helm et al. 2011, Smith, Koch et al. 2012). Hoecsht 33342 (Invitrogen) or

DRAQ5 (ABcam) was used to counterstain some slides. All slides were rinsed with PBS, cover-slipped with fluoromount-G (Southern Biotech, Birmingham, AL) and stored at

4°C.

Tracheas for whole mounts were prepared and removed as for immunofluorescence but were cut down both sides and the ventral surface mounted on slides in fluoromount-G. Overlapping photographs were taken along the length of the tracheas at 40x using an epifluorescence microscope. Images were aligned and fitted together in Adobe Photoshop (Adobe Systems, Mountain View, CA) using a maximum intensity projection for regions of overlap.

Cell counts

Cells from at least 20 tracheal rings (cross-sections) from each animal were counted. Counted rings were selected from throughout the length of the trachea and separated from each other by at least 10 sections (160 µm) to adequately sample the full length of the specimen. Cells were counted as α-gustducin positive only if a non-reactive nucleus was visible in the center of the cell. For a cell to be considered double–labeled, the nucleus had to be immunoreactive for BrdU. The number of epithelial cells per µm of epithelium was determined by counting the number of epithelial cells in 20 DAPI-stained tracheal rings at each age. The total number of stained cells in a trachea ring was estimated by multiplying the density of cells per unit length of epithelium by the

58 circumference of the ring. The circumference of tracheal rings was measured by tracing the epithelia in a photograph of the ring with the freehand selection tool in ImageJ.

In vitro experiments

The procedure for producing air liquid interface (ALI) cultures from mouse tracheal epithelial cells has been described previously (You, Richer et al. 2002, Ghosh,

Brechbuhl et al. 2011). Briefly, mice were anesthetized with 250 mg/kg Avertin and tracheas dissected free of any adherent connective tissue. The tracheas were opened in the midsagittal plane and digested overnight in 0.15% Pronase (Sigma-Aldrich) in Ham’s

F-12 medium at 4°C. Tracheas were gently agitated to remove epithelial cells and the media with dislodged epithelial cells was centrifuged at 2000 rpm for 5 minutes at 4C.

The pellet of epithelial cells was resuspended in warm DMEM and transferred to a T-100 cell culture dish at 37C in a 5% CO2 incubator for 2-4 hours. Media containing epithelial cells was centrifuged, and the pellet resuspended in proliferation medium, consisting of

1:1 DMEM:Ham’s F-12 with 15mM HEPES, 2.5mM L-glutamine (Gibco 11330), 25 ng/ml Hydrocortisone (MP Biomedicals), 25 ng/ml Epidermal growth factor (BD

Biosciences), 0.1 mg/ml Cholera toxin (Sigma-Aldrich), 0.1 μg/ml Insulin-Transferrin-

Selenite (ITS, Gibco), 0.03 mg/ml Bovine pituitary extract (Gibco), 50 nM Retinoic acid

(Sigma-Aldrich) and 5% Fetal bovine serum (FBS) (Hy Clone) added. The resuspended cells were plated onto collagen-coated 24-transwell plates at 105 cells/cm2. Proliferation medium was replaced on the apical and basal side of the transwell every 48 to 72 hours until trans-epithelial resistance reached at least 4000 Ω. The medium was then removed from the apical side of the transwell and the proliferation medium on the basal side was replaced with differentiation medium: 1:1 DMEM:Ham’s F-12 with 15mM HEPES,

2.5mM L-glutamine (Gibco 11330), 50 nM Retinoic acid (Sigma-Aldrich) and 2%

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NuSerum (BD Biosciences 355100). The differentiation medium was changed every 48 to 72 hours (Figure 3.1C).

Figure 3.1: Diagram of BrdU treatment and culture methods. (A) Adult mice received three injections of 200mg/kg BrdU and were sacked at 5-day intervals to assess if BrdU had labeled BCs. (B) 5-day-old (P5) mouse pups received a single injection of 100mg/kg BrdU and were sacked at 5, 15, 30 and 60 days post injection to assess if BrdU had labeled BCs. (C) Dissociated tracheal epithelial cells are used to seed ALI cultures. A and B were produced using Servier Medical Art.

To label dividing cells, 10 µmol/L BrdU was added to the culture medium for a

12-hour period. Cultures were fixed for immunofluorescence by pipetting 500µL of 4%

PFA plus 20% sucrose in PBS onto the apical side of the transwell membrane.

Membranes were stored in PBS at 4°C. The immunofluorescence staining protocol for

ALI cultures was similar to the method for staining tracheal sections with the following exceptions: antigen retrieval was conducted at room temperature and culture membranes were mounted onto slides after staining. Cell density measurements were collected from

4 cultures per time point. Three distinct sampling fields spread evenly across each culture

60 membrane were selected for counting. Sampling fields from each culture were photographed at 20x and cells counted manually. The same criterion for counting cells was used as in the in vivo experiments.

Results

Identification of chemosensory brush cells

BCs are readily identifiable as a unique cell type within the tracheal epithelium.

They are elongate and typically have an apical process which reaches the lumen as well as one or more basal processes (Figure 3.2, Figure 3.3). Consistent with previous reports

(Krasteva, Canning et al. 2011, Tizzano, Cristofoletti et al. 2011), BCs are present at every level of the trachea from larynx to bronchi and co-express TRPM5, choline acetyltransferase (ChAT) and Gα-gustducin (Figure 3.2BC, Figure 3.3). BCs are not, however, immunoreactive for the ciliated cell marker, acetylated tubulin (ACT, (Figure

3.2D) or the Clara-like cell marker, Clara Cell Secretory Protein (CSSP, Figure 3.2E).

Although morphologically similar to neuroendocrine cells of the trachea, BCs are not immunoreactive for PGP9.5 (Figure 3.1A, Figure 3.3C), CGRP (Figure 3.2F) or 5HT

(Figure 3.2G), three markers of neuroendocrine cells. BCs are present throughout the tracheal epithelium and occur both as solitary cells (Figure 3.2ABDEG) and in clusters

(Figure 3.2CF). Cells that are similar to BCs occasionally occur in submucosal glands as indicated by Gα-gustducin immunoreactivity (Figure 3.2B, arrow). A similar distribution of BCs is demonstrated by GFP positivity in transgenic mice that harbor transgenes regulated by the ChAT- or TRPM5- promoters (Figure 3.2A-C).

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Figure 3.2: BCs are a distinct cell type in the tracheal epithelium (Part 1).

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Figure 3.2: BCs are a distinct cell type in the tracheal epithelium (Part 1). (A) Triple- labeled whole mount tracheal epithelium showing TRPM5-GFP (green), PGP9.5 (Blue) and acetylated tubulin (=ACT in red). BCs appear green. Nerve fibers (magenta) are immunoreactive for both PGP9.5 and ACT. Neuroendocrine cells are immunoreactive for PGP9.5 (Blue), while cilia appear red, being immunoreactive for ACT. (B) BCs co- express Gα-gustducin (red) and ChAT (green) and occur within the tracheal epithelium (arrow heads) and in submucosal glands (arrow) (C) Most BCs are immunoreactive for both Gα-gustducin (red) and TRPM5 (green) and have multiple processes. (D) Gα- gustducin (green) immunoreactive BCs are not immunoreactive ACT (magenta), a marker for ciliated cells. (E) Gα-gustducin (green) immunoreactive BCs are not immunoreactive for CCSP (red), a marker for Clara-like cells. (F and G) TRPM5 (green) expressing BCs appear morphologically similar to neuroendocrine cells but are not immunoreactive for known neuroendocrine cell markers: (E) CGRP (red) and (F) 5HT (red). (B-G) Counter stained with DRAQ5 shown in Cyan. Scale bars = 10μm.

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Figure 3.3: BCs are a distinct cell type in the tracheal epithelium (Part 2). (A) BCs co- express α-gustducin (red) and ChAT (green); counterstained with DAPI in blue. (B) Bipolar BC immunoreactive for both α-gustducin (red) and TRPM5 (green). Counterstained with DRAQ5 in cyan. (C) α-gustducin (green) expressing BCs appear morphologically similar to neuroendocrine cells but are not immunoreactive for known neuroendocrine cell marker PGP9.5 (Red). Counter stained with DRAQ5 in cyan. (D) Trachea from 5-day-old mouse, injected with BrdU and sacrificed after 5 hours. BrdU (green) labeled nuclei of Cytokeratin 14 (K14, magenta) expressing basal cells or nuclei closely associated with Cytokeratin 14 expressing basal cells. Inserts show single color channel. Scale bars=10µm

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Chemosensory brush cells do not turnover in adult mice

In adult mice, BrdU immunoreactive nuclei were present throughout the depth of the tracheal epithelium 5, 10, 15 and 20 days after BrdU injection (Figure 3.4A, Figure

3.5A). BrdU+ nuclei were more common near the basement membrane at shorter time intervals. Five days after BrdU injection, 2.37±0.03% of tracheal epithelia cells were labeled (Figure 3.4B). At this rate, 100% of cells would be replaced in 211 days assuming no premature cell death. The frequency of BrdU+ cells decreased to 1.86 ± 0.05% on day

10; 1.07 ± 0.03% on day 15; and, 0.92 ± 0.07% on day 20. Extrapolation of these values predicts that epithelial cell lifespan is greater than 211 days and is consistent with findings reported by others (Basbaum and Jany 1990, Rawlins and Hogan 2008). In the

16 mice utilized in this experiment, 2849 BCs were examined. No BrdU positive BC were detected at any time (Figure 3.4B). Thus, BrdU labeling revealed no evidence for replacement of BCs in adult epithelium. We conclude that BC replacement, if it occurs at all in healthy adult mice, is significantly slower than that of the surrounding epithelium

(Figure 3.4B, p<0.001 by Chi-square test).

Perinatal generation of chemosensory brush cells

Since BCs are a relatively stable population in adults, we investigated the origin of BCs during development. The trachea lengthens and enlarges during postnatal development (Van Winkle, Fanucchi et al. 2004) resulting in a substantial increase in epithelial surface area (Figure 3.3AB). If BCs are a static population that is established prior to birth, the density of BCs in the trachea should decrease as the trachea enlarges.

Conversely, if the density of BCs is constant during this period of expansion, then new

BCs must be added over time. To evaluate this question, we imaged whole-mounted tracheas from TRPM5-GFP mice. The density of BCs did not change significantly from post-natal day (p) 5 to p30 implying significant cell addition occurs during this period

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(Figure 3.3C; 43.20 ± 4.92 BCs/mm2 on p5, 38.40 ± 4.26 BCs/mm2 on p10, 42.80 ± 6.95

2 2 BCs/mm on p15, and 57.20 ± 6.09 BCs/mm on p30, one-way ANOVA, F(3,16)=2.09, p=0.14).

To test if BCs themselves are mitotic during postnatal growth, we stained the trachea for the proliferateration marker Ki67, which is present during all phases of the cell cycle except G0 (Scholzen and Gerdes 2000). Numerous epithelial cells, but no Gα- gustducin-immunoreactive BCs were immunoreactive for Ki67. The mitotic index (% cells immunoreactive for Ki67) within the tracheal epithelium decreased significantly as mice aged from p10 to p65 (Figure 3.4D, one-way ANOVA, F(3,8)=23.80, p<0.001; p10,

5.75 ± 0.06%; p20, 4.11 ± 0.11%; p35, 2.66±0.05%; p65, 2.57±0.05%). In the 12 mice utilized in this experiment, 415 BCs and 3767 Ki67 positive nuclei were examined. At no time were Ki67 and Gα-gustducin double-positive cells detected (Figure 3.4C).

To test if BCs are added from a proliferative population during early development, we injected mice with BrdU at p5 and examined the tracheas for labeled

BCs at 5 hours, and at 5, 15, 30 and 60 days post-injection (Figure 3.1B). In tracheas examined 5 hours after BrdU injection, the majority of BrdU labeled cells expressed cytokeratin 14 or were adjacent to cytokeratin 14-expressing basal cells (Figure 3.3D).

No BCs were BrdU-labeled at this time. This finding is consistent with previous reports that the main tracheal progenitor is a basal cell which expresses cytokeratin 14 (Reynolds and Malkinson 2010, Brechbuhl, Ghosh et al. 2011, Ghosh, Brechbuhl et al. 2011). As early as 5 days post-injection (chase day 5 = p10), BCs with BrdU labeled nuclei (Figure

3.4E) were detected along with BrdU-labeled non-BC epithelial cells. Apparently, these labeled BCs arise from the proliferative basal cells labeled shortly after the BrdU injection. The fraction of BCs labeled with BrdU increased with post-injection time,

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Figure 3.4: BCs are a static population in the adult trachea but are generated by a population of proliferative progenitor cells during perinatal development.

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Figure 3.4: BCs are a static population in the adult trachea but are generated by a population of proliferative progenitor cells during perinatal development. (A) Double-labeled image with Nomarski overlay showing BrdU (red) labeled epithelial cell nuclei near an unlabeled Gα-gustducin (green) immunoreactive BC in an adult mouse, 20 days after BrdU treatment. (B) Percentage of tracheal epithelial cells and BCs labeled with BrdU at 5 to 20 days after BrdU treatment of adult mice. Bars depict mean±SEM (n=4) for each point. (C) Double-labeled image showing Ki57 (red) labeled epithelial cell nuclei near a Gα-gustducin (green) immunoreactive BC in a trachea from a 35 day old mouse. (D) Percentage of perinatal tracheal epithelial cells and BCs immunoreactive for the mitotic marker Ki67. Bars depict mean±SEM (n=3) for each time. (E) Double-labeled image with Nomarski overlay showing BrdU (red) labeled Gα-gustducin (green) immunoreactive BC (arrow) in a 35 day old mouse, 30 days after BrdU treatment. (F) Percentage of perinatal tracheal epithelial cells and BCs labeled with BrdU at different times after injection of 5day old mice. Bars depict mean + SEM (n=4) for each time point. **p<0.01, *** p<0.001 by Chi-squared-test. Inserts show single color channels. Scale bars = 10μm.

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Figure 3.5: Chemosensory brush cells are present in the lower airways of prenatal mice (A) Whole-mounted tracheas from TRPM5-GFP (green) mice 5 (P05), 10 (P10), 15 (P15) and 30 (P30) days after birth. Red channel is background auto-fluorescence to display gross anatomy of tracheal rings. Scale bar = 1mm. (B) Surface area of the tracheas increased during the perinatal period. (C) Density of BCs does not significantly change during the perinatal period. Bars depict mean + SEM (n=5) for each time point. suggesting a slow addition of new BCs from the proliferative basal cell pool. In contrast, the fraction of BrdU-labeled non-BC epithelial cells decreased (Figure 3.4F). On chase day 5, the proportion of BrdU-labeled to total BCs was significantly lower than the proportion of labeled to total non-BC epithelial cells (0.38 ± 0.38% of BCs compared to

4.16 ± 0.29% of non-BC epithelial cells, n=4, Chi squared analysis, p<0.01). By chase day 30, the proportion of BrdU-labeled to total BCs (2.46 ± 0.93%) was not significantly different from the proportion of BrdU-labeled to total non-BC epithelial cells (2.12 ±

0.32%). On chase day 60, the proportion of BrdU-labeled BCs was significantly higher than the proportion of non-BC epithelial cells labeled by BrdU (7.71 ± 0.12% of BCs compared to 1.44 ± 0.03% of other epithelial cells, n=4, Chi squared analysis, p<0.001).

The BC progenitor was labeled by BrdU on p5 but BC progeny were not detected for 5

69 days. Taken together, these findings indicate that the BC progenitor is proliferative at p5, but that differentiation of BCs occurs over a period of weeks.

New chemosensory brush cells can be generated from adult tracheal epithelial progenitors

To test whether the adult tracheal epithelium retains the capacity to generate new

BCs, we employed air-liquid-interface (ALI) cultures (Figure 3.1C). ALI cultures are produced by seeding dissociated tracheal epithelial cells at low density to model epithelial regeneration after injury (You, Richer et al. 2002, Brechbuhl, Ghosh et al. 2011,

Ghosh, Helm et al. 2011, Smith, Koch et al. 2012). These epithelial cells are grown to confluence on a porous polycarbonate membrane and differentiate into an epithelial layer that recapitulates the anatomy and gene expression of native mouse trachea (You, Richer et al. 2002, Brechbuhl, Ghosh et al. 2011, Smith, Koch et al. 2012). This model can be used to determine the cell types generated by adult tracheal facultative progenitor cells

(You, Richer et al. 2002, Brechbuhl, Ghosh et al. 2011) and was adapted to evaluate tissue stem cell differentiation (Ghosh, Brechbuhl et al. 2011).

Immediately after transition to differentiation conditions, rare Gα-gustducin immunoreactive cells were noted (Figure 3.6A) but constituted only 0.01 ± 0.01% of the total population (Figure 3.6G). In culture, as in vivo, BCs expressed both Gα-gustducin and TRPM5 (Figure 3.5B) but were not immunoreactive for acetylated tubulin (Figure

3.6C). In vitro BCs were multi-polar with two to three processes of various lengths

(Figure 3.6ABC) similar to their appearance in vivo in adult trachea. Doublet BCs, i.e., pairs of adjacent Gα-gustducin-positive BCs, were common (Figure 3.6E-F). Closely- spaced BCs were also present in vivo but were not as intimately associated as those in culture (c.f Figure 3.2C and Figure 3.6EF).

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The overall cell density within the cultures is stable after the switch to differentiation conditions (Figure 3.6G), but the proportion of BCs increases significantly

(one-way ANOVA, F(4,85)=62.71, p<0.001) to 0.38 ± 0.09 % at ALI day 3, 0.93 ± 0.22% at ALI day 7 and 1.12 ± 0.18% by ALI day 14 (Figure 3.6G). To test the possibility that this increase in BC density was due to cell division, BrdU was added to the cultures on proliferation day 3 (2 days before the switch to ALI) or on ALI day 0. No BrdU labeled

BCs were present in cultures fixed immediately after the 12 hour BrdU pulse, indicating that differentiated BCs are not proliferative. BrdU labeled BCs are, however, present on

ALI day 14 (chase day 16 or 14; Figure 3.6D) indicating that BCs differentiate from a proliferative population labeled in culture some days previous during cell division. It is possible that the proliferative cells undergo asymmetric divisions, so that one daughter cell remains proliferative while the other assumes a differentiated cell fate, potentially becoming one of a variety of cell types including ciliated, Clara-like or BC.

Determination of an exact lineage relationship within the epithelial population will require further study.

Significantly more BCs were labeled when BrdU was added on ALI0 (Figure

3.6H, t-test, p<0.001), than when BrdU was added on proliferation day 3 suggesting that progenitor cells that proliferate on proliferation day 3 are biased toward population expansion. In contrast, progenitors that proliferate after transition to ALI have the potential to generate differentiated BCs. Finally, continued cell divisions between proliferation day 3 and onset of ALI would likely dilute the BrdU label in the proliferative population resulting in no detectable label in later-forming BCs.

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Figure 3.6: New BCs can be generated from adult tracheal epithelium in vitro.

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Figure 3.6: New BCs can be generated from adult tracheal epithelium in vitro. (A) A Gα-gustducin (red) immunoreactive BC in tracheal epithelial cell culture on ALI day 7. Cultured BCs have multiple processes like their in vivo counterparts. (B) BCs in cultures produced from TRPM5-GFP (green) transgenic mice were immunoreactive for Gα-gustducin (red) and expressed GFP. ALI day 7. (C) Gα-gustducin (green) immunoreactive BCs were negative for the ciliated cell marker acetylated tubulin (ACT, magenta). ALI day 7. (D) Image of a double-labeled cell showing a BrdU (green) and Gα-gustducin (red) double immunoreactive BC (arrow). BrdU also labeled other cells in culture. ALI day 14. (E) Several doublet Gα-gustducin (red) immunoreactive BCs, suggestive of daughter cells, were observed culture. ALI day 1. (F) Double labeled image showing an Gα-gustducin (red) immunoreactive doublet where both nuclei are labeled with BrdU (green). ALI day 14. Counterstained with DAPI shown in blue. Scale bars = 10μm. (G) Tracheal epithelial cell density (red line) and the faction of BCs (green line) over time in ALI cultures. Points depict mean ± SEM (n=3) for each time. (H) Fraction of cultured BCs labeled when dosed with BrdU either during the proliferation period (Pro 3) or at ALI day 0. *** p<0.001 by t-test. Bars depict mean ± SEM (n=3) for each point

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Thus, the present study faithfully evaluates early BC generation but may not detect BCs that are generated as a consequence of multiple rounds of progenitor proliferation.

Taken together, the in vitro results indicate that the adult tracheal epithelium has the capacity to generate new BCs, although few are generated under normal healthy conditions. Since the ALI culture system tests the ability of tracheal facultative progenitor cells to reconstitute the tracheal epithelium after disruption (You, Richer et al.

2002, Brechbuhl, Ghosh et al. 2011), generation of substantial numbers of BCs in the adult epithelium is likely dependent on epithelial injury or damage. While ALI cultures mimic many aspects of the naphthalene injury model (Brechbuhl, Ghosh et al. 2011), in vitro models may not always faithfully recapitulate in vivo circumstances, and so these data must be viewed with that caveat in mind.

In summary, our results indicate that BCs are a relatively static population in the healthy adult trachea and that these cells show no evidence of the turnover typical of the surrounding epithelial cells. The majority of BCs present in the healthy adult mouse is generated perinatally, while the trachea is expanding. Although tracheal BCs are a stable population, adult epithelial progenitor cells (Brechbuhl, Ghosh et al. 2011, Ghosh,

Brechbuhl et al. 2011) retain the ability to generate new BCs in vitro suggesting that BCs can be replaced as part of an epithelial repair response.

Discussion

Tracheal brush cells are a stable population

Respiratory epithelia, including the tracheal epithelium, are at risk of damage by inhaled particulates, toxins and pathogens. In the normal healthy trachea, ciliated cells and Clara-like cells have the most exposure to the airway lumen and are replaced approximately every 6-9 months. Nascent luminal cells are derived from basal cell progenitors which proliferate and then differentiate into ciliated and Clara-like cells. In

74 contrast to luminal cells, the tracheal BC population is essentially static. However, our in vitro studies demonstrate that BCs can be regenerated if they are lost due to tracheal damage. Additionally, BC lifespan, and consequently, their turnover rate, is distinct from that of other tracheal epithelial cell types. These data raise interesting questions regarding the mechanisms that determine BC longevity.

The lack of BC cell turnover is unusual for chemosensory epithelia. Most chemosensory cells are replaced every 2-4 weeks (e.g., taste buds, cells and even solitary chemosensory cells of the nasal epithelium) (Beidler and Smallman

1965, Moulton 1974, Farbman 1980, Mackay-Sim and Kittel 1991, Gulbransen and

Finger 2005, Hamamichi, Asano-Miyoshi et al. 2006). However, some chemosensory cells, including those of the carotid body, are quite stable (Wang, Olson et al. 2008).

These cells are internal chemosensors (Milsom and Burleson 2007) and their lack of turnover may be due to their protected intraepithelial position and minimal interaction with the luminal environment. Similarly, other sensory cells in protected endorgans are not replaced in a normal healthy individual, (e.g., photoreceptors (Zeiss and Johnson

2004), auditory hair cells (Lopez-Schier 2004), and Merkel cells (Vaigot, Pisani et al.

1987)). The stability of murine tracheal BCs may also reflect the relatively protected environment in which they reside. The rodent nose provides an excellent filtering mechanism, leaving the trachea relatively free of inhaled toxins. In contrast, the is not as efficient at this protective function (Harkema, Carey et al. 2006). Thus, the human tracheobronchial BC population is likely to be more susceptible to damage and therefore is less stable than its murine counterpart. Following a similar line of logic,

SCCs in the nasal cavity of adult mice are likely to be exposed to a higher concentration of inhaled toxins than are tracheal BCs. Thus, the finding that nasal SCCs turnover every

75 few weeks, (Gulbransen and Finger 2005) while BCs are stable, may reflect cell location rather than cell-intrinsic differences in maximal lifespan.

New brush cells are generated during tracheal growth

During the perinatal growth period, the surface area of the trachea increases and new epithelial cells are added. In agreement with previous reports (Basbaum and Jany

1990), we find that the mitotic index of tracheal epithelium during the postnatal period is higher than in adult mice (Figure 3.3D). The density of ciliated and Clara-like cells remains constant even as new cells are generated (Van Winkle, Fanucchi et al. 2004).

Likewise, new BCs are generated during this period (Figure 3.4EF), resulting in a constant proportion of BCs to total cells (Figure 3.6). The constant BC, ciliated and

Clara-like cell density suggests that signaling within the epithelium regulates the rate at which various tracheal cell types are generated during tracheal growth.

During adulthood, tracheal dimensions are constant and the BC population is stable (Figure 3.4B). While non-BC epithelial cells are replaced during adulthood, their lifespan increases considerably with aging (Basbaum and Jany 1990, Rock, Randell et al.

2010). Conversely after injury, the adult respiratory epithelium increases the rate at which new epithelial cells are produced (Hong, Reynolds et al. 2004, Hong, Reynolds et al.

2004, Cole, Smith et al. 2010) and new cell addition results in the regeneration of all of the major respiratory cell types. Our results demonstrate that BCs, like ciliated and Clara- like cells, can be regenerated after damage (Figure 3.6DGH). The airway epithelium is a remarkably resilient tissue, capable of reconstituting the diverse cell types that compose the tissue from a small reservoir of progenitor cells (Basbaum and Jany 1990, Reynolds,

Giangreco et al. 2000). The idea that tracheal progenitor cells are capable of regenerating

BCs is completely consistent with current models of airway plasticity.

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BC function may play a role in regulating the plasticity of the tracheal epithelium.

BCs respond to bacteria metabolites by synthesizing and releasing acetylcholine

(Krasteva, Canning et al. 2011, Krasteva, Canning et al. 2012) (Figure 3.2B, Figure

3.3B). The acetylcholine released by BCs lying close to mitotic basal cells (Figure 3.4C) may act in paracrine fashion on basal cell nicotinic acetylcholine receptors which modulate proliferation and differentiation of the mitotic population (Maouche, Polette et al. 2009). This model provides a mechanism by which BC activation could result in changes in proliferation and differentiation of tracheal progenitor cells.

The main progenitor of neonatal and adult tracheal epithelial cells is a cytokeratin

(K) 5- and 14-expressing basal cell (Daniely, Liao et al. 2004, Reynolds and Malkinson

2010, Brechbuhl, Ghosh et al. 2011, Ghosh, Brechbuhl et al. 2011). In the adult tracheal epithelium, basal cells self-renew (Reynolds and Malkinson 2010) while also producing

CCSP+ Clara-like secretory cells (Ghosh, Brechbuhl et al. 2011). Clara-like cells then are capable of producing ciliated cells in normal conditions (Ghosh, Brechbuhl et al. 2011) or undergoing injury-induced metaplasia to become mucus-secreting cells (Reynolds and

Malkinson 2010). Ciliated cells and mucus secreting cells appear to be terminally differentiated, whereas Clara-like cells are capable of self-renewal (Reynolds and

Malkinson 2010). Similarly, our data indicate that BCs are also terminally-differentiated cells.

BCs have yet to be placed within a tracheal epithelial cell linage. The lack of reliable BC markers and their scarcity (1% of total epithelial cells) has hindered previous examination of lineage relationships (Reid, Meyrick et al. 2005). We found that nearly all

BrdU-labeled cells expressed K14 or were adjacent to K14+ basal cells within 5 hours of injection (Figure 3.3D) while BrdU+ BCs appear with longer chase times. Although, our results suggest that the K14- and K5-expressing basal cells give rise to BCs, as well as

77 the more common cell types of the tracheal epithelium, lineage-tracing in vivo will be needed to establish a lineage relationship.

Reliable markers for chemosensory brush cells

The lack of reliable markers for BCs has presented a significant impediment to elucidating details of their biology (Reid, Meyrick et al. 2005). Initial speculation that

BCs were chemosensory was provoked by ultrastructural analysis of tracheal epithelium

(Luciano, Reale et al. 1969). This observation was not confirmed until 40 years later

(Krasteva, Canning et al. 2011, Krasteva, Canning et al. 2012), largely because BCs could not be identified among the more common cells types in tracheal epithelium (Reid,

Meyrick et al. 2005).

In the current study, we used Gα-gustducin as a marker for BCs because this protein was detectable even after the harsh antigen retrieval methods required to detect

BrdU. Gα-gustducin has been used extensively as a marker for bitter, sweet and umami sensitive type II taste receptor cells, including several BrdU-dating studies (Cho,

Farbman et al. 1998, Nguyen, Reyland et al. 2012). Similarly, Gα-gustducin is a reliable marker for nasal solitary chemosensory cells.

The majority of BCs are immunoreactive for Gα-gustducin, but a small fraction can lack one or more of the taste-related markers (Tizzano, Cristofoletti et al. 2011,

Krasteva, Hartmann et al. 2012). In the auditory tube, 3% of ChAT-expressing BCs are not immunoreactive for Gα-gustducin (Krasteva, Hartmann et al. 2012). Likewise, the occasional tracheal BC that does not express all of the phenotypic markers has also been noted (Krasteva, Canning et al. 2011). While this small fraction of cells could be explained by experimental variation, other possibilities exist. In the taste system, sub- populations of receptor cells utilize different downstream transduction elements (Tizzano,

Dvoryanchikov et al. 2008). Thus, in BCs, as in taste receptor cells, different second

78 messengers could be utilized by different subpopulations. Alternatively, these “atypical” non-Gα-gustducin BCs could be immature cells which have yet to express the full complement of chemosensory transduction proteins.

A previous study reported that ciliated cells of cultured human respiratory epithelium are immunoreactive for Gα-gustducin (Shah, Ben-Shahar et al. 2009), although others noted the absence of Gα-gustducin in ciliated cells of the respiratory epithelium of rat (Merigo, Benati et al. 2012). Likewise, we never observed cilia that were immunoreactive for Gα-gustducin in our in vivo (Figure 3.2D) or in vitro (Figure

3.6C) preparations of mouse tissue. This disparity may represent differences in species- specific expression patterns or differences between cultured and native ciliated cells. The distinct morphology of ciliated cells and BCs is sufficient to ensure that no ciliated cells were counted as BCs in the current study (Figure 3.1D, Figure 3.3C).

Conclusions

In summary, BCs in healthy adult mice are a static population which is generated during perinatal development. This is unusual for epithelial cells. Despite the lack of turnover in adult epithelium, BCs can be regenerated from proliferative epithelial cells following tracheal damage allowing for full reconstitution of the tracheal epithelium.

Acknowledgements

We thank Robert Margolskee (Monell Chemical Senses Center) for providing the

TRPM5-GFP mice and Sukumar Vijayaraghavan (University of Colorado School of

Medicine) for providing the ChAT-tauGFP mice. Additionally, we thank Russell Smith for his assistance with BrdU-cytokeratin 14 immunohistochemistry and Jennifer Strafford for statistical advice. Parts of this manuscript were presented at the 2010, 2011, 2012

79 annual meetings of the Association of Chemoreceptive Sciences and 2011 American

Thoracic Society international conference.

Grants

This work was supported by National Institutes of Health Grants R01 009820

(T.E.F.), P30 DC004657 (D. Restrepo) and RO1 HL075585 and Supplement HL075585-

S1 (S.D.R.).

80

CHAPTER IV

CONCLUSION

Solitary chemosensory cells (SCCs) and brush cells (BCs) along the respiratory tract provide a warning system by which the lumen of the airway can be monitored for potentially toxic chemicals. Chapter II described experiments, which established that

SCCs, like BCs, require nicotinic acetylcholine receptors and cholinergic neurotransmission to trigger inflammation. Chapter III described experiments, which established that BCs are not renewed at the same rate as the surrounding tracheal epithelial cells but, rather, are generated shortly after birth and are a relatively static population in healthy adults. These new findings fill in several gaps that existed in our current understanding of SCC and BC biology and raise several interesting avenues for future research.

The elucidation of the basic function of SCC’s and BC’s is an exciting new development in the fields of sensory neuroscience and pulmonology. These findings raise thrilling new questions about the role of these cells in the epithelium. As more data are collected, BCs and SCCs appear to have more similarities than differences, raising the possibility that they are in fact the same cell type. Perhaps taste cell-like chemosensors are a basic component of muco-cilliary epithelium and part of that tissue’s innate defense system against pathogens? Both BCs and SCCs activate nAChRs on nerve fibers but

AChRs are also found on cells throughout the epithelium, raising the possibility of paracrine ACh signaling (Kummer, Lips et al. 2008). Additionally, since new SCCs and

(under some circumstances) new BCs can be generated (Gulbransen and Finger 2005), these cells must utilize a currently unknown signaling molecule to attract nerve fibers and form synapses. Finally, since innervation appears to be one of the few differences between the two cell types (Krasteva, Canning et al. 2011, Tizzano, Cristofoletti et al.

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2011), perhaps understanding how SCC and BC innervation is regulated will help establish if the cells are the same type.

Are Sccs And Bcs The Same Thing?

SCCs and BCs are more similar than different. Both SCCs and BCs are capable of triggering irritation and inflammation along the length of the airway (Finger, Bottger et al. 2003, Krasteva, Canning et al. 2011)(Chapter II). The same molecular machinery is present in both cell types and allows for the detection of bitter compounds and bacterial metabolites (Finger, Bottger et al. 2003, Tizzano, Gulbransen et al. 2010, Krasteva,

Canning et al. 2011) (Chapter II, Chapter III). Irritation induced by SCCs and BCs is dependent on nicotinic cholinergic neurotransmission (Krasteva, Canning et al. 2011)

(Chapter I). However, there are several differences in ultrastructure, renewal and innervation between the cells. Since, SCCs and BCs have many characteristics in common, but several discrete differences, any attempt to establish commonality of these cells must start with a discussion of the criteria used to establish that cells represent different types.

Cells of the same type will share a common morphology, function, origin and molecular character. If SCCs and BCs represent the same cell type, then these cells should share the same basic morphology and anatomical arrangement within the epithelium. Most importantly, cells of the same type should achieve a comparable function through common physiological mechanisms. Additionally, cells belonging to the same category should have similar origins and develop from homologous tissues. Finally, since a cell’s morphology and physiology are largely dictated by the proteins made, cells of the same type should have a similar gene expression or molecular character.

82

Morphology and anatomy

SCCs and BCs have a generally similar morphology and similar relationship with the surrounding cells of the epithelium. SCCs and BC have both been described as bottle- shaped and both possess multiple intricate processes that wend throughout the epithelium

(Finger, Bottger et al. 2003, Krasteva, Canning et al. 2011, Tizzano, Cristofoletti et al.

2011) (Chapter II, Chapter III). One of these processes usually penetrate into the lumen of the respiratory tract where they terminate in a microvillus tuft (Finger, Bottger et al.

2003, Krasteva, Canning et al. 2011). While both SCCs and BCs possess a microvillus tuft, some have observed that the SCC microvillus tuft appears shorter than that of the

BC (Finger, Bottger et al. 2003, Krasteva, Canning et al. 2011). However, there are several caveats to this already subtle difference. First, the microvillus tufts of SCCs have only been observed via transmission electron micrographs (Finger, Bottger et al. 2003).

The characteristic “brush” of BCs is best observed by scanning electron microscopy, which avoids artifacts that might occur as a result of sectioning the epithelium (Brody

2005). Secondly, the nasal cavity purifies inhaled air before it reaches the trachea

(Harkema, Carey et al. 2006). As such, perhaps the shortened microvillus tufts observed on SCCs are a result of damage due to its exposed location, not a difference due to different biological properties. Finally, even if there are differences in the size or length of the “brush” between SCCs and BCs, this difference would hardly be significant enough to establish them as two different types of cells.

A more pronounced difference between SCCs and BCs than the size of each cell’s microvillus tuft are differences observed in innervation between SCCs and BCs. Both

SCCs and BCs are innervated by peptidergic nociceptive fibers, but while nerve fibers intimately intertwine SCCs, they make only passing contact with BCs (Krasteva,

Canning et al. 2011, Tizzano, Cristofoletti et al. 2011). While this is certainly a difference

83 in the anatomy associated with cells, this distinction may represent a difference in trigeminal and vagal nociceptive fibers, not a difference in the taste cell-like chemosensors of the nose and trachea. Specifically, the innervation of the nasal epithelium is denser than that of the trachea (Gulbransen, Silver et al. 2008) (Chapter III), so there are more pain fibers present in the nasal epithelium to innervate SCCs.

Furthermore, whatever differences there are between the innervation of the trachea and nose, the innervation of both cells is sufficient to allow for their stimulation to cause changes to respiratory physiology by activating nerves (Finger, Bottger et al. 2003,

Tizzano, Gulbransen et al. 2010, Krasteva, Canning et al. 2011, Krasteva, Canning et al.

2012).

Physiology and function

There are very few differences between the physiology of SCCs and BCs. Both cells are stimulated by bitter substances and bacteria metabolites through the canonical bitter taste transduction cascade (Tizzano, Gulbransen et al. 2010, Krasteva, Canning et al. 2011). Activation of SCCs or BCs via this pathway results in protective respiratory reflexes indicative of irritation (Tizzano, Gulbransen et al. 2010, Krasteva, Canning et al.

2011). Additionally, inflammation or irritation induced by SCCs or BCs is blocked when animals are treated with nAChR antagonists (Krasteva, Canning et al. 2011) (Chapter II).

For cells to be the same type, they should have the same function and utilize the same general mechanism to achieve this function. Since SCCs and BCs both utilize the same transduction pathway to detect potentially noxious compounds and induce the same protective respiratory reflexes, they are functionally the same cell type.

Development

In addition to having similar functions, cells of the same type should have a similar origin. While only four publications, including chapter III, have explored the

84 developmental biology of SCCs and BCs, the information that is available suggests that

SCCs and BCs originate in the same manner (Gulbransen and Finger 2005, Gulbransen,

Silver et al. 2008, Ohmoto, Yamaguchi et al. 2013) (Chapter III). Both SCCs and BCs have been identified in the airway of neonatal mice. In contrast to taste receptor cells, in vivo experiments with SCCs and in vitro experiments with BCs suggest that both SCCs and BCs arise even in the absence of innervation (Gulbransen, Silver et al. 2008)

(Chapter III). However, while the two cells appear in the absence of innervation, the rate at which the two cells arise and are replaced appears to be different (Gulbransen and

Finger 2005) (Chapter III).

The main developmental difference between the two cell types appears to be that

SCCs are replaced with the surrounding epithelium, while BCs are a relatively static population in the trachea. However, this difference, like the possible differences in the morphology of the microvillus tuft, might be due to the amount of damage endured by the nasal and tracheal epithelium, rather than a difference between the cells (Harkema,

Carey et al. 2006). Since the nasal epithelium endures more damage than the tracheal epithelium, the turnover rate of nasal epithelium is higher for all the cell types of the nose, including SCCs (Basbaum and Jany 1990). This difference in cell turnover is one of the few distinctions between SCCs and BCs, but other possible differences have yet to be explored.

If SCCs and BCs were shown to originate from distinct cell lineages, they could be classified as two different cell types. Currently, no experiments have looked at the developmental lineages of SCCs. Additionally, the only information on BCs lineages we have comes from experiments demonstrating that BrdU labels cytokeratin-14 immunoreative basal cells after short chase periods, before any BrdU labeled BCs are observed (Chapter III). To address the question of SCC and BC lineage, future studies

85 should utilize a Cre-Lox recombination system to label the progeny of cytokeratin-5 and

-14 positive basal cells and determine if these cells give rise to both classes of taste cell- like airway chemosensors (Ghosh, Brechbuhl et al. 2011). If SCCs and BCs are produced by a similar stem cell found throughout the epithelium of the conducting airways, then this would support the hypothesis that SCCs and BCs are, in fact, the same cell type.

Conversely, demonstrating that SCCs and BCs originate from separate cell lineages would be strong evidence that SCCs and BCs are distinct cells types.

Molecular character

While morphology, function and developmental origin are all criteria that distinguish different types of cells, differences in all these factors are largely determined by gene expression. For example, the microvillus tuft present in both SCCs and BCs exists because both cells produce the structural protein villin (Hansen 2007, Krasteva,

Canning et al. 2011). Additionally, SCCs and BCs would be unable to respond to the bitter substances if the elements of the canonical bitter taste transduction cascade—T2Rs,

α-gustducin, PLCβ2, IP3R3 and TRPM5—were not present (Tizzano, Gulbransen et al.

2010) (Chapter II). Furthermore, without the presence of the synthetic and vesicular packaging enzymes, ChAT and VAChT, neither SCCs nor BCs would be capable of activating nAChRs on the post-synaptic nerve fiber by releasing ACh (Krasteva, Canning et al. 2011)(Chapter II). While little is known about the ontology of SCCs and BCs, it is likely that the expression of one or more genes triggers a developmental program, which produces airway chemosensory cells, similar to the events that control the development of some gut chemoreceptor cells (Delgiorno, Hall et al. 2013, Ohmoto, Yamaguchi et al.

2013). In short, if cells of the same type share similar morphology, function and ontology, then they will express the same genes.

86

For SCCs and BCs to be the same cell type, they must express the same genes.

Next generation sequencing technology allows us to measure the genes transcribed by a cell. SCCs and BCs can be separated from the surrounding epithelium by using fluorescently activated cell sorting (FACS) to purify these cells from transgenic mice, where a taste transduction gene, like TRPM5, drives GFP expression (Bezencon, le

Coutre et al. 2007). Once purified, the transcriptome of these cells can be quantified by next generation RNA-sequencing and compared to determine if they are the same cell type. The critical phase of this experiment would be determining by what degree the transcriptome of SCCs and BCs could differ and still be classified as the same cell type.

Since nasal and tracheal respiratory epithelium both contain ciliated cells that are generally considered to be the same cell type, a comparison of the transcriptome of nasal and tracheal ciliated cells would provide a baseline for SCCs and BCs to be compared against. Next generation RNA-sequencing provides a method by which differences between the two populations of cells can be quantified.

Paracrine Signaling

In addition to SCCs and BCs, several other cell types compose the respiratory epithelium. Ciliated cells, secretory cells and basal cells compose the majority of epithelium (Rock, Randell et al. 2010). Each of these three cell types plays a unique role in maintaining the health of the epithelium. Furthermore, the function of each of these cells is capable of being modulated by chemical signals released from other nearby cells, a phenomenon known as paracrine signaling (Kummer, Lips et al. 2008).

ACh is a potent paracrine signal to all three of the most common cell types of the respiratory epithelium (Kummer, Lips et al. 2008). Ciliated cells are responsible for clearing the airway and beat faster when exposed to ACh (Kummer and Lips 2006).

Additionally, secretory cells release their bio-active and potentially protective contents of

87 their secretory granules on stimulation with ACh (Kummer and Lips 2006). Basal cells are widely thought to be sensitive to stimulation by ACh, although there are conflicting reports as to the exact effect of this stimulation (Metzen, Bittinger et al. 2003, Maouche,

Polette et al. 2009). SCCs and BCs represent a possible reservoir of ACh in the epithelium that could be utilized for paracrine signaling.

On stimulation with bitter irritants and xenobiotics, SCCs and BCs may release

ACh in a paracellular manner to trigger protective responses in nearby cells. Our current understanding of SCCs and BCs focuses on their role as neural end organs; however, this focus does not preclude the possibility that these cells also play a role in signaling within the epithelium. This idea is generally consistent with the concept that SCCs and BCs act as sentinels of the airway, detecting potential dangers and triggering a protective response. For example, if ACh from taste cell-like epithelial chemosensors causes an increase in ciliary beating, then bacteria growing in the airway might be dislodged by the cilia and subsequently expelled when activated nociceptors trigger a cough. Similarly, if

SCCs and BCs are capable of triggering the release of bio-active chemicals from secretory cells, then those secretions might aid in fighting off a bacterial infection of the airway.

In addition to having an immediate protective effect in the respiratory epithelium,

ACh released from chronically stimulated SCCs and BCs may trigger morphological changes. When subjected to chronic inflammation, respiratory epithelium undergoes a series of morphological changes (Ren, Shah et al. 2013). These changes are generally described as a thickening of the epithelium or a change in the frequency of cell types in the epithelium, and these changes correspond to an increased incidence of cancer

(Manuyakorn, Howarth et al. 2013). ACh added to cultures of respiratory epithelium can induce several of these morphological changes (Metzen, Bittinger et al. 2003). This

88 suggests that sources of ACh in the epithelium, like SCCs and BCs, might be able to induce changes in the composition of the epithelium.

Air liquid interface (ALI) cultures could be used to test if SCCs and BCs are capable of causing changes in the composition of the epithelium. After growing ALI cultures, an activator of taste cell-like airway chemosensors, like denatonium, could be added and the culture and the epithelium assessed for changes in thickness or composition. If changes are observed, AChR antagonists, like mecamylamine or atropine, could be used to identify which ACh receptor subtype is mediating the response (Metzen,

Bittinger et al. 2003). Additionally, control experiments with cultures produced from gustducin or TRPM5 knockout mice could be used to show that any changes seen are due to the activation of the canonical taste transduction cascade. Variations of these experiments could be used to explore the effect of paracrine signaling by BCs and SCCs in the respiratory epithelium.

The Airway Chemofensor Complex

Chemofensor complex is a neologism used to describe the traditional chemosensory signaling found outside of the classical chemosensory systems, which may have a defensive effect (Green 2012). Essentially, the chemofensor complex can be viewed as part of the innate immune system, in the same way that pain signaling is generally considered an essential component of triggering the innate and adaptive immunity (DeLeo 2006). SCCs and BCs are detectors of bacterial infection and have a role in defending the airway from pathogens, either by expelling the pathogens through protective airway reflexes or by recruiting the immune system by inducing inflammation

(Tizzano, Gulbransen et al. 2010, Krasteva, Canning et al. 2012). However, the airway chemofensor complex consists of several cell types in addition to SCCs and BCs.

89

A chemofensor role has also recently been assigned to ciliated cells and pulmonary neuroendocrine cells in the respiratory epithelium in addition to SCCs and

BCs. In humans, ciliated cells express elements of the canonical taste transduction cascade (Shah, Ben-Shahar et al. 2009). When stimulated by bitter substances, ciliated cells increase the rate at which their cilia beat and produce reactive oxygen species that kill bacteria (Lee and Cohen 2013). We can easily imagine a situation where a respiratory pathogen, like Pseudomonas aeruginosa, produces chemicals that activate the canonical

“bitter” taste transduction cascades in both ciliated cells and taste cell-like airway chemosensors causing synergistic protective effects (Lee and Cohen 2013).

For example, AHLs from P. aeruginosa would cause the production of reactive oxygen species to kill some of the bacteria growing in the respiratory tract. At the same time, these AHLs are also causing an increase in ciliary beat frequency to dislodge bacteria growing in the respiratory tract. In parallel, the AHLs are also stimulating SCCs and BCs, which induce protective respiratory reflexes to expel dislodged bacteria and cause inflammation to recruit the immune system to fight off the resident infection utilizing classical immunological pathways. While this example focuses exclusively on the canonical bitter taste transduction pathway, recent experiments have also demonstrated olfactory signaling elements in some cells of the respiratory tract (Van

Lommel, Bolle et al. 1999).

Pulmonary neuroendocrine cells (PNECs) are a neuro-epithelial cell in found in the trachea and may utilize the elements of canonical olfactory signaling in a similar fashion to how SCCs and BCs utilize the canonical “bitter” taste transduction cascade.

Much like BCs, decades ago, PNECs were described morphologically but their exact function has remained a mystery (Van Lommel, Bolle et al. 1999). Also like BCs, PNECs are contacted by peptidergic nociceptors and should trigger inflammation on stimulation

90

(Linnoila 2006). Several reports have suggested that PNECs are hypoxia sensors, based on their expression of a hypoxia-sensitive potassium channel (Pan, Bear et al. 2002).

However, more recent reports have claimed that PNECs also express olfactory receptors

(Ben-Shahar 2013). Perhaps, PNECs are a parallel element of the airway chemofensor complex, utilizing olfactory receptors signaling to provide an additional mechanism for detecting respiratory pathogens and causing inflammation and irritation to defend the airway against invaders.

Concluding Thoughts

Since G.H. Parker coined the term “common chemical sense” to refer to the chemical sensitivity of epithelia, our understanding of chemical irritation has been constantly evolving. The identification of taste cell-like airway chemosensors and the elucidation of their function represent a paradigm shift in our understanding of the causes of chemical irritation. Additionally, linking SCCs and BCs to inflammation and the innate immune system have suggested their possible involvement in several pathological conditions and highlighted the possibility of translational significance. Future research on taste cell-like chemosensors will likely continue to have not only translation importance but also challenge our basic of mucociliary epithelium.

We define tissue types based on the constituent cells, and the classical definition of mucociliary epithelium consists of ciliated cells, secretory cells and basal cells.

However, cells similar to SCCs and BCs have also been identified in mucosal epithelia from the eustation tubes to the urinary tract (Krasteva, Hartmann et al. 2012, Kummer,

Flipski et al. 2012). If SCCs and BCs might be the same cell type, then perhaps there is

91 commonality between the taste cell-like chemosensors in all these epithelial layers. While further research would be required to establish unity between of these chemosensory cells, their reoccurring presence in these tissues highlights the fundamental role of epithelial layers as chemosensory end organs.

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REFERENCES

Abe, J., H. Hosokawa, M. Okazawa, M. Kandachi, Y. Sawada, K. Yamanaka, K. Matsumura and S. Kobayashi (2005). "TRPM8 protein localization in trigeminal ganglion and taste papillae." Brain Res Mol Brain Res 136(1-2): 91-98. Acs, G., T. Biro, P. Acs, S. Modarres and P. M. Blumberg (1997). "Differential activation and desensitization of sensory neurons by resiniferatoxin." J Neurosci 17(14): 5622-5628. Alarie, Y. (1966). "Irritating properties of airborne materials to the upper respiratory tract." Arch Environ Health 13(4): 433-449. Alarie, Y. (1973). "Sensory irritation by airborne chemicals." CRC Crit Rev Toxicol 2(3): 299-363. Alarie, Y. (1973). "Sensory irritation of the upper airways by airborne chemicals." Toxicol Appl Pharmacol 24(2): 279-297. Alimohammadi, H. and W. L. Silver (2000). "Evidence for nicotinic acetylcholine receptors on nasal trigeminal nerve endings of the rat." Chem Senses 25(1): 61- 66. Antony-Babu, S. and M. Goodfellow (2008). "Biosystematics of alkaliphilic streptomycetes isolated from seven locations across a beach and dune sand system." Antonie Van Leeuwenhoek 94(4): 581-591. Armstrong, D., R. M. Dry, C. A. Keele and J. W. Markham (1953). "Observations on chemical excitants of cutaneous pain in man." J Physiol 120(3): 326-351. Bandell, M., L. J. Macpherson and A. Patapoutian (2007). "From chills to chilis: mechanisms for thermosensation and chemesthesis via thermoTRPs." Curr Opin Neurobiol 17(4): 490-497. Bandell, M., G. M. Story, S. W. Hwang, V. Viswanath, S. R. Eid, M. J. Petrus, T. J. Earley and A. Patapoutian (2004). "Noxious Cold Ion Channel TRPA1 Is Activated by Pungent Compounds and Bradykinin." 41(6): 849-857. Barham, H. P., S. E. Cooper, C. B. Anderson, M. Tizzano, T. T. Kingdom, T. E. Finger, S. C. Kinnamon and V. R. Ramakrishnan (2013). "Solitary chemosensory cells and bitter taste receptor signaling in human sinonasal mucosa." Int Forum Allergy Rhinol 3(6):450-7. Barratt, W. (1898). "On the Anatomical Structure of the Vagus Nerve." J Anat Physiol 32(Pt 3): 422-427. Basbaum, A. I., D. M. Bautista, G. Scherrer and D. Julius (2009). "Cellular and molecular mechanisms of pain." Cell 139(2): 267-284. Basbaum, C. and B. Jany (1990). "Plasticity in the airway epithelium." Am J Physiol 259(2 Pt 1): L38-46. Bathla, G. and A. N. Hegde (2013). "The trigeminal nerve: an illustrated review of its imaging anatomy and pathology." Clin Radiol 68(2): 203-213.

93

Bautista, D. M., P. Movahed, A. Hinman, H. E. Axelsson, O. Sterner, E. D. Hogestatt, D. Julius, S. E. Jordt and P. M. Zygmunt (2005). "Pungent products from garlic activate the sensory ion channel TRPA1." Proc Natl Acad Sci U S A 102(34): 12248-12252. Bautista, D. M., Y. M. Sigal, A. D. Milstein, J. L. Garrison, J. A. Zorn, P. R. Tsuruda, R. A. Nicoll and D. Julius (2008). "Pungent agents from Szechuan peppers excite sensory neurons by inhibiting two-pore potassium channels." Nat Neurosci 11(7): 772-779. Beidler, L. M. and R. L. Smallman (1965). "Renewal of cells within taste buds." J Cell Biol 27(2): 263-272. Beitel, R. E. and R. Dubner (1976). "Response of unmyelinated (C) polymodal nociceptors to thermal stimuli applied to monkey's face." J Neurophysiol 39(6): 1160-1175. Ben-Shahar, Y. (2013). Human Pulmonary Neuroendocrine Cells are Olfactory Sentinels The Association for Chemoreception Sciences. 35th Annual Meeting. April 17- 20, 2013. Huntington Beach, CA. Benninger, M. S., B. J. Ferguson, J. A. Hadley, D. L. Hamilos, M. Jacobs, D. W. Kennedy, D. C. Lanza, B. F. Marple, J. D. Osguthorpe, J. A. Stankiewicz, J. Anon, J. Denneny, I. Emanuel and H. Levine (2003). "Adult chronic rhinosinusitis: definitions, diagnosis, epidemiology, and pathophysiology." Otolaryngol Head Neck Surg 129(3 Suppl): S1-32. Bezencon, C., J. le Coutre and S. Damak (2007). "Taste-signaling proteins are coexpressed in solitary intestinal epithelial cells." Chem Senses 32(1): 41-49. Blumenkrantz, N. and G. Asboe-Hansen (1975). "A selective stain for mast cells." Histochem J 7(3): 277-282. Boulais, N. and L. Misery (2008). "The epidermis: a sensory tissue." Eur J Dermatol 18(2): 119-127. Bousquet, J., W. Fokkens, P. Burney, S. R. Durham, C. Bachert, C. A. Akdis, G. W. Canonica, S. E. Dahlen, T. Zuberbier, T. Bieber, S. Bonini, P. J. Bousquet, J. L. Brozek, L. O. Cardell, R. Crameri, A. Custovic, P. Demoly, R. G. Van Wijk, M. Gjomarkaj, C. Holland, P. Howarth, M. Humbert, S. L. Johnston, F. Kauffmann, M. L. Kowalski, B. Lambrecht, S. Lehmann, B. Leynaert, K. Lodrup-Carlsen, J. Mullol, B. Niggemann, E. Nizankowska-Mogilnicka, N. Papadopoulos, G. Passalacqua, H. J. Schünemann, H. U. Simon, A. Todo-Bom, E. Toskala, R. Valenta, M. Wickman and J. P. Zock (2008). "Important research questions in allergy and related diseases: nonallergic rhinitis: a GA2LEN paper." Allergy 63(7): 842-853. Bousquet, J., N. Khaltaev, A. A. Cruz, J. Denburg, W. J. Fokkens, A. Togias, T. Zuberbier, C. E. Baena-Cagnani, G. W. Canonica, C. Van Weel, I. Agache, N. Aït- Khaled, C. Bachert, M. S. Blaiss, S. Bonini, L. P. Boulet, P. J. Bousquet, P. Camargos, K. H. Carlsen, Y. Chen, A. Custovic, R. Dahl, P. Demoly, H. Douagui, S. R. Durham, R. G. Van Wijk, O. Kalayci, M. A. Kaliner, Y. Y. Kim, M. L. Kowalski, P. Kuna, L. T. T. Le, C. Lemiere, J. Li, R. F. Lockey, S. Mavale-

94

Manuel, E. O. Meltzer, Y. Mohammad, J. Mullol, R. Naclerio, R. E. O’Hehir, K. Ohta, S. Ouedraogo, S. Palkonen, N. Papadopoulos, G. Passalacqua, R. Pawankar, T. A. Popov, K. F. Rabe, J. Rosado-Pinto, G. K. Scadding, F. E. R. Simons, E. Toskala, E. Valovirta, P. Van Cauwenberge, D. Y. Wang, M. Wickman, B. P. Yawn, A. Yorgancioglu, O. M. Yusuf, H. Zar, I. Annesi-Maesano, E. D. Bateman, A. B. Kheder, D. A. Boakye, J. Bouchard, P. Burney, W. W. Busse, M. Chan-Yeung, N. H. Chavannes, A. Chuchalin, W. K. Dolen, R. Emuzyte, L. Grouse, M. Humbert, C. Jackson, S. L. Johnston, P. K. Keith, J. P. Kemp, J. M. Klossek, D. Larenas-Linnemann, B. Lipworth, J. L. Malo, G. D. Marshall, C. Naspitz, K. Nekam, B. Niggemann, E. Nizankowska-Mogilnicka, Y. Okamoto, M. P. Orru, P. Potter, D. Price, S. W. Stoloff, O. Vandenplas, G. Viegi and D. Williams (2008). "Allergic Rhinitis and its Impact on Asthma (ARIA) 2008*." Allergy 63: 8-160. Bragulla, H. H. and D. G. Homberger (2009). "Structure and functions of keratin proteins in simple, stratified, keratinized and cornified epithelia." J Anat 214(4): 516-559. Brechbuhl, H. M., M. Ghosh, M. K. Smith, R. W. Smith, B. Li, D. A. Hicks, B. B. Cole, P. R. Reynolds and S. D. Reynolds (2011). "beta-catenin dosage is a critical determinant of tracheal basal cell fate determination." Am J Pathol 179(1): 367- 379. Breuel, K. F. and W. K. De Ponti (2006). "Measurement of mast cell cytokine release by multiplex assay." Methods Mol Biol 315: 217-230. Brewer, C., R. G. Wunderink, C. B. Jones and K. V. Leeper (1996). "Ventilator- associated pneumonia due to Pseudomonas aeruginosa." Chest 109(4): 1019- 1029. Brody, A. R. (2005). "The brush cell." Am J Respir Crit Care Med 172(10): 1349. Bryant, B. P. and W. Silver (2000). Chemesthesis: The common chemical sense. Neurobiology of Taste and Smell. T. E. Finger, W. Silver and D. Restrepo, Wiley- Liss, Inc: 73-100. Bryant, B. P. and W. L. Silver (2000). Chemesthesis: The common chemical sense. Neurobiology of Taste and Smell. F. TE, S. W and R. D. New York, Wiley-Liss, Inc: 73–100. Canning, B. J. and D. Spina (2009). "Sensory nerves and airway irritability." Handb Exp Pharmacol(194): 139-183. Carr, M. J. and B. J. Undem (2003). "Bronchopulmonary afferent nerves." Respirology 8(3): 291-301. Carreno, C., A. Domenech, N. Prats, M. Miralpeix and I. Ramis (2012). "Characterization of a model of tracheal plasma extravasation in passively sensitized rats using anti-allergic and anti-inflammatory drugs by oral and intratracheal route." Pulm Pharmacol Ther 25(1): 87-93. Caspani, O. and P. A. Heppenstall (2009). "TRPA1 and cold transduction: an unresolved issue?" J Gen Physiol 133(3): 245-249.

95

Caterina, M. J., M. A. Schumacher, M. Tominaga, T. A. Rosen, J. D. Levine and D. Julius (1997). "The capsaicin receptor: a heat-activated ion channel in the pain pathway." Nature 389(6653): 816-824. Cauna, N., K. H. Hinderer and R. T. Wentges (1969). "Sensory receptor organs of the human nasal respiratory mucosa." Am J Anat 124(2): 187-209. Cavanaugh, D. J., A. T. Chesler, J. M. Braz, N. M. Shah, D. Julius and A. I. Basbaum (2011). "Restriction of transient receptor potential vanilloid-1 to the peptidergic subset of primary afferent neurons follows its developmental downregulation in nonpeptidergic neurons." J Neurosci 31(28): 10119-10127. Cho, Y. K., A. I. Farbman and D. V. Smith (1998). "The timing of alpha-gustducin expression during cell renewal in rat vallate taste buds." Chem Senses 23(6): 735- 742. Chung, M. K., H. Lee, A. Mizuno, M. Suzuki and M. J. Caterina (2004). "TRPV3 and TRPV4 mediate warmth-evoked currents in primary mouse keratinocytes." J Biol Chem 279(20): 21569-21575. Clapp, T. R., K. F. Medler, S. Damak, R. F. Margolskee and S. C. Kinnamon (2006). "Mouse taste cells with G protein-coupled taste receptors lack voltage-gated calcium channels and SNAP-25." BMC Biol 4: 7. Cole, B. B., R. W. Smith, K. M. Jenkins, B. B. Graham, P. R. Reynolds and S. D. Reynolds (2010). "Tracheal Basal cells: a facultative progenitor cell pool." Am J Pathol 177(1): 362-376. Cosens, D. J. and A. Manning (1969). "Abnormal electroretinogram from a Drosophila mutant." Nature 224(5216): 285-287. Daniely, Y., G. Liao, D. Dixon, R. I. Linnoila, A. Lori, S. H. Randell, M. Oren and A. M. Jetten (2004). "Critical role of p63 in the development of a normal esophageal and tracheobronchial epithelium." Am J Physiol Cell Physiol 287(1): C171-181. Dawicki, W. and J. S. Marshall (2007). "New and emerging roles for mast cells in host defence." Curr Opin Immunol 19(1): 31-38. del Camino, D., S. Murphy, M. Heiry, L. B. Barrett, T. J. Earley, C. A. Cook, M. J. Petrus, M. Zhao, M. D'Amours, N. Deering, G. J. Brenner, M. Costigan, N. J. Hayward, J. A. Chong, C. M. Fanger, C. J. Woolf, A. Patapoutian and M. M. Moran (2010). "TRPA1 contributes to cold hypersensitivity." J Neurosci 30(45): 15165-15174. DeLeo, J. A. (2006). "Basic science of pain." J Bone Joint Surg Am 88 Suppl 2: 58-62. Delgiorno, K. E., J. C. Hall, K. K. Takeuchi, F. C. Pan, C. J. Halbrook, M. K. Washington, K. P. Olive, J. Spence, B. Sipos, C. V. Wright, J. M. Wells and H. C. Crawford (2013). "Identification and Manipulation of Biliary Metaplasia in Pancreatic Tumors." Gastroenterology. [Epub ahead of print]

96

Deshpande, D. A., W. C. Wang, E. L. McIlmoyle, K. S. Robinett, R. M. Schillinger, S. S. An, J. S. Sham and S. B. Liggett (2010). "Bitter taste receptors on airway smooth muscle bronchodilate by localized calcium signaling and reverse obstruction." Nat Med 16(11): 1299-1304. Dhaka, A., T. J. Earley, J. Watson and A. Patapoutian (2008). "Visualizing cold spots: TRPM8-expressing sensory neurons and their projections." J Neurosci 28(3): 566-575. Eberhard, A., A. L. Burlingame, C. Eberhard, G. L. Kenyon, K. H. Nealson and N. J. Oppenheimer (1981). "Structural Identification of Autoinducer of Photobacterium-Fischeri Luciferase." Biochemistry 20(9): 2444-2449. Farbman, A. I. (1980). "Renewal of taste bud cells in rat circumvallate papillae." Cell Tissue Kinet 13(4): 349-357. Finger, T. E. (1997). " of taste and solitary chemoreceptor cell systems." Brain Behav Evol 50(4): 234-243. Finger, T. E., B. Bottger, A. Hansen, K. T. Anderson, H. Alimohammadi and W. L. Silver (2003). "Solitary chemoreceptor cells in the nasal cavity serve as sentinels of respiration." Proc Natl Acad Sci U S A 100(15): 8981-8986. Finger, T. E., V. Danilova, J. Barrows, D. L. Bartel, A. J. Vigers, L. Stone, G. Hellekant and S. C. Kinnamon (2005). "ATP signaling is crucial for communication from taste buds to gustatory nerves." Science 310(5753): 1495-1499. Finger, T. E., V. L. St Jeor, J. C. Kinnamon and W. L. Silver (1990). "Ultrastructure of substance P- and CGRP-immunoreactive nerve fibers in the nasal epithelium of rodents." J Comp Neurol 294(2): 293-305. Flückiger, F. and D. Hanbury (1874). History of quinine. Pharmacographia: A history of the principal drugs of vegetable origin, met with in Great Britain and British India. London, England, Macmillan and Co. Folkesson, A., L. Jelsbak, L. Yang, H. K. Johansen, O. Ciofu, N. Hoiby and S. Molin (2012). "Adaptation of Pseudomonas aeruginosa to the cystic fibrosis airway: an evolutionary perspective." Nat Rev Microbiol 10(12): 841-851. Frasnelli, J., S. Heilmann and T. Hummel (2004). "Responsiveness of human nasal mucosa to trigeminal stimuli depends on the site of stimulation." Neurosci Lett 362(1): 65-69. Fuchs, E. (1995). "Keratins and the skin." Annu Rev Cell Dev Biol 11: 123-153. Fuqua, C. and E. P. Greenberg (2002). "Listening in on bacteria: Acyl-homoserine lactone signalling." Nature Reviews Molecular Cell Biology 3(9): 685-695. Fuqua, W. C., S. C. Winans and E. P. Greenberg (1994). "Quorum Sensing in Bacteria - the Luxr-Luxi Family of Cell Density-Responsive Transcriptional Regulators." Journal of Bacteriology 176(2): 269-275.

97

Galli, S. J., J. Kalesnikoff, M. A. Grimbaldeston, A. M. Piliponsky, C. M. Williams and M. Tsai (2005). "Mast cells as "tunable" effector and immunoregulatory cells: recent advances." Annu Rev Immunol 23: 749-786. Galli, S. J., S. Nakae and M. Tsai (2005). "Mast cells in the development of adaptive immune responses." Nat Immunol 6(2): 135-142. Gelardi, M., A. Maselli del Giudice, M. L. Fiorella, R. Fiorella, C. Russo, P. Soleti, M. Di Gioacchino and G. Ciprandi (2008). "Non-allergic rhinitis with eosinophils and mast cells constitutes a new severe nasal disorder." Int J Immunopathol Pharmacol 21(2): 325-331. Gerhold, K. A. and D. M. Bautista (2009). "Molecular and cellular mechanisms of trigeminal chemosensation." Ann N Y Acad Sci 1170: 184-189. Ghosh, M., H. M. Brechbuhl, R. W. Smith, B. Li, D. A. Hicks, T. Titchner, C. M. Runkle and S. D. Reynolds (2011). "Context-dependent differentiation of multipotential keratin 14-expressing tracheal basal cells." Am J Respir Cell Mol Biol 45(2): 403-410. Ghosh, M., K. M. Helm, R. W. Smith, M. S. Giordanengo, B. Li, H. Shen and S. D. Reynolds (2011). "A single cell functions as a tissue-specific stem cell and the in vitro niche-forming cell." Am J Respir Cell Mol Biol 45(3): 459-469. Gilfillan, A. M. and C. Tkaczyk (2006). "Integrated signalling pathways for mast-cell activation." Nat Rev Immunol 6(3): 218-230. Gliklich, R. E. and R. Metson (1995). "The health impact of chronic sinusitis in patients seeking otolaryngologic care." Otolaryngol Head Neck Surg 113(1): 104-109. Green, B. G. (1985). "Menthol modulates oral sensations of warmth and cold." Physiol Behav 35(3): 427-434. Green, B. G. (2012). "Chemesthesis and the chemical senses as components of a "chemofensor complex"." Chem Senses 37(3): 201-206. Green, B. G. and L. J. Flammer (1989). "Localization of chemical stimulation: capsaicin on hairy skin." Somatosens Mot Res 6(5-6): 553-566. Green, B. G., J. R. Mason and M. R. Kare (1990). Chemical senses, vol. 2: Irritation. New York, Marcel Dekker, Inc. Greiff, L., I. Erjefalt, P. Wollmer, U. Pipkorn and C. G. Persson (1991). "Effects of histamine, ethanol, and a detergent on exudation and absorption across guinea pig airway mucosa in vivo." Thorax 46(10): 700-705. Gri, G., B. Frossi, F. D'Inca, L. Danelli, E. Betto, F. Mion, R. Sibilano and C. Pucillo (2012). "Mast cell: an emerging partner in immune interaction." Front Immunol 3: 120. Grybko, M. J., E. T. Hahm, W. Perrine, J. A. Parnes, W. S. Chick, G. Sharma, T. E. Finger and S. Vijayaraghavan (2011). "A transgenic mouse model reveals fast nicotinic transmission in hippocampal pyramidal neurons." Eur J Neurosci 33(10): 1786-1798.

98

Guimaraes, M. Z. P. and S. E. Jordt (2007). TRPA1 : A Sensory Channel of Many Talents. TRP Ion Channel Function in Sensory Transduction and Cellular Signaling Cascades. W. B. Liedtke and S. Heller. Boca Raton (FL). Gulbransen, B. (2007). Nasal solitary chemoreceptor cells: cell turnover, nerve dependence and detection capabilities. PhD Dissertation, University of Colorado- Denver. Gulbransen, B., W. Silver and T. E. Finger (2008). "Solitary chemoreceptor cell survival is independent of intact trigeminal innervation." J Comp Neurol 508(1): 62-71. Gulbransen, B. D., T. R. Clapp, T. E. Finger and S. C. Kinnamon (2008). "Nasal solitary chemoreceptor cell responses to bitter and trigeminal stimulants in vitro." J Neurophysiol 99(6): 2929-2937. Gulbransen, B. D. and T. E. Finger (2005). "Solitary chemoreceptor cell proliferation in adult nasal epithelium." J Neurocytol 34(1-2): 117-122. Hamamichi, R., M. Asano-Miyoshi and Y. Emori (2006). "Taste bud contains both short- lived and long-lived cell populations." Neuroscience 141(4): 2129-2138. Hansen, A. (2007). "Olfactory and solitary chemosensory cells: two different chemosensory systems in the nasal cavity of the American alligator, Alligator mississippiensis." BMC Neurosci 8: 64. Hansen, A. and T. E. Finger (2008). "Is TrpM5 a reliable marker for chemosensory cells? Multiple types of microvillous cells in the main olfactory epithelium of mice." BMC Neurosci 9: 115. Hargreave, F. E., J. Dolovich, P. M. O'Byrne, E. H. Ramsdale and E. E. Daniel (1986). "The origin of airway hyperresponsiveness." J Allergy Clin Immunol 78(5 Pt 1): 825-832. Harkema, J. R., S. A. Carey and J. G. Wagner (2006). "The nose revisited: a brief review of the comparative structure, function, and toxicologic pathology of the nasal epithelium." Toxicol Pathol 34(3): 252-269. Hensel, H. and Y. Zotterman (1951). "The effect of menthol on the ." Acta Physiol Scand 24(1): 27-34. Hinman, A., H. H. Chuang, D. M. Bautista and D. Julius (2006). "TRP channel activation by reversible covalent modification." Proc Natl Acad Sci U S A 103(51): 19564- 19568. Hong, K. U., S. D. Reynolds, S. Watkins, E. Fuchs and B. R. Stripp (2004). "Basal cells are a multipotent progenitor capable of renewing the bronchial epithelium." Am J Pathol 164(2): 577-588. Hong, K. U., S. D. Reynolds, S. Watkins, E. Fuchs and B. R. Stripp (2004). "In vivo differentiation potential of tracheal basal cells: evidence for multipotent and unipotent subpopulations." Am J Physiol Lung Cell Mol Physiol 286(4): L643- 649.

99

Ikoma, A., M. Steinhoff, S. Stander, G. Yosipovitch and M. Schmelz (2006). "The neurobiology of itch." Nat Rev Neurosci 7(7): 535-547. Ishizaka, T., C. M. Sian and K. Ishizaka (1972). "Complement fixation by aggregated IgE through alternate pathway." J Immunol 108(3): 848-851. Ivanusic, J. J., M. M. Kwok and E. A. Jennings (2011). "Peripheral targets of 5-HT(1D) receptor immunoreactive trigeminal ganglion neurons." Headache 51(5): 744- 751. Jones, N. S. (2005). "Sinogenic facial pain: diagnosis and management." Otolaryngol Clin North Am 38(6): 1311-1325, x-xi. Julius, D. and A. I. Basbaum (2001). "Molecular mechanisms of nociception." Nature 413(6852): 203-210. Kane, L. E., C. S. Barrow and Y. Alarie (1979). "A short-term test to predict acceptable levels of exposure to airborne sensory irritants." Am Ind Hyg Assoc J 40(3): 207- 229. Kaske, S., G. Krasteva, P. Konig, W. Kummer, T. Hofmann, T. Gudermann and V. Chubanov (2007). "TRPM5, a taste-signaling transient receptor potential ion- channel, is a ubiquitous signaling component in chemosensory cells." BMC Neurosci 8: 49. Keele, C. A. (1962). "The common chemical sense and its receptors." Arch Int Pharmacodyn Ther 139: 547-557. Klein-Schwartz, W. (1991). "Denatonium benzoate: review of efficacy and safety." Vet Hum Toxicol 33(6): 545-547. Knipping, S., H. J. Holzhausen, A. Riederer and T. Schrom (2009). "Allergic and idiopathic rhinitis: an ultrastructural study." Eur Arch Otorhinolaryngol 266(8): 1249-1256. Kobayashi, K., T. Fukuoka, K. Obata, H. Yamanaka, Y. Dai, A. Tokunaga and K. Noguchi (2005). "Distinct expression of TRPM8, TRPA1, and TRPV1 mRNAs in rat primary afferent neurons with adelta/c-fibers and colocalization with trk receptors." J Comp Neurol 493(4): 596-606. Kobayashi, K., T. Fukuoka, K. Obata, H. Yamanaka, Y. Dai, A. Tokunaga and K. Noguchi (2005). "Distinct expression of TRPM8, TRPA1, and TRPV1 mRNAs in rat primary afferent neurons with aδ/c-fibers and colocalization with trk receptors." The Journal of Comparative 493(4): 596-606. Kotrschal, K., M. Whitear and T. E. Finger (1993). "Spinal and facial innervation of the skin in the gadid fish Ciliata mustela (Teleostei)." J Comp Neurol 331(3): 407- 417. Krasteva, G., B. J. Canning, P. Hartmann, T. Z. Veres, T. Papadakis, C. Muhlfeld, K. Schliecker, Y. N. Tallini, A. Braun, H. Hackstein, N. Baal, E. Weihe, B. Schutz, M. Kotlikoff, I. Ibanez-Tallon and W. Kummer (2011). "Cholinergic chemosensory cells in the trachea regulate breathing." Proc Natl Acad Sci U S A 108(23): 9478-9483.

100

Krasteva, G., B. J. Canning, T. Papadakis and W. Kummer (2012). "Cholinergic brush cells in the trachea mediate respiratory responses to quorum sensing molecules." Life Sci 91(21-22): 992-996. Krasteva, G., P. Hartmann, T. Papadakis, M. Bodenbenner, L. Wessels, E. Weihe, B. Schutz, A. C. Langheinrich, V. Chubanov, T. Gudermann, I. Ibanez-Tallon and W. Kummer (2012). "Cholinergic chemosensory cells in the auditory tube." Histochem Cell Biol 137(4):483-97 Kummer, W., K. Flipski, T. Papadakis, M. Wolff, I. Ibanez-Tallon, T. Bschleiper and G. Krasteva (2012). Cholinergic chemosensory Brush cells in the Murine Urethra. Poster presented at: Society for Neuroscience; 2012 Oct 13-17; New Orleans, LA Kummer, W. and K. S. Lips (2006). "Non-neuronal acetylcholine release and its contribution to COPD pathology." Drug Discovery Today: Disease Mechanisms 3(1): 47-52. Kummer, W., K. S. Lips and U. Pfeil (2008). "The epithelial cholinergic system of the airways." Histochem Cell Biol 130(2): 219-234. Lagunoff, D., T. W. Martin and G. Read (1983). "Agents that release histamine from mast cells." Annu Rev Pharmacol Toxicol 23: 331-351. Langley, J. N. (1900). "On axon-reflexes in the pre-ganglionic fibres of the sympathetic system." J Physiol 25(5): 364-398. Lawrence, T., D. A. Willoughby and D. W. Gilroy (2002). "Anti-inflammatory lipid mediators and insights into the resolution of inflammation." Nat Rev Immunol 2(10): 787-795. Lee, R. J. and N. A. Cohen (2013). "The emerging role of the bitter taste receptor T2R38 in upper respiratory infection and chronic rhinosinusitis." Am J Rhinol Allergy 27(4): 283-286. Lee, R. J., G. Xiong, J. M. Kofonow, B. Chen, A. Lysenko, P. Jiang, V. Abraham, L. Doghramji, N. D. Adappa, J. N. Palmer, D. W. Kennedy, G. K. Beauchamp, P. T. Doulias, H. Ischiropoulos, J. L. Kreindler, D. R. Reed and N. A. Cohen (2012). "T2R38 taste receptor polymorphisms underlie susceptibility to upper respiratory infection." J Clin Invest 122(11): 4145-4159. Liedtke, W., Y. Choe, M. A. Marti-Renom, A. M. Bell, C. S. Denis, A. Sali, A. J. Hudspeth, J. M. Friedman and S. Heller (2000). "Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor." Cell 103(3): 525-535. Lin, W., T. Ogura, R. F. Margolskee, T. E. Finger and D. Restrepo (2008). "TRPM5- expressing solitary chemosensory cells respond to odorous irritants." J Neurophysiol 99(3): 1451-1460. Linnoila, R. I. (2006). "Functional facets of the pulmonary neuroendocrine system." Lab Invest 86(5): 425-444.

101

Liu, L. and S. A. Simon (1996). "Similarities and differences in the currents activated by capsaicin, piperine, and zingerone in rat trigeminal ganglion cells." Journal of 76(3): 1858-1869. Lopez-Schier, H. (2004). "Regeneration: did you hear the news?" Curr Biol 14(3): R127- 128. Luciano, L., E. Reale and H. Ruska (1969). "[Brush cells in the alveolar epithelium of the rat lung]." Z Zellforsch Mikrosk Anat 95(2): 198-201. Lumpkin, E. A. and M. J. Caterina (2007). "Mechanisms of sensory transduction in the skin." Nature 445(7130): 858-865. Lundberg, J. M., E. Brodin, X. Hua and A. Saria (1984). "Vascular permeability changes and smooth muscle contraction in relation to capsaicin-sensitive substance P afferents in the guinea-pig." Acta Physiol Scand 120(2): 217-227. Lundblad, L. and J. M. Lundberg (1984). "Capsaicin sensitive sensory neurons mediate the response to nasal irritation induced by the vapour phase of cigarette smoke." Toxicology 33(1): 1-7. Lundblad, L., J. M. Lundberg, E. Brodin and A. Anggard (1983). "Origin and distribution of capsaicin-sensitive substance P-immunoreactive nerves in the nasal mucosa." Acta Otolaryngol 96(5-6): 485-493. MacGlashan, D. W., Jr. (2012). "IgE-dependent signaling as a therapeutic target for allergies." Trends Pharmacol Sci 33(9): 502-509. Mack, J. A., S. Anand and E. V. Maytin (2005). "Proliferation and cornification during development of the mammalian epidermis." Birth Defects Res C Embryo Today 75(4): 314-329. Mackay-Sim, A. and P. Kittel (1991). "Cell dynamics in the adult mouse olfactory epithelium: a quantitative autoradiographic study." J Neurosci 11(4): 979-984. Macpherson, L. J., S. W. Hwang, T. Miyamoto, A. E. Dubin, A. Patapoutian and G. M. Story (2006). "More than cool: Promiscuous relationships of menthol and other sensory compounds." Molecular and 32(4): 335-343. Macpherson, L. J., B. Xiao, K. Y. Kwan, M. J. Petrus, A. E. Dubin, S. Hwang, B. Cravatt, D. P. Corey and A. Patapoutian (2007). "An ion channel essential for sensing chemical damage." J Neurosci 27(42): 11412-11415. Mandadi, S., T. Sokabe, K. Shibasaki, K. Katanosaka, A. Mizuno, A. Moqrich, A. Patapoutian, T. Fukumi-Tominaga, K. Mizumura and M. Tominaga (2009). "TRPV3 in keratinocytes transmits temperature information to sensory neurons via ATP." Pflugers Arch 458(6): 1093-1102. Manuyakorn, W., P. H. Howarth and S. T. Holgate (2013). "Airway remodelling in asthma and novel therapy." Asian Pac J Allergy Immunol 31(1): 3-10.

102

Maouche, K., M. Polette, T. Jolly, K. Medjber, I. Cloez-Tayarani, J. P. Changeux, H. Burlet, C. Terryn, C. Coraux, J. M. Zahm, P. Birembaut and J. M. Tournier (2009). "{alpha}7 nicotinic acetylcholine receptor regulates airway epithelium differentiation by controlling basal cell proliferation." Am J Pathol 175(5): 1868- 1882. McCoy, D. D., W. M. Knowlton and D. D. McKemy (2011). "Scraping through the ice: uncovering the role of TRPM8 in cold transduction." Am J Physiol Regul Integr Comp Physiol 300(6): R1278-1287. McDonald, D. M., G. Thurston and P. Baluk (1999). "Endothelial gaps as sites for plasma leakage in inflammation." Microcirculation 6(1): 7-22. McKemy, D. D., W. M. Neuhausser and D. Julius (2002). "Identification of a cold receptor reveals a general role for TRP channels in thermosensation." Nature 416(6876): 52-58. Merigo, F., D. Benati, M. Cristofoletti, F. Amaru, F. Osculati and A. Sbarbati (2012). "Glucose transporter/T1R3-expressing cells in rat tracheal epithelium." J Anat 221(2): 138-150. Merigo, F., D. Benati, M. Tizzano, F. Osculati and A. Sbarbati (2005). "alpha-Gustducin immunoreactivity in the airways." Cell Tissue Res 319(2): 211-219. Metcalfe, D. D., D. Baram and Y. A. Mekori (1997). "Mast cells." Physiol Rev 77(4): 1033-1079. Metzen, J., F. Bittinger, C. J. Kirkpatrick, H. Kilbinger and I. Wessler (2003). "Proliferative effect of acetylcholine on rat trachea epithelial cells is mediated by nicotinic receptors and muscarinic receptors of the M1-subtype." Life Sci 72(18- 19): 2075-2080. Milsom, W. K. and M. L. Burleson (2007). "Peripheral arterial chemoreceptors and the evolution of the carotid body." Respir Physiol Neurobiol 157(1): 4-11. Moulton, D. G. (1974). "Dynamics of cell populations in the olfactory epithelium." Ann N Y Acad Sci 237(0): 52-61. Muroi, Y. and B. J. Undem (2011). "Targeting peripheral afferent nerve terminals for cough and dyspnea." Curr Opin Pharmacol 11(3): 254-264. Nakae, S., H. Suto, M. Kakurai, J. D. Sedgwick, M. Tsai and S. J. Galli (2005). "Mast cells enhance T cell activation: Importance of mast cell-derived TNF." Proc Natl Acad Sci U S A 102(18): 6467-6472. Nestle, F. O., P. Di Meglio, J. Z. Qin and B. J. Nickoloff (2009). "Skin immune sentinels in health and disease." Nat Rev Immunol 9(10): 679-691. Nguyen, H. M., M. E. Reyland and L. A. Barlow (2012). "Mechanisms of taste bud cell loss after head and neck irradiation." J Neurosci 32(10): 3474-3484. Nielsen, G. D. (1991). "Mechanisms of activation of the sensory irritant receptor by airborne chemicals." Crit Rev Toxicol 21(3): 183-208.

103

O'Connor, T. M., J. O'Connell, D. I. O'Brien, T. Goode, C. P. Bredin and F. Shanahan (2004). "The role of substance P in inflammatory disease." J Cell Physiol 201(2): 167-180. Ohmoto, M., T. Yamaguchi, J. Yamashita, A. A. Bachmanov, J. Hirota and I. Matsumoto (2013). "Pou2f3/Skn-1a Is Necessary for the Generation or Differentiation of Solitary Chemosensory Cells in the Anterior Nasal Cavity." Biosci Biotechnol Biochem 77(10):2154-6 Okumura, Y., M. Narukawa, Y. Iwasaki, A. Ishikawa, H. Matsuda, M. Yoshikawa and T. Watanabe (2010). "Activation of TRPV1 and TRPA1 by black pepper components." Biosci Biotechnol Biochem 74(5): 1068-1072. Olson, T. H., M. S. Riedl, L. Vulchanova, X. R. Ortiz-Gonzalez and R. Elde (1998). "An acid sensing ion channel (ASIC) localizes to small primary afferent neurons in rats." Neuroreport 9(6): 1109-1113. Pan, J., C. Bear, S. Farragher, E. Cutz and H. Yeger (2002). "Cystic fibrosis transmembrane conductance regulator modulates neurosecretory function in pulmonary neuroendocrine cell-related tumor cell line models." Am J Respir Cell Mol Biol 27(5): 553-560. Parker, G. H. (1912). "The relations of smell taste and the common chemical sense in vertebrates." J Acad Nat Sci Philadelphia 14:221-234. Parker, G. H. (1922). Smell Taste and Allied Senses in the Vertebrates. Philadelphia, J.B. Lippincott Co. Peier, A. M., A. Moqrich, A. C. Hergarden, A. J. Reeve, D. A. Andersson, G. M. Story, T. J. Earley, I. Dragoni, P. McIntyre, S. Bevan and A. Patapoutian (2002). "A TRP channel that senses cold stimuli and menthol." Cell 108(5): 705-715. Peier, A. M., A. J. Reeve, D. A. Andersson, A. Moqrich, T. J. Earley, A. C. Hergarden, G. M. Story, S. Colley, J. B. Hogenesch, P. McIntyre, S. Bevan and A. Patapoutian (2002). "A heat-sensitive TRP channel expressed in keratinocytes." Science 296(5575): 2046-2049. Peters, R. C., K. Kotrschal and W. D. Krautgartner (1991). "Solitary Chemoreceptor Cells of Ciliata-Mustela (Gadidae, Teleostei) Are Tuned to Mucoid Stimuli." Chemical Senses 16(1): 31-42. Qureshi, R. and B. A. Jakschik (1988). "The role of mast cells in thioglycollate-induced inflammation." J Immunol 141(6): 2090-2096. Rather, L. J. (1971). "Disturbance of function (functio laesa): the legendary fifth cardinal sign of inflammation, added by Galen to the four cardinal signs of Celsus." Bull N Y Acad Med 47(3): 303-322. Rawlins, E. L. and B. L. Hogan (2008). "Ciliated epithelial cell lifespan in the mouse trachea and lung." Am J Physiol Lung Cell Mol Physiol 295(1): L231-234. Rawlins, E. L., T. Okubo, J. Que, Y. Xue, C. Clark, X. Luo and B. L. Hogan (2008). "Epithelial stem/progenitor cells in lung postnatal growth, maintenance, and repair." Cold Spring Harb Symp Quant Biol 73: 291-295.

104

Rawlins, E. L., T. Okubo, Y. Xue, D. M. Brass, R. L. Auten, H. Hasegawa, F. Wang and B. L. Hogan (2009). "The role of Scgb1a1+ Clara cells in the long-term maintenance and repair of lung airway, but not alveolar, epithelium." Cell Stem Cell 4(6): 525-534. Reid, L., B. Meyrick, V. B. Antony, L. Y. Chang, J. D. Crapo and H. Y. Reynolds (2005). "The mysterious pulmonary brush cell: a cell in search of a function." Am J Respir Crit Care Med 172(1): 136-139. Ren, X., T. A. Shah, V. Ustiyan, Y. Zhang, J. Shinn, G. Chen, J. A. Whitsett, T. V. Kalin and V. V. Kalinichenko (2013). "FOXM1 promotes allergen-induced goblet cell metaplasia and pulmonary inflammation." Mol Cell Biol 33(2): 371-386. Reynolds, S. D., A. Giangreco, J. H. Power and B. R. Stripp (2000). "Neuroepithelial bodies of pulmonary airways serve as a reservoir of progenitor cells capable of epithelial regeneration." Am J Pathol 156(1): 269-278. Reynolds, S. D. and A. M. Malkinson (2010). "Clara cell: Progenitor for the bronchiolar epithelium." The International Journal of Biochemistry & Cell Biology 42(1): 1- 4. Rhodin, J. and T. Dalhamn (1956). "Electron microscopy of the tracheal ciliated mucosa in rat." Z Zellforsch Mikrosk Anat 44(4): 345-412. Richards, P., C. J. Saunders and W. Silver (2010). Functional of the Upper Airway in Experimental Animals. Toxicology of the Nose and Upper Airways: 45-64. Richards PM, Johnson EC, Silver WL: Four Irritating Odorants Target the Trigeminal Chemoreceptor TRPA1. Chemosensory Perception. 2010; 3(3–4): 190–199. Rock, J. R., S. H. Randell and B. L. M. Hogan (2010). "Airway basal stem cells: a perspective on their roles in epithelial homeostasis and remodeling." Disease Models & Mechanisms 3(9-10): 545-556. Rodger, I. W., C. Tousignant, D. Young, C. Savoie and C. C. Chan (1995). "Neurokinin receptors subserving plasma extravasation in guinea pig airways." Can J Physiol Pharmacol 73(7): 927-931. Rothschild, A. M. (1970). "Mechanisms of histamine release by compound 48-80." Br J Pharmacol 38(1): 253-262. Ruffoli, R., F. S. Giorgi, C. Pizzanelli, L. Murri, A. Paparelli and F. Fornai (2011). "The chemical neuroanatomy of vagus nerve stimulation." J Chem Neuroanat 42(4): 288-296. Saunders, C. J., S. D. Reynolds and T. E. Finger (2013). "Chemosensory Brush Cells of the Trachea: A Stable Population in a Dynamic Epithelium." Am J Respir Cell Mol Biol 49(2):190-6.

105

Saunders, C. J., L. Y. Winston, T. D. Patel, J. A. Muday and W. L. Silver (2013). "Dissecting the role of TRPV1 in detecting multiple trigeminal irritants in three behavioral assays for sensory irritation [v1; ref status: indexed, http://f1000r.es/p8]." F1000Research 2: 74. Sbarbati, A., F. Merigo, D. Benati, M. Tizzano, P. Bernardi, C. Crescimanno and F. Osculati (2004). "Identification and characterization of a specific sensory epithelium in the rat larynx." J Comp Neurol 475(2): 188-201. Sbarbati, A. and F. Osculati (2006). "Allelochemical communication in vertebrates: Kairomones, allomones and synomones." Cells Tissues Organs 183(4): 206-219. Schafer, K., H. A. Braun and C. Isenberg (1986). "Effect of menthol on cold receptor activity. Analysis of receptor processes." J Gen Physiol 88(6): 757-776. Scholzen, T. and J. Gerdes (2000). "The Ki67 protein: from the known and the unknown." J Cell Physiol 182(3): 311-322. Seo, M. W., D. S. Yang, S. J. Kays, G. P. Lee and K. W. Park (2009). "Sesquiterpene Lactones and Bitterness in Korean Leaf Lettuce Cultivars." Hortscience 44(2): 246-249. Settipane, R. A. and M. A. Kaliner (2013). "Chapter 14: Nonallergic rhinitis." Am J Rhinol Allergy 27 Suppl 1: 48-51. Shah, A. S., Y. Ben-Shahar, T. O. Moninger, J. N. Kline and M. J. Welsh (2009). "Motile cilia of human airway epithelia are chemosensory." Science 325(5944): 1131- 1134. Shusterman, D. and M. A. Murphy (2007). "Nasal hyperreactivity in allergic and non- allergic rhinitis: a potential risk factor for non-specific building-related illness." Indoor Air 17(4): 328-333. Silver, W., P. Roe and C. J. Saunders (2010). Functional Neuroanatomy of the Upper Airway in Experimental Animals. Toxicology of the Nose and Upper Airways. J. B. Morris and D. Shusterman. New York, Informa Healthcare: 45 - 64. Silver, W. L. and T. E. Finger (2009). "The Anatomical and Electrophysiological Basis of Peripheral Nasal Trigeminal Chemoreception." Annals of the New York Academy of Sciences 1170(1): 202-205. Smith, G. D., M. J. Gunthorpe, R. E. Kelsell, P. D. Hayes, P. Reilly, P. Facer, J. E. Wright, J. C. Jerman, J. P. Walhin, L. Ooi, J. Egerton, K. J. Charles, D. Smart, A. D. Randall, P. Anand and J. B. Davis (2002). "TRPV3 is a temperature-sensitive vanilloid receptor-like protein." Nature 418(6894): 186-190. Smith, M. K., P. J. Koch and S. D. Reynolds (2012). "Direct and Indirect Roles for B- Catenin in Facultative Basal Progenitor Cell Differentiation." Am J Physiol Lung Cell Mol Physiol 15;302(6):L580-94. Song, M. Y. and J. X. Yuan (2010). "Introduction to TRP channels: structure, function, and regulation." Adv Exp Med Biol 661: 99-108.

106

Souslova, V., P. Cesare, Y. Ding, A. N. Akopian, L. Stanfa, R. Suzuki, K. Carpenter, A. Dickenson, S. Boyce, R. Hill, D. Nebenuis-Oosthuizen, A. J. Smith, E. J. Kidd and J. N. Wood (2000). "Warm-coding deficits and aberrant inflammatory pain in mice lacking P2X3 receptors." Nature 407(6807): 1015-1017. Stripp, B. R., S. D. Reynolds, I. M. Boe, J. Lund, J. H. Power, J. T. Coppens, V. Wong, P. R. Reynolds and C. G. Plopper (2002). "Clara cell secretory protein deficiency alters clara cell secretory apparatus and the protein composition of airway lining fluid." Am J Respir Cell Mol Biol 27(2): 170-178. Suto, H., S. Nakae, M. Kakurai, J. D. Sedgwick, M. Tsai and S. J. Galli (2006). "Mast cell-associated TNF promotes dendritic cell migration." J Immunol 176(7): 4102- 4112. Symanowicz, P. T., G. Gianutsos and J. B. Morris (2004). "Lack of role for the vanilloid receptor in response to several inspired irritant air pollutants in the C57Bl/6J mouse." Neurosci Lett 362(2): 150-153. Szallasi, A. and P. M. Blumberg (1992). "Vanilloid receptor loss in rat sensory ganglia associated with long term desensitization to resiniferatoxin." Neurosci Lett 140(1): 51-54. Szallasi, A., F. Joo and P. M. Blumberg (1989). "Duration of desensitization and ultrastructural changes in dorsal root ganglia in rats treated with resiniferatoxin, an ultrapotent capsaicin analog." Brain Res 503(1): 68-72. Szolcsanyi, J. (1977). "A pharmacological approach to elucidation of the role of different nerve fibres and receptor endings in mediation of pain." J Physiol (Paris) 73(3): 251-259. Szolcsanyi, J. (2004). "Forty years in capsaicin research for sensory pharmacology and physiology." Neuropeptides 38(6): 377-384. Szolcsanyi, J., A. Szallasi, Z. Szallasi, F. Joo and P. M. Blumberg (1990). "Resiniferatoxin: an ultrapotent selective modulator of capsaicin-sensitive primary afferent neurons." J Pharmacol Exp Ther 255(2): 923-928. Talavera, K., M. Gees, Y. Karashima, V. M. Meseguer, J. A. Vanoirbeek, N. Damann, W. Everaerts, M. Benoit, A. Janssens, R. Vennekens, F. Viana, B. Nemery, B. Nilius and T. Voets (2009). "Nicotine activates the chemosensory cation channel TRPA1." Nat Neurosci 12(10): 1293-1299. Talavera, K., K. Yasumatsu, T. Voets, G. Droogmans, N. Shigemura, Y. Ninomiya, R. F. Margolskee and B. Nilius (2005). "Heat activation of TRPM5 underlies thermal sensitivity of sweet taste." Nature 438(7070): 1022-1025. Tizzano, M., M. Cristofoletti, A. Sbarbati and T. E. Finger (2011). "Expression of taste receptors in solitary chemosensory cells of rodent airways." BMC Pulm Med 11: 3. Tizzano, M., G. Dvoryanchikov, J. K. Barrows, S. Kim, N. Chaudhari and T. E. Finger (2008). "Expression of Galpha14 in sweet-transducing taste cells of the posterior tongue." BMC Neurosci 9: 110.

107

Tizzano, M., B. D. Gulbransen, A. Vandenbeuch, T. R. Clapp, J. P. Herman, H. M. Sibhatu, M. E. Churchill, W. L. Silver, S. C. Kinnamon and T. E. Finger (2010). "Nasal chemosensory cells use bitter taste signaling to detect irritants and bacterial signals." Proc Natl Acad Sci U S A 107(7): 3210-3215. Tominaga, M., M. J. Caterina, A. B. Malmberg, T. A. Rosen, H. Gilbert, K. Skinner, B. E. Raumann, A. I. Basbaum and D. Julius (1998). "The cloned capsaicin receptor integrates multiple pain-producing stimuli." Neuron 21(3): 531-543. Tso, T.C. "Tobacco." In: Elvers B, Hawkins S,Schulz G, editors. Ullmann's Encyclopedia of Industrial Chemistry. fifth ed. Weinheim, Germany:Wiley-VCH, 2000; A27. : 123–146. Undem, B. J. and M. J. Carr (2001). "Pharmacology of airway afferent nerve activity." Respir Res 2(4): 234-244. Urb, M. and D. C. Sheppard (2012). "The role of mast cells in the defence against pathogens." PLoS Pathog 8(4): e1002619. Vaigot, P., A. Pisani, Y. M. Darmon and J. P. Ortonne (1987). "The majority of epidermal Merkel cells are non-proliferative: a quantitative immunofluorescence analysis." Acta Derm Venereol 67(6): 517-520. Van Delden, C. and B. H. Iglewski (1998). "Cell-to-cell signaling and Pseudomonas aeruginosa infections." Emerg Infect Dis 4(4): 551-560. Van Hees, J. and J. M. Gybels (1972). "Pain related to single afferent C fibers from ." Brain Res 48: 397-400. Van Lommel, A., T. Bolle, W. Fannes and J. M. Lauweryns (1999). "The pulmonary neuroendocrine system: the past decade." Arch Histol Cytol 62(1): 1-16. Van Winkle, L. S., M. V. Fanucchi, L. A. Miller, G. L. Baker, L. J. Gershwin, E. S. Schelegle, D. M. Hyde, M. J. Evans and C. G. Plopper (2004). "Epithelial cell distribution and abundance in rhesus monkey airways during postnatal lung growth and development." J Appl Physiol 97(6): 2355-2363. Vanbeek, T. A., P. Maas, B. M. King, E. Leclercq, A. G. J. Voragen and A. Degroot (1990). "Bitter Sesquiterpene Lactones from Chicory Roots." Journal of Agricultural and Food Chemistry 38(4): 1035-1038. Venge, P. (1994). "Soluble markers of allergic inflammation." Allergy 49(1): 1-8. Vijayaraghavan, R., M. Schaper, R. Thompson, M. F. Stock and Y. Alarie (1993). "Characteristic modifications of the breathing pattern of mice to evaluate the effects of airborne chemicals on the respiratory tract." Archives of Toxicology 67(7): 478-490. Voedisch, S., S. Rochlitzer, T. Z. Veres, E. Spies and A. Braun (2012). "Neuropeptides control the dynamic behavior of airway mucosal dendritic cells." PLoS One 7(9): e45951.

108

Vriens, J., H. Watanabe, A. Janssens, G. Droogmans, T. Voets and B. Nilius (2004). "Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4." Proc Natl Acad Sci U S A 101(1): 396-401. Wang, Y. Y., R. B. Chang, S. D. Allgood, W. L. Silver and E. R. Liman (2011). "A TRPA1-dependent mechanism for the pungent sensation of weak acids." J Gen Physiol 137(6): 493-505. Wang, Z. Y., E. B. Olson, Jr., D. E. Bjorling, G. S. Mitchell and G. E. Bisgard (2008). "Sustained hypoxia-induced proliferation of carotid body type I cells in rats." J Appl Physiol 104(3): 803-808. Wershil, B. K., T. Murakami and S. J. Galli (1988). "Mast cell-dependent amplification of an immunologically nonspecific inflammatory response. Mast cells are required for the full expression of cutaneous acute inflammation induced by phorbol 12-myristate 13-acetate." J Immunol 140(7): 2356-2360. Whitear, M. (1965). "Presumed Sensory Cells in Fish Epidermis." Nature 208(5011): 703-704. Whitear, M. (1992). Solitary chemosensory cells. Fish Chemoreception. T. J. Hara. London, Chapman & Hall: 103-125. Widdicombe, J. (2001). "Airway receptors." Respir Physiol 125(1-2): 3-15. Wilson, S. R., A. M. Nelson, L. Batia, T. Morita, D. Estandian, D. M. Owens, E. A. Lumpkin and D. M. Bautista (2013). "The ion channel TRPA1 is required for chronic itch." J Neurosci 33(22): 9283-9294. Wong, G. T., K. S. Gannon and R. F. Margolskee (1996). "Transduction of bitter and sweet taste by gustducin." Nature 381(6585): 796-800. Wu, S. V., N. Rozengurt, M. Yang, S. H. Young, J. Sinnett-Smith and E. Rozengurt (2002). "Expression of bitter taste receptors of the T2R family in the gastrointestinal tract and enteroendocrine STC-1 cells." Proc Natl Acad Sci U S A 99(4): 2392-2397. Xiang, Z., X. Bo and G. Burnstock (1998). "Localization of ATP-gated P2X receptor immunoreactivity in rat sensory and sympathetic ganglia." Neurosci Lett 256(2): 105-108. Xu, H., M. Delling, J. C. Jun and D. E. Clapham (2006). "Oregano, thyme and clove- derived and skin sensitizers activate specific TRP channels." Nat Neurosci 9(5): 628-635. You, Y., E. J. Richer, T. Huang and S. L. Brody (2002). "Growth and differentiation of mouse tracheal epithelial cells: selection of a proliferative population." Am J Physiol Lung Cell Mol Physiol 283(6): L1315-1321. Zeiss, C. J. and E. A. Johnson (2004). "Proliferation of Microglia, but not Photoreceptors, in the Outer Nuclear Layer of the rd-1 Mouse." Investigative Ophthalmology & Visual Science 45(3): 971-976.

109

Zheng, J. (2013). "Molecular mechanism of TRP channels." Compr Physiol 3(1): 221- 242.