Novel Aspects of Ciliary Beat Regulation in Human Bronchial Epithelial Cells

Ph.D. Theses

Zoltán Süttő M.D.

Semmelweis University Budapest School of Ph.D. Studies in Clinical Medicine

Tutor: Gábor Horváth M.D., Ph.D. Reviewers: Attila Szabó M.D., Ph.D. Tamás Major jr. M.D., Ph.D. Scientific Committee:

President: Prof. Imre Hutás M.D., D.Sc. Members: Attila Somfay M.D., Ph.D

Prof. György Böszörményi Nagy M.D., Ph.D.

Budapest 2006.

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TABLE OF CONTENTS

ABBREVIATIONS.......................................................................................................... 5

1. INTRODUCTION........................................................................................................ 8

1.1. Anatomy and structure of the mucociliary apparatus .......................................... 8

1.1.1. Bronchial epithelium..................................................................................... 9 1.1.2. Airway surface liquid.................................................................................... 9 1.1.3. Cilia ............................................................................................................. 10

1.2. Regulation of the mucociliary apparatus ........................................................... 11

1.2.1. Neuronal regulation..................................................................................... 11 1.2.2. Effect of autonomic agonists on the mucociliary apparatus ....................... 12

1.2.2.1 Cholinergic agonists ........................................................................... 12 1.2.2.2 β-adrenergic agonists.......................................................................... 13 1.2.2.3 Other mediators .................................................................................. 14

1.2.3. Autocrine and paracrine regulation............................................................. 15 1.2.3.1 Nucleotides ......................................................................................... 15 1.2.3.2 Acetylcholine...................................................................................... 15

1.3. Intracellular signaling of ciliary beating: role of cAMP and Ca2+.................... 18

1.3.1. cAMP .......................................................................................................... 18 1.3.2. Calcium ....................................................................................................... 19 1.3.3. Interplay between cAMP and [Ca2+]i .......................................................... 19 1.3.4. Synergism between cAMP and [Ca2+]i ....................................................... 21 1.3.5. Soluble adenylyl cyclase (sAC): a unique source of cAMP ....................... 21

1.3.5.1 Enzymatic characteristics and cellular distribution of sAC................ 21 1.3.5.2 Cytosolic ions: role of HCO3

- and H+ in ciliary activity .................... 22

1.4. : Regulation of catecholamine concentration at adrenergic receptor site: role of catecholamine transporters ................................................................................ 23

2. AIMS .......................................................................................................................... 26

3. MATERIALS AND METHODS ............................................................................... 27

3.1. Chemicals ........................................................................................................... 27

3.2. Solutions ............................................................................................................. 27

3.3. Cell isolation and culture techniques ................................................................. 28

3.3.1. Preparation of submerged tracheal epithelial cell cultures.......................... 28 3.3.2. Preparation of air liquid interface cultures of tracheal epithelium.............. 29

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3.3.3. Human bronchial arterial smooth muscle cell isolation.............................. 30 3.3.4. Preparation of primary cultures of bronchial arterial SMCs....................... 31 3.3.5. Caki-1 cell culture ....................................................................................... 31

3.4. Selective permeabilization of the basolateral membrane of cells grown at the ALI ...................................................................................................................... 31

3.5. Measurement of CBF.......................................................................................... 32

3.6. Measurement of pHi............................................................................................ 33

3.6.1. Fluorometric pHi measurements ................................................................. 33 3.6.2. Calibration of pHi measurements ................................................................ 33

3.6.2.1 Preparation of the pH calibration curve.............................................. 33 3.6.2.2 pHi estimation..................................................................................... 34

3.6.3. Experimental procedures for pHi measurements......................................... 35

3.7. Measurement of [Ca2+]i...................................................................................... 35

3.8. Simultaneous measurement of CBF and pHi or [Ca2+]i ..................................... 36

3.9. mRNA-expression of sAC and catecholamine transporters................................ 36

3.9.1. RT-PCR....................................................................................................... 36 3.9.2. Cloning of cDNA fragments of sAC variants ............................................. 37

3.10. Expression of sAC-protein................................................................................ 38

3.10.1. Western blotting for sAC .......................................................................... 38 3.10.2. sAC-specific immunohystochemistry and immunocytochemistry ........... 38

3.11. Single-cell, fluorometric NE uptake measurements.......................................... 39

3.11.1. NE uptake experiments ............................................................................. 39 3.11.2. NE uptake assay ........................................................................................ 39

3.12. Immunochemical detection of a plasma membrane binding site for corticosterone in bronchial arterial SMC .......................................................... 40

3.13. Statistical analysis ............................................................................................ 41

4. RESULTS................................................................................................................... 42

4.1. Regulation of human airway ciliary beat frequency by pHi ............................... 42

4.1.1. The effect of changing pHi on CBF during ammonium prepulse ............... 42 4.1.2. The effect of changing pHi on CBF during removal of extracellular CO2.. 44 4.1.3. Inhibition and stimulation of PKA does not prevent the effect of pHi on

CBF.............................................................................................................. 46

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4.1.4. Inhibition of protein phosphatases does not influence the effect of pHi on CBF.............................................................................................................. 48

4.1.5. pHi does not regulate CBF via changes in [Ca2+]i....................................... 49 4.1.6. Basolaterally permeabilized cells grown at the ALI ................................... 52 4.1.7. Correlation between pHi and CBF in permeabilized cells .......................... 53

4.2. sAC expression and cellular distribution in HAECs .......................................... 54

4.2.1. sAC mRNA expression ............................................................................... 54 4.2.2. Expression and cellular distribution of sAC protein ................................... 57 4.2.3. Effect of cytosolic HCO3

- on ciliary beating in HAECs ............................. 59

4.3. NE transporter mRNA expression profile in human airway epithelial cells ...... 61

4.4. Pharmacological characterization of NE uptake ............................................... 62

4.4.1. NE uptake characteristics in bronchial arterial SMCs ................................ 62 4.4.2. Inhibition of NE uptake by GSs .................................................................. 64 4.4.3. EMT inhibition by corticosterone is a nongenomic effect.......................... 65

5. DISCUSSION............................................................................................................. 69

5.1. Regulation of human airway ciliary beat frequency by pHi ............................... 69

5.2. Expression of sAC in human airway epithelial cells .......................................... 73

5.3. Expression and putative function of catecholamine transporters in human airway epithelial cells......................................................................................... 76

6. CONCLUSIONS ........................................................................................................ 81

SUMMARY ................................................................................................................... 82

ÖSSZEFOGLALÁS ....................................................................................................... 84

REFERENCES ............................................................................................................... 86

PUBLICATIONS LIST................................................................................................ 111

Publications relevant to theses ................................................................................ 111

Other publications ................................................................................................... 112

ACKNOWLEDGEMENTS ......................................................................................... 115

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ABBREVIATIONS

acetyl-CoA acetyl coenzyme-A

ACh acetylcholine

AChE acethylcholinesterase

AKAP A-kinase anchoring protein

ALI air-liquid interface

ARS ATP regenerating system

ASL airway surface liquid

ATCC American Type Culture Collection

ATP adenosine 5'-triphosphate

ATPase adenosine triphosphatase

BCECF 2’7’-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein

BEGM bronchial epithelial growth medium

BK channel large conductance Ca2+-activated K+ channel

bp base pairs

BSA bovine serum albumin

[Ca2+]i intracellular calcium concentration

cAMP cyclic adenosine monophosphate

CBF ciliary beat frequency

cDNA complementary DNA

CFTR cystic fibrosis transmembrane conductance regulator

ChAT choline acetyltransferase

CHT choline high affinity transporter

COMT catecholamine-O-methyltransferase

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COPD chronic obstructive pulmonary disease

DAPI 4,6-diamidino-2-phenylindole

DIC differential-interference-contrast

DMEM Dulbecco’s modified Eagle’s medium

DNase deoxyribonuclease

EMT extraneuronal monoamine transporter

F fluorescence

FBS fetal bovine serum

FFT fast Fourier transform

Fn mean fluorescence intensity value

GAPDH glyceraldehyde-3-phosphate dehydrogenase

GS stimulatory G-protein

GS glucocorticosteroid

HAEC human airway epithelial cell

[HCO3-]i intracellular bicarbonate concentration

HRP horseradish peroxidase

IC50 inhibitor concentration at 50% inhibition

IP3 inositol trisphosphate

KD dissociation constant

Ki inhibition constant

Km Michaelis constant (substrate concentration at half-

maximal enzyme velocity)

MAO monoamine oxidase

mRNA messenger RNA

NE norepinephrine

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NET norepinephrine transporter

NFR normalized fluorescence ratio

OCT organic cation transporter

PBS phosphate-buffered saline

PCL periciliary liquid layer

PIP2 phosphatidyl inositol disphosphate

PKA cAMP-dependent protein kinase

PLC phospholipase C

RAR rapidly adapting stretch receptor

ROI region of interest

RT-PCR reverse transcriptase polymerase chain reaction

sAC soluble adenylyl cyclase

SEM standard error of the mean

SMC smooth muscle cell

SPG sucrose-potassium phosphate-glyoxylic acid

Tm annealing temperature

tmAC transmembrane adenylyl cyclase

TRITC tetramethylrhodamine isothiocyanate

TSA tyramide signal amplification

UTP uridine 5'-triphosphate

VIP vasoactive intestinal peptide

Vmax maximal enzyme velocity

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1. INTRODUCTION

1.1. Anatomy and structure of the mucociliary apparatus

The ventilation of the lung ranges between 1,000 and 21,000 liters per 24 hours in

humans depending on body size and physical activity. This exposes the extensive

epithelial surface of the respiratory tract to a large burden of microorganisms as well as

inorganic and organic particulate and gaseous material. A series of defense systems

protect the airways against these potentially injurious actions. First, the mucociliary

apparatus serves as a mechanical barrier by trapping particulates in the surface liquid

and removing them from the airway. Second, a number of macromolecules secreted by

the airway epithelial cells, e.g. lysozyme, defensin, lactoferrin, leukocyte protease

inhibitor and lactoperoxidase, exhibits antimicrobial effects and thus, form a „chemical

shield”. Third, the surface liquids provide a „biologic barrier” function by interacting

with microorganisms and luminal inflammatory cells, thereby preventing them from

adhering to and migrating through the airway epithelium. The different defense systems

work together and are complementary to each other (Wanner et al., 1996; Chilvers &

O'Callaghan, 2000; Knowles & Boucher, 2002).

The present work will focus on the regulation of ciliary beat frequency (CBF), the

primary determinant of the effectiveness of the mucociliary apparatus and mucus

clearance. Mucociliary dysfunction is a frequent consequence of chronic airway

diseases such as asthma (Foster et al., 1982; Bateman et al., 1983a; Pavia et al., 1985;

O'Riordan et al., 1992), chronic bronchitis (Camner et al., 1973; Santa Cruz et al.,

1974b; Goodman et al., 1978; Vastag et al., 1986), and cystic fibrosis (Yeates et al.,

1976; Matthys & Kohler, 1986; Regnis et al., 1994), but can cause by itself airway

disease as illustrated by primary ciliary dyskinesia (Kollberg et al., 1978; Baum et al.,

1990; Regnis et al., 2000). Thus, the better understanding of CBF regulation could

provide useful therapeutic tools in restoring normal clearance in airway diseases.

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1.1.1. Bronchial epithelium

Although the airway epithelium contains a number of resident cells, only ciliated

columnar cells and secretory cells in the surface epithelium and in submucosal glands

seem to contribute directly to mucociliary function (Wanner et al., 1996). The bronchial

tree is lined by a ciliated epithelium extending from the proximal trachea to the terminal

bronchioles. Surface secretory cells in the central airways referred to as goblet cells are

mainly mucous cells, whereas the secretory cells of submucosal glands are either serous

or mucous. In central airways, the ratio of ciliated columnar cells to goblet cells is

approximately 5:1 (Rhodin, 1966). Whereas the surface epithelium in the central

airways is pseudostratified and the cells are columnar in shape, the surface epithelium in

the most peripheral bronchioles is characterized by cuboidal cells, is less densely

ciliated and mucus producing goblet cells are replaced with other secretory cells called

Clara cells (Wanner et al., 1996).

1.1.2. Airway surface liquid

The airway epithelium is covered by a surface liquid throughout its length

produced by the airway secretory cells. This liquid layer, referred to airway surface

liquid (ASL), consists of at least two layers in the central airways: a mucus layer (which

is missing in the most peripheral airways) and a periciliary liquid layer (PCL) (Wanner

et al., 1996; Knowles & Boucher, 2002). The mucus layer contains high–molecular

weight, heavily glycosylated macromolecules, that behave as a tangled network of

polymers. The properties of the mucin gel are determined by this tangled network,

which reflects, in part, the water content, ion concentrations, and pH of the ASL. It

appears that mucin macromolecules are well adapted to binding and trapping inhaled

particles for clearance from the lung, at least in part because of the extraordinary

diversity of their carbohydrate side chains. Because they provide a combinatorial library

of carbohydrate sequences, mucins can bind to virtually all particles that land on airway

epithelia and can thus clear them from the lung. It is still not known whether the mucus

layer is continuous or discontinuous in human airways in vivo (Knowles & Boucher,

2002).

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The mucus-free zone at the cell surface, which approximates the height of the

outstretched cilia, is the PCL, also known as "sol" layer. PCL is crucial both because it

provides a low-viscosity solution in which cilia can beat rapidly and because it shields

the epithelial cell surface from the overlying mucus layer. The absence of the PCL

allows adhesive interactions between mobile mucins in the mucus layer and tethered

cell-surface mucins - the so called "glycocalyx", which covers the apical surface of

airway epithelial cells - greatly reducing the efficiency of mucociliary and cough

clearance (Knowles & Boucher, 2002).

1.1.3. Cilia

The surface of each tracheal columnar cell contains approximately 200 cilia with

an average length of 6 μm and a diameter of 0.1-0.2 μm. With an increase in airway

generations, there is a decrease in the relative number of ciliated cells, which are

characterized by fewer and shorter cilia than in the large bronchi (Wanner et al., 1996).

The schematic structure of a normal cilium is depicted in Figure 1-1. The typical 9 + 2

microtubular structure is a consistent feature of all known cilia. The microtubules are

constructed primarily from dynein and heterodimers of α- and β-tubulin, but more than

200 other proteins are associated with a variety of circumferential, spoke, and radial

linkages that serve to maintain the axonemal structure and possibly have a role in the

control of ciliary movement (Wanner et al., 1996; Chilvers & O'Callaghan, 2000).

Figure 1-1. The schematic structure of a normal cilium in cross section as seen from the ciliary tip towards the base. Two central microtubules (CT) are enclosed within a central sheath to form the central axis of the axoneme. This is surrounded by nine microtubule doublets (MD). The doublets consist of an A and B subfiber. Each doublet is connected by nexin links (N) to the next. From the A doublet, inner (IA) and outer (OA) dynein arms project to the adjacent doublet. Radial spokes (RS) connect the central sheath to the outer microtubule doublets (Chilvers & O'Callaghan, 2000).

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The outer nine doublet microtubules consist of a complete A subfiber (13

protofilaments) upon which an incomplete B subfiber (10 to 11 protofilaments)

assembles. The nine outer doublet microtubules are connected to each other by nexin

links, and radial spokes connect the outer microtubules with projections originating

from the inner singlet microtubules. Dynein, the motor for ciliary movement, is an

adenosine triphosphatase (ATPase). An outer and an inner dynein arm originate from

each A subfiber of the outer doublet microtubule and project in a counterclockwise

direction (as seen from the tip) toward the B subfiber of the next doublet. The

mammalian outer arm dynein is a two-headed structure; less is known about the

morphology of inner arm dynein. Each dynein head contains a heavy chain ATPase to

which several dynein light chains and at least one intermediate chain are attached. These

chains anchor the dynein arm to the A microtubule and are involved in the regulation of

ciliary beating. At the tip of the cilium, the nine outer doublets simplify into a single A

fiber, which inserts into a transmembrane complex called the ciliary crown. The ciliary

crown carries three to seven short “claws”, which probable function is to mechanically

or chemically engage the overlying mucus layer during the effective ciliary stroke. At

the base, the nine outer doublets end in the basal body, which is anchored to the

cytoskeleton. The outer ciliary membrane covering the axoneme is continuous with the

cell surface membrane. Bridges between the ciliary membrane and the outer doublets

exist along the entire length of the cilium (Wanner et al., 1996; Chilvers & O'Callaghan,

2000).

1.2. Regulation of the mucociliary apparatus

1.2.1. Neuronal regulation

The mucociliary apparatus is innerved predominantly by the parasympathetic

nervous system. Catecholamine and vasoactive intestinal peptide (VIP) containing nerve

fibers as well as a rich network of cholinergic nerves have been shown in submucosal

glands, but not within the surface epithelium, of human bronchi (Partanen et al., 1982;

Laitinen, 1985a, b; Laitinen et al., 1985). Apparently, there are no efferent neurons

within the surface epithelium but this issue has not been fully clarified (Wanner et al.,

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1996). Instead, electron microscopic studies have revealed intraepithelial sensory

(afferent) axons, either close to the lumen or near the basement membrane of human

bronchi (Laitinen, 1985b). These fibers are primarily unmyelinated capsaicin sensitive

C-fibers or thin, myelinated Aδ axons of rapidly adapting stretch receptors (RAR) and

are thought to participate in defensive autonomic reflex mechanisms like mucus

hypersecretion, cough and bronchoconstriction in response to certain chemical or

mechanical stimuli. Excitation of C-fiber endings may cause local axon reflex, a feature

also referred to as the “efferent” function of sensory nerves, resulting in the release of

neuropeptides (Wanner et al., 1996; Belvisi, 2002; Widdicombe, 2003).

It appears that baseline mucociliary function is not under autonomic control. For

example, the sympathetic and parasympathetic nervous system seems to lack an effect

on baseline glycoprotein secretion in cat (Wanner et al., 1996). However, both vagal

and sympathetic nerve stimulation as well as excitation of sensory nerve endings might

alter mucosal function by increasing mucus secretion and water transport towards the

airway (Wanner et al., 1996; Belvisi, 2002). In contrast to secretory responses, very

little is known about the effects of autonomic nerve stimulation on ciliary activity or

mucociliary clearance in the airways. Vagal section or vagal stimulation of dogs seems

to have no effect on in vivo ciliary activity in the large airways (Wanner et al., 1996).

1.2.2. Effect of autonomic agonists on the mucociliary apparatus

1.2.2.1 Cholinergic agonists

Although airway surface epithelium is not innerved by cholinergic nerves, these

cells express muscarinic receptors, similarly to submucosal gland cells. Human

bronchial epithelial cells express both M1 and M3 muscarinic receptor subtypes (Racke

& Matthiesen, 2004) while human submucosal gland cells express only M3 receptors

(Mak et al., 1992). M1 receptors seem to be restricted to the peripheral airways < 2 mm

(Racke et al., 2006). Furthermore, functionally active nicotinic receptors have recently

been demonstrated on human airway epithelial cells (Maus et al., 1998; Wang et al.,

2001; Wessler & Kirkpatrick, 2001; Carlisle et al., 2004).

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Cholinergic antagonists, likewise other autonomic receptor antagonists, generally

do not influence baseline mucociliary clearance (Mercke et al., 1982; Wanner et al.,

1996). Atropine is a notable exception, which, in contrast to another anticholinergic

agent ipratropium bromide, depressed airway mucociliary transport in most studies,

probably by cilioinhibition (Foster et al., 1976; Mercke et al., 1982; Whiteside et al.,

1984; Groth et al., 1991; Wanner et al., 1996), yet this finding is not consistent

(Hybbinette & Mercke, 1982b; Mercke et al., 1982). However, almost all in vitro and in

vivo studies have found an enhancement of mucociliary clearance and ciliary activity by

the cholinergic agonists acetylcholine (ACh) and methacholine, but not vagal

stimulation, in different species (Cammer et al., 1974; Berger et al., 1978; Hybbinette &

Mercke, 1982b; Mercke et al., 1982; Wong et al., 1988a, b; Seybold et al., 1990;

Salathe & Bookman, 1995; Wanner et al., 1996). Cholinergic ciliostimulation in sheep

is mediated by M3 receptors coupled to pertussis-toxin-insensitive G-proteins involving

Ca2+-release from inositol trisphosphate (IP3) sensitive Ca2+-stores (Salathe &

Bookman, 1995; Salathe et al., 1997; Salathe & Bookman, 1999; Salathe et al., 2001).

1.2.2.2 β-adrenergic agonists

As mentioned above, microscopic studies have failed to show efferent autonomic

innervation in the surface epithelium of human bronchi (Partanen et al., 1982), however,

there is a high density of β-adrenergic receptors on human airway epithelial cells

(HAEC): surface cells express mainly β2-adrenergic receptors, whereas submucosal

gland cells express both β1- and β2-adrenergic receptors (Wanner et al., 1996).

Therefore it is not surprising that β-adrenergic receptor agonists, which are primarily

used as bronchodilators in the treatment of obstructive airway diseases, also influence

airway epithelial cell function. Besides opening apical ion channels, regulating cytokine

secretion and enhancing epithelial wound repair, one of their most important effects, at

least in terms of mucociliary clearance, is ciliostimulation (Salathe, 2002).

A number of clinical studies have demonstrated the short term stimulatory effect

of β-adrenergic receptor agonists on mucociliary clearance (Santa Cruz et al., 1974a;

Mossberg et al., 1976; Foster et al., 1982; Weiss et al., 1983; Vastag & Zsamboki,

1985; Matthys & Kohler, 1986; Yeates et al., 1986; Gatto, 1993). Most of the clinical

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studies have proved that β-adrenergic receptor agonists augment mucociliary clearance

of healthy individuals (Tay et al., 1997; Svartengren et al., 1998), COPD and asthma

patients (Fazio & Lafortuna, 1981; Matthys et al., 1987; Hasani et al., 2003; Hasani et

al., 2005; Bennett et al., 2006) and animals (Hybbinette & Mercke, 1982a; Mercke et

al., 1982; Sabater et al., 2005) (but not delta F508 homozygous cystic fibrosis patients

(Mortensen et al., 1993)). However, some report contrast these results (Pavia et al.,

1980; Bateman et al., 1983b; Davies & Webster, 1989; Svartengren et al., 1998). The

interpretation of radioaerosol studies on mucociliary clearance is complicated by the

bronchodilatory effect of β-adrenergic receptor agonists, which permit a greater

penetration of the radioaerosol into the smaller airways where mucociliary clearance is

markedly slower (Salathe et al., 1996). This methodological problem is eliminated when

not the clearance of labeled particles but CBF, the primary determinant of mucociliary

clearance rate, is measured. The results of these studies are consistent in proving that β-

adrenergic receptor agonists increase CBF of mammalian, including human, airway

epithelial cells in vitro (Verdugo et al., 1980; Wong et al., 1988a; Sanderson & Dirksen,

1989; Devalia et al., 1992; Ingels et al., 1992; Tamaoki et al., 1993; Kanthakumar et al.,

1994; Lindberg et al., 1995; Frohock et al., 2002; Salathe, 2002; Allen-Gipson et al.,

2004; Shiima-Kinoshita et al., 2004; Piatti et al., 2005; Zhang et al., 2005) as well as in

vivo (Wong et al., 1988b; Lindberg et al., 1995).

1.2.2.3 Other mediators

α-adrenergic receptors are expressed in submucosal gland cells, mainly in serous

cells. Peptidergic receptors such as those mediating the effects of VIP and substance P

(neurokinin receptors) have also been identified on airway epithelial cells in animals

and in humans. In vagally induced mucus hypersecretion, neuropeptides also seem to be

involved, besides ACh (Wanner et al., 1996). Ephedrine has been reported to increase

CBF by some but not all investigators. Reported effects of alpha-adrenergic agonists on

CBF are conflicting: stable, decreasing and increasing CBF have all been reported.

Peptide mediators like substance P has a slight ciliostimulatory effect, but this has not

been found consistently, while neuropeptide Y was found to decrease CBF (Wanner et

al., 1996).

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1.2.3. Autocrine and paracrine regulation

Recent data suggest that autocrine and paracrine mediators like 5' nucleotides and

ACh may have a role in the regulation of mucociliary clearance.

1.2.3.1 Nucleotides

As mentioned above, neuronal mechanisms are probably not involved in the

regulation of CBF, and the role of endocrine mechanisms are also unlikely as systemic

responses, for example to epinephrine, have time constants that are too long for this

function (Lazarowski et al., 2000; Lazarowski & Boucher, 2001; Knowles & Boucher,

2002; Lazarowski et al., 2003). Adenosine 5'-triphosphate (ATP) and uridine 5'-

triphosphate (UTP) are released by the airway epithelial cells across both the apical and

basolateral membrane in response to mechanical and osmotic stresses (Homolya et al.,

2000) and also under basal conditions. Released nucleotides interact with purinoceptors

at both membranes to trigger Ca2+-mobilization. Apical purinoceptors modulate cellular

functions like ciliary beating (Morse et al., 2001; Lieb et al., 2002), mucus secretion

(Knowles et al., 1991; Davis et al., 1992; Lethem et al., 1993) and ion channel activities

(Korngreen et al., 1998), i.e. the elements of the mucociliary apparatus, while

basolateral nucleotide release constitutes a paracrine mechanism by which mechanical

stresses signal adjacent cells not only within the epithelium, but also in the submucosa.

Airways also express a large number of extracellular nucleotidases that regulate

extracellular nucleotide concentrations. Furthermore, the hydrolysates of 5' nucleotides

also serve as ligands for specific receptors. These data strongly suggest that nucleotide

release by epithelial cells regulates mucus clearance rates in response to luminal stresses

(Knowles & Boucher, 2002; Picher & Boucher, 2003; Picher et al., 2003).

1.2.3.2 Acetylcholine

ACh represents the “classical” neurotransmitter of the parasympathetic nervous

system, but evidence has been provided that ACh is also synthesized and secreted by a

variety of non-neuronal tissues including human bronchial epithelium (Wessler &

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Kirkpatrick, 2001; Proskocil et al., 2004). In fact, more or less all human cells fulfill the

three crucial conditions for the synthesis of acetylcholine: presence of the synthesizing

enzyme choline acetyltransferase (ChAT) and the substrates choline and acetyl

coenzyme-A (acetyl-CoA) (Wessler & Kirkpatrick, 2001).

The bronchial epithelium, however, not only secretes ACh, but also possesses all

the other elements essential for cholinergic regulation (Proskocil et al., 2004; Lips et al.,

2005), a system referred to as non-neuronal cholinergic system. While acetyl-CoA is

produced in the citric acid cycle, choline, the other substrate of ACh synthesis is

transported into the cells by specific transporters, which is the rate-limiting step in ACh

synthesis (Lips et al., 2003). Previously it was thought that choline high affinity

transporter (CHT) is restricted to the neurons, while non-neuronal cells express only a

low affinity choline uptake system (Wessler & Kirkpatrick, 2001). Recently CHT has

been shown to be expressed in human bronchial epithelial cells as well (Proskocil et al.,

2004). Immunolocalization has revealed that CHT is restricted to the apical membrane

of ciliated cells in rats (Pfeil et al., 2003). ChAT, the ACh-synthesizing enzyme, is

expressed in the surface epithelial and submucosal gland cells of human airways. The

most prominent ChAT immunoreactivity in ciliated bronchial cells has been shown at

basal bodies, where the cilia are embedded (Wessler & Kirkpatrick, 2001). ACh-release

from non-neuronal cells seems to be mediated by organic cation transporters (OCT),

especially by the isoforms OCT-1 and OCT-2 (Wessler & Kirkpatrick, 2001; Wessler et

al., 2003; Lips et al., 2005). OCTs, as discussed below in paragraph 1.4, are

polyspecific cation transporters and are best known about their role in cellular

catecholamine uptake. All three isoforms of OCT has been shown to be expressed in

human bronchial epithelium and to be localized (but not restricted) to the apical

membrane of ciliated cells (Lips et al., 2005). The effectors of ACh regulation, i.e.

muscarinic as well as nicotinic ACh-receptors are also expressed in HAEC (Mak et al.,

1992; Wanner et al., 1996; Racke & Matthiesen, 2004). The inactivating enzyme

acethylcholinesterase (AChE) has also been demonstrated in monkey bronchial

epithelial cell culture by virtue of its catalytic activity (Proskocil et al., 2004).

There are some basic differences between neuronal and non-neuronal cholinergic

systems. Non-neuronal cells are not endowed with effective storing organelles like

cholinergic vesicles and thus fail to concentrate ACh. In contrast to neuronal exocytotic

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ACh-release, non-neuronal ACh release via OCTs seems to be continuously active upon

concentration gradient (Wessler et al., 2001; Lips et al., 2005). Furthermore, in contrast

to the cholinergic synapse, there are not known “hot spots” of highly concentrated ACh-

receptors; and finally, the elimination of non-neuronal ACh is much slower because of

low cholinesterase activity (Wessler et al., 2003).

These features of non-neuronal cholinergic system agree with the concept that

non-neuronal ACh acts continuously as a local signaling molecule involved in the

regulation of basic cell functions (Wessler et al., 2003), however, the physiological

relevance of airway non-neuronal cholinergic system remains to be determined. In

airway epithelium, there are substantial speculations concerning its function including

regulation of inflammatory mediator release from epithelial cells and contribution to the

regulation of epithelial layer integrity by the control of epithelial cell adhesion, cell–cell

interactions and proliferation (Racke & Matthiesen, 2004). Although non-neuronal ACh

is considered to be a local mediator, a recent study suggests that ACh secreted by the

mouse airway epithelium causes bronchoconstriction (Moffatt et al., 2004), however,

the specificity of this result has been questioned (Racke et al., 2006).

Several data mentioned above suggest that nonneuronal ACh might be involved in

CBF regulation. First of all, ACh is a well known regulator of CBF in mammalian

ciliated bronchial epithelial cells via M3 muscarinic receptor (Salathe et al., 1997),

though there is no cholinergic innervation in the surface epithelium (Partanen et al.,

1982). Furthermore, the subcellular distribution of ChAT in airway ciliated cells

characterized by an intense immunoreactivity close to the basal bodies of cilia (Racke &

Matthiesen, 2004) also suggests a functional relationship between non-neuronal ACh

and CBF regulation. The immunolocalization of OCTs and CHT in the apical

membrane of ciliated cells also supports this hypothesis (Lips et al., 2005). In addition

to bronchial epithelial cells, migrating immune cells in the airways also synthesize ACh

(Wessler & Kirkpatrick, 2001). Whether ACh released by these cells is involved in CBF

regulation remains to be answered.

18

1.3. Intracellular signaling of ciliary beating: role of cAMP and Ca2+

External stimuli affecting ciliary beating in HAECs are transduced mainly by two

intracellular second messengers, cAMP and intracellular calcium.

1.3.1. cAMP

Intracellular cAMP plays a crucial role in the regulation of CBF. Cytoplasmic

cAMP levels have been modulated in vitro by β2-adrenergic agonists that stimulate

transmembrane adenylyl cyclase (tmAC), forskolin that directly activates tmAC, and

membrane permeable cAMP analogues (e.g., Verdugo et al., 1980; Sanderson &

Dirksen, 1989; Tamaoki et al., 1989; Di Benedetto et al., 1991b; Lansley et al., 1992;

Wyatt et al., 1998). Increasing intracellular cAMP has been shown to increase human,

bovine, ovine, and rabbit CBF, and this increase was blockable by a broad specificity

protein kinase inhibitor implicating cAMP-dependent protein kinase (PKA) (Yang et

al., 1996; Wyatt et al., 1998; Salathe et al., 2000b; Frohock et al., 2002). The molecular

mechanism by which PKA increases mammalian CBF is likely analogous to the

stimulation of ciliary activity by cAMP in Paramecium. There, Hamasaki et al. (1989)

showed that PKA phosphorylated specific axonemal targets both in vitro and in

permeabilized cells. One with an apparent molecular weight of 29 kDa was extractable

by procedures used for the isolation of outer arm dynein and was identified to be a

dynein light chain (Hamasaki et al., 1991). The phosphorylation of this light chain

increases the velocity of microtubule gliding across dynein-coated surfaces in vitro and

the swimming speed of Paramecium in vivo (Hamasaki et al., 1991). In mammalian

respiratory cilia, axonemal PKA targets with similar molecular weights have been

identified by virtue of their cAMP-dependent phosphorylation (Salathe et al., 1993;

Sisson et al., 2000; Kultgen et al., 2002) and, in an ovine study, the target was found on

outer dynein arms (Salathe et al., 2000a). Since these dynein arms are responsible for

frequency regulation (Brokaw & Kamiya, 1987), this is the correct place for

phosphorylation targets to influence CBF. Experiments using isolated cilia also revealed

that PKA must be localized to cilia (Salathe et al., 1993; Kultgen et al., 2002), a fact

supported by the presence of an anti-PKA immunopositive protein in bovine axonemes

19

(Sisson et al., 2000) and a specific A-kinase anchoring protein (AKAP) in human

airway cilia (Kultgen et al., 2002).

1.3.2. Calcium

The mechanism by which CBF is adjusted by the intracellular calcium

concentration ([Ca2+]i) seems to differ between unicellular organisms and mammals. In

Paramecium, rising [Ca2+]i slows CBF to the point where the beat direction is reversed

(Naito & Kaneko, 1972). Mammalian cilia never reverse direction and [Ca2+]i rises

increase CBF (Verdugo, 1980; Girard & Kennedy, 1986; Sanderson & Dirksen, 1989;

Villalon et al., 1989; Sanderson et al., 1990; Di Benedetto et al., 1991a; Lansley et al.,

1992; Salathe & Bookman, 1995) whereas [Ca2+]i decreases lower CBF (Salathe &

Bookman, 1995). Recent evidence suggests that Ca2+ acts directly to change CBF, and

the Ca2+-binding protein is probably an integral part of the axonemal machinery

(Salathe & Bookman, 1999).

1.3.3. Interplay between cAMP and [Ca2+]i

The “classical” way to achieve intracellular cAMP production in HAECs is to

stimulate β-adrenergic receptors. β-adrenergic agonists augment CBF in airway

epithelial cells through a signaling cascade involving β2-adrenergic receptors, the α-

subunit of an adenylyl cyclase stimulatory G-protein (GS) to stimulate tmAC, cAMP,

PKA and the phosphorylation of a ciliary target protein. β3-adrenergic receptors might

also be involved but the data are conflicting (Tamaoki et al., 1993; Thomas & Liggett,

1993; Kelsen et al., 1995; Salathe, 2002). Nevertheless, a more complex signaling

pathway has also been observed in rabbit and ovine ciliated tracheal cells (Lansley et

al., 1992; Frohock et al., 2002; Zhang et al., 2005), where a calcium-coupled

acceleration of CBF, blockable by the inhibition of PKA, was also observed (left side of

the panel in Figure 1-2) (Frohock et al., 2002).

Conversely, cytoplasmic Ca2+ transients may stimulate cAMP production. Short-

term purinergic receptor stimulation of HAECs by ATP results in a transient increase of

[Ca2+]i through calcium-release from internal stores, which, in turn leads to a calcium-

20

coupled CBF peak. However, CBF fails to follow [Ca2+]i when the latter returns to

baseline. The prolonged increase in CBF after purinergic stimulation has been shown to

be the consequence of cAMP production by calcium-sensitive adenylyl cyclase(s) and

consecutive PKA-activation (Lieb et al., 2002). The exact isoforms of adenylyl cyclase

involved is not known. Proposed mechanism of interplay between the cAMP- and

[Ca2+]i-dependent signal transduction pathways is shown in Figure 1-2.

Figure 1-2. Interplay between cAMP- and [Ca2+]i-dependent signal transduction mechanisms in the regulation of CBF. Black arrows represent the primary mechanisms by which intracellular cAMP production and [Ca2+]i is increased. Red arrows depict the interplay between cAMP- and [Ca2+]i-dependent mechanisms. Left side of the panel: stimulation of β2-adrenergic receptors activates transmembrane adenylyl cyclase to produce cAMP, which activates an axonemal PKA on outer dynein arms (not shown). Activated PKA increases CBF by the phosphorylation of a dynein light chain. In some ciliated cells, cytosolic PKA activation also releases calcium from intracellular stores (red arrows), which increases CBF through an additional mechanism (Salathe, 2002). Right side of the panel: stimulation of purinergic receptor (P2Y) results in PLC activation and IP3-production, which in turn releases Ca2+ from internal stores. Ca2+ stimulates CBF probably acting directly on the axoneme. In HAECs, increasing [Ca2+]i also activates Ca2+-dependent adenylyl cyclase(s) to generate cAMP (red arrows) and thus, to stimulate CBF (question mark indicates that whether Ca2+-sensitive adenylyl cyclase is a transmembrane and/or soluble adenylyl cyclase is not known). Abbreviations: β2, β2-adrenergic receptor; IP3, inositol trisphosphate; PIP2, phosphatidyl inositol disphosphate; PKA, cAMP-dependent protein kinase; PLC, phospholipase C; P2Y, P2Y purinergic receptor.

+ATP cAMP

PKA

Ca2+

P2Y PLC

tmAC β2

IP3 +

PIP2

endoplasmic reticulum

cilia

+

Ca2+

?+

cilia

ted

airw

ay e

pith

elia

l cel

l

21

1.3.4. Synergism between cAMP and [Ca2+]i

Besides the interplay of the two second messengers described above, a

fundamental synergistic interrelationship between cytoplasmic cAMP and [Ca2+]i has

also been demonstrated in a recent study (Ma et al., 2002). In these experiments,

cytoplasmic fluid of patch-clamped rabbit ciliated tracheal cells was replaced by the

pipette solution and thus, was directly controlled. These experiments showed that

spontaneous beating requires only MgATP, whereas stimulation of ciliary motility

requires both Ca2+ and cyclic nucleotides, in addition to MgATP: in the absence of

cytoplasmic cAMP, [Ca2+]i rises alone do not augment CBF and, on the other hand, in

the absence of cytoplasmic Ca2+, cAMP exerts only a slight ciliostimulatory effect

compared to that in the presence of cytoplasmic Ca2+. While the physiological [Ca2+]i

seems to be sufficient for synergistic augmentation of cAMP’s effect on CBF, little is

known about the regulation of baseline cAMP production in the axoneme, a

permissive factor necessary for CBF stimulation by [Ca2+]i transients.

1.3.5. Soluble adenylyl cyclase (sAC): a unique source of cAMP

1.3.5.1 Enzymatic characteristics and cellular distribution of sAC

Transmembrane adenylyl cyclase (tmAC), the commonly found adenylyl cyclase

in mammalian cells is sensitive to G-protein and Mg2+, can be activated by forskolin

and has a transmembrane domain. Another, distinct adenylyl cyclase activity was

described in testis (Braun & Dods, 1975; Braun et al., 1977; Neer, 1978) and called

soluble adenylyl cyclase (sAC). sAC differs from tmAC enzymatically by its 10-fold

lower affinity for substrate (Neer, 1978) and its insensitivity to G-protein activation or

forskolin (Buck et al., 1999). Rat testicular (Buck et al., 1999) and human sAC (Geng et

al., 2005) was only recently purified and cloned. Surprisingly, sAC is closely related to

cyanobacterial adenylyl cyclases but not to tmAC, possibly explaining the over 20-year

period from enzymatic characterization to cloning. Mammalian sAC is directly

activated by HCO3- in a pH-independent manner (Chen et al., 2000) and by Ca2+

(Jaiswal & Conti, 2003; Litvin et al., 2003). HCO3- stimulates sAC activity by

22

increasing its Vmax and relieving substrate inhibition, while Ca2+ decreases the enzyme’s

Km for its substrate, ATP-Mg2+ (Litvin et al., 2003).

In the C-terminal portion of sAC, a region not required for enzymatic activity, a

potential leucine zipper or coiled–coiled interaction domain of unknown significance

can be found. With the recent documentation of sAC’s presence within specific

microdomains of many cells (Zippin et al., 2003), this region could serve as a protein–

protein interaction domain, possibly tethering sAC to the cytoskeleton at these specific

locations and in close contact to its major effector PKA, anchored to the cytoskeleton by

AKAPs. Such microdomains where sAC can be found include mitochondria, centrioles,

mitotic spindles, mid-bodies, and nuclei, all of which contain cAMP targets (Zippin et

al., 2003).

These data suggest that cAMP signaling plays a fundamental role in many

biological systems and make a good case that cilia, or structures close to cilia such as

basal bodies, contain sAC as well (already shown to be present in centrioles) (Zippin et

al., 2003). Sperm motility and hyperactivation has long been known to require HCO3-

and cAMP; the process has recently been shown to depend on sAC activity (Esposito et

al., 2004; Luconi et al., 2005).

Whether sAC is expressed in ciliated HAECs is not known. Since airway cilia

and sperm flagella share both structural and functional similarities and the beating

of cilia and flagella are regulated by similar mechanisms at least with respect to

cAMP, it is possible that sAC also plays a role in regulating airway ciliary beating,

perhaps by maintaining a certain cAMP level in cilia necessary for Ca2+-related CBF

actions (as mentioned in paragraph 1.3.4). One objective of these studies was to test

the hypothesis that sAC is expressed in ciliated HAECs and co-localized to the cilia.

1.3.5.2 Cytosolic ions: role of HCO3- and H+ in ciliary activity

Intracellular bicarbonate concentration ([HCO3-]i), the regulator of sAC, is one of

the most important regulators of pHi as well. Therefore, when HCO3--related CBF

changes are being investigated, it is necessary to identify whether CBF changes are

caused by cAMP production (via sAC activation) or by pHi-alterations (independently

of sAC). However, it has not been clarified how pHi itself influences CBF in HAECs.

23

The few papers investigated the effect of extracellular pH changes (without measuring

pHi) reported no significant effect on CBF within a certain pH-range (Ingels et al.,

1991; Clary-Meinesz et al., 1998b). CBF, for example, was not significantly modified

when extracellular pH was varied between 7.5 and 10.5. Reversible and significantly

lower frequencies were observed below pH 7.0 for bronchi and below pH 5.0 for

bronchioles. Extreme pH values (> 11.0, < 3.0) caused ciliostasis within a few minutes

(Clary-Meinesz et al., 1998a). Similar results were found when exposing cell cultures to

SO2 making the bathing solutions extremely acid (Kienast et al., 1994). In addition,

although controversial, one study showed that human spermatozoa lacking outer dynein

arms, in contrast to normal spermatozoa, failed to show higher beat frequency during

mild alkalization, suggesting that outer dynein arms might be involved in the response

to changing pH, maybe independent of HCO3- (Keskes et al., 1998b). Thus, there is

some evidence that outer dynein arm activity, the ciliary frequency-sensitive location

(Brokaw & Kamiya, 1987), may be directly sensitive to pH. The kinase/phosphatase

system is also a potential target of pHi changes. The catalytic efficiency of PKA is

optimal at near neutral pH and is inhibited at acidic pH (Cox & Taylor, 1995); in

addition, acidic pH not only inhibits PKA but activates phosphatase to de-

phosphorylation PKA targets and vice versa (Reddy et al., 1998b).

Considering that no publication had ever reported the relationship of pHi to

CBF, these studies also aimed to clarify this issue.

1.4. : Regulation of catecholamine concentration at adrenergic

receptor site: role of catecholamine transporters

Adrenergic neurotransmitters and hormones have powerful physiologic effects,

therefore their concentration at the adrenergic synaptic cleft and in the bloodstream is

strictly regulated. Since catecholamines can be metabolized only intracellularly (the

metabolizing enzymes, catecholamine-O-methyltransferase (COMT) and monoamine

oxidase (MAO), reside intracellularly), and, as organic cations, cannot pass the

plasmamembrane freely, their inactivation is determined mainly by cellular uptake

processes via specialized transporters (Koepsell et al., 2003; Ciarimboli & Schlatter,

24

2005). The diverse group of these transport proteins can be divided into neuronal and

extraneuronal catecholamine transporters.

Neuronal uptake of norepinephrine (NE), also referred to as uptake-1 or reuptake,

takes place through an oligospecific catecholamine plasma membrane transporter, the

norepinephrine transporter (NET). Besides being expressed in neuronal tissue and

neuroendocrine cells of the adrenal medulla, NET is also present in some extraneuronal

tissue like placenta (Bottalico et al., 2004) and pulmonary capillary endothelial cells,

where its main function is to clear catecholamines from the circulation (Catravas &

Gillis, 1983; Bryan-Lluka et al., 1992; Westwood et al., 1996; Eisenhofer, 2001;

Koepsell et al., 2003).

Extraneuronal catecholamine uptake is mediated by the polyspecific electrogenic

organic cation transporters (OCT). Three isoforms of OCTs have been identified in

mammals including man: OCT-1 (Grundemann et al., 1994; Gorboulev et al., 1997),

OCT-2 (Okuda et al., 1996; Gorboulev et al., 1997) and the ‘classic’ uptake-2, the

corticosterone-sensitive extraneuronal monoamine transporter (EMT, also referred to as

OCT-3) (Grundemann et al., 1998; Kekuda et al., 1998; Verhaagh et al., 1999). The

different OCT isoforms have different organ distribution that may reflect their

specialized roles. In man, OCT-1 appears to be confined to the liver and intestine, OCT-

2 is mainly expressed in the kidney but also detectable in human placenta and in

cerebral neurons, whereas the classic EMT has a broad tissue distribution being

expressed in skeletal muscle, liver, placenta, kidney, heart, lung and brain (Gorboulev et

al., 1997; Eisenhofer, 2001; Koepsell et al., 2003). Most recently, all the three OCT

isoforms have been identified in human airway epithelium (Lips et al., 2005) where two

of them, OCT-1 and OCT-2 , seem to act also as ACh transporters, participating in the

local non-neuronal cholinergic system (see above in paragraph 1.2.3.2).

The role of catecholamine transporters in the inactivation of circulating

catecholamines and terminating the activation of postsynaptic receptors in the

adrenergic synaptic cleft has been widely investigated (Eisenhofer, 2001). For example,

Figure 1-3 demonstrates the mechanism of bronchial arterial vasoconstriction resulting

from the inhibition of extraneuronal catecholamine uptake by glucocorticosteroids (GS)

25

and consequent increase in extracellular catecholamine concentration (Kumar et al.,

2000; Horvath et al., 2001; Mendes et al., 2003; Horvath & Wanner, 2006).

Figure 1-3. Proposed mechanism of the acute vasoconstrictor effect of inhaled corticosteroids in the airway. Corticosteroids facilitate the noradrenergic (sympathetic) neuromuscular signal transmission by rapidly (within 5 min) inhibiting the extraneuronal monoamine transporter (EMT) in vascular smooth muscle cells (Horvath & Wanner, 2006).

However, little is known about the local cellular uptake and metabolism of

inhaled β2-adrenergic agonist drugs in airway epithelium, which might be a critical

determinant of the magnitude and duration of their ciliostimulating effect.

Thus, in these studies we tested the catecholamine transporter expression

profile of HAECs. Furthermore, pharmacological characteristics of NE transport as

well as its inhibition by budesonide and methylprednisolone was also studied in a

model of human bronchial arterial smooth muscle cells expressing OCT-1 and EMT.

26

2. AIMS

1. sAC is a recently cloned adenylyl cyclase exhibiting completely different

structural and functional features than tmACs, the only known adenylyl cyclases in

bronchial epithelial cells so far. In contrast to tmACs, sAC is insensitive to G-protein

activation, but can be stimulated by HCO3-, independently of pH. Considering that

[HCO3-]i and pHi strictly correlate, and neither has ever been tested for ciliary effects,

we aimed to clarify how isolated pHi-changes influence CBF, a key issue for further

investigations of [HCO3-]i mediated CBF-effects. For this purpose, we performed

simultaneous CBF and pHi measurements on cultured HAECs.

2. To analyze the signaling mechanism by which pHi regulates CBF, we used

specific inhibitors of cAMP- and Ca2+-dependent signaling pathways and also measured

[Ca2+]i. Furthermore, we measured CBF on basolaterally permeabilized cells under the

direct control of pHi.

3. We also tested whether sAC mRNA and protein is expressed in HAECs. In

order to determine the subcellular localization of sAC, we performed

immunofluorescence studies on human tissue sections and cultured HAECs. Our

preliminary physiological experiments aimed to investigate sAC’s function in HAECs.

4. Another set of experiments targeted catecholamine transporter proteins, which

mediate cellular catecholamine uptake. Inhibition of cellular catecholamine uptake by

GSs has been shown to enhance adrenergic effects in vascular smooth muscle cells by

increasing extracellular catecholamine concentration. To test the hypothesis that GSs

have a similar effect on HAECs and thus, could enhance ciliostimulatory effect of

adrenergic agonists, we investigated the expression profile of HAECs for catecholamine

transporter mRNAs.

5. Finally we examined the pharmacologic characteristics of cellular

catecholamine uptake, including the effect of GSs on it as well as the mechanism by

which GSs inhibit cellular catecholamine uptake.

27

3. MATERIALS AND METHODS

3.1. Chemicals

LHC basal medium, Trace elements 100x, Stock 4 100x, and Stock 11 100x were

purchased from Biosource International (Rockville, MD, USA); Ham's nutrient F-12

and gentamicin from Gibco BRL Laboratories (Grand Island, NY, USA); the

acetoxymethyl ester form of the pH-sensitive dye 2’7’-bis-(2-carboxyethyl)-5-(and-6)-

carboxyfluorescein (BCECF) and fura-2 from Molecular Probes (Eugene, OR, USA);

nigericin from Molecular Probes (Eugene, OR, USA) and Calbiochem (La Jolla, CA,

USA); thapsigargin from Calbiochem (La Jolla, CA, USA); cyclosporin A from Fluka

(Buchs, Switzerland); and okadaic acid from Research Biochemicals International

(Natick, MA, USA). All other reagents were from Sigma Chemicals (St. Louis, MO,

USA) unless otherwise noted.

3.2. Solutions

Table 3-1 lists the compositions of solutions used for pHi experiments. The free

Ca2+ and Mg2+ concentration of EGTA- and ATP-containing solutions was estimated

using WebMAXC Standard software by Chris Patton from Stanford University,

available at http://www.stanford.edu/~cpatton/webmaxcS.html (constants used:

CMC1002.TCM).

28

Table 3-1: Composition of solutions. All concentrations are given in mM unless otherwise indicated. *Approximate free [Mg2+] after chelation by EGTA is 0.9 mM; †approximate free [Ca2+] after chelation by ATP is 0.1 mM; ‡pH was adjusted with concentrated NaOH solution unless otherwise indicated. CPK, creatine phosphokinase; CrP, creatine phosphate disodium salt.

3.3. Cell isolation and culture techniques

Human tissue was obtained from organ donors whose lungs were rejected for

transplant through the Life Alliance Organ Recovery Agency of the University of

Miami. Institutional Review Board-approved consents for use of these tissues for

research were obtained by the Life Alliance Organ Recovery Agency.

3.3.1. Preparation of submerged tracheal epithelial cell cultures

Primary cultures of tracheal epithelial cells were prepared as previously described

(Salathe & Bookman, 1995), with minor modifications. The mucosa from freshly

obtained tracheas was dissected from the underlying cartilage under sterile conditions

and incubated in 0.05 % protease (Sigma, Type 14) in Dulbecco’s modified Eagle’s

medium (DMEM) overnight at 4°C. After protease treatment, epithelial cells were

29

released by vigorous shaking, harvested by centrifugation and then plated on collagen-

coated glass coverslips (human placental collagen, Sigma) at a density of 1.5 – 6 x 104

cells·cm-2. The culture medium consisted of 47.5% DMEM, 47.5% Ham's F-12 nutrient,

5% fetal bovine serum (FBS) + penicillin (10 U/ml) and streptomycin (10 µg/ml) for the

first two days after plating. Then, it was changed to growth factor medium (50 %

DMEM, 50 % Ham's F-12 medium supplemented with insulin (10 µg·ml-1), transferrin

(5 µg·ml-1), hydrocortisone (0.36 µg·ml-1), triiodothyronine (20 ng·ml-1), endothelial cell

growth supplement (7.5 µg·ml-1), penicillin (10 U·ml-1), and streptomycin (10 µg·ml-1).

The medium was exchanged every other day. Cells from these cultures were used for

measurements within 6 days after plating.

3.3.2. Preparation of air liquid interface cultures of tracheal epithelium

Human air-liquid interface (ALI) cultures were prepared according to published

methods (Adler et al., 1990; Bernacki et al., 1999; Nlend et al., 2002).

Tracheobronchial epithelial cells isolated as above were first plated on collagen-coated

plastic dishes and grown to confluence in bronchial epithelial growth medium (BEGM),

yielding undifferentiated airway epithelial cells. BEGM contained 50% DMEM and

50% LHC basal medium supplemented with insulin (5 µg·ml-1), hydrocortisone (72

ng·ml-1), epidermal growth factor (31.3 ng·ml-1), triiodothyronine (6.5 ng·ml-1),

transferrin (10 µg·ml-1), epinephrine (0.6 µg·ml-1), phosphorylethanolamine (70 ng·ml-

1), ethanolamine (31 ng·ml-1), bovine pituitary extract (1% vol/vol), bovine serum

albumin (0.5 mg·ml-1), CaCl2 (0.08 mM), trace elements 100x (1% vol/vol), stock 4

100x (1% vol/vol), stock 11 100x (1% vol/vol), retinoic acid (15 ng·ml-1), penicillin

(100 U·ml-1), streptomycin (100 µg·ml-1), gentamicin (10 µg·ml-1) and amphotericin

(0.25 µg·ml-1). Cells were passaged after enzyme dissociation with trypsin onto 24 mm

diameter, 3 µm pore-sized Transwell collagen-coated inserts (Corning Costar

Corporation, Cambridge, MA, USA). The ALI medium used for both the apical and

basolateral side of the cells was the same as BEGM except for the concentration of

epidermal growth factor (0.63 ng·ml-1), penicillin (10 U·ml-1), streptomycin (10 µg·ml-1)

and the absence of gentamicin and amphotericin. Cells were grown in an incubator at 37

°C in ambient air supplemented with 5 % CO2. The medium was exchanged every other

30

day. The apical medium was removed as soon as cells reached confluence (usually after

1 week). The ALI cultures were used for measurements after the cells fully re-

differentiated (about 4-8 weeks, Figure 3-1).

Figure 3-1. Human cells at ALI. Human cells grown and differentiated for 21 days at the ALI were fixed with formaldehyde/glutaraldehyde, dehydrated, and embedded in Spurr's resin. Thick section stained with toluidine blue. Differentiated, pseudostratified epithelium is seen with mainly ciliated and a few goblet cells. The support filter is visible below (bar = 10 µm).

3.3.3. Human bronchial arterial smooth muscle cell isolation

Major branches of bronchial arteries (approximately 0.5 to 1-mm diameter) from

the main bronchi of freshly obtained lungs were excised using a dissecting

stereomicroscope under sterile conditions. To confirm that the dissected structure was in

fact an artery, a small portion of each vessel was fixed in 4% formaldehyde in

phosphate-buffered saline (PBS), processed according to regular procedures for

histology, and stained with hematoxylin and eosin. The rest of the vessel was dissected

from adhering fat and connective tissue and opened longitudinally. Endothelial cells

were removed by scraping the inside surface. From this muscle preparation, strips were

cut transversely and immediately used for smooth muscle cell (SMC) isolation or RNA

extraction.

SMC isolation from human bronchial arterial smooth muscle was carried out

based on the method of Clapp and Gurney (Clapp & Gurney, 1991) with some

modifications. For isolation of SMCs, muscle strips were transferred to a constantly

oxygenated incubation solution (137 mM NaCl, 4.17 mM NaHCO3, 0.34 mM

NaH2PO4, 5.37 mM KCl, 0.44 mM KH2PO4, 7 mM glucose, 0.15 mM CaCl2, 2 mM

MgCl2, 10 mM HEPES, 0.02% bovine serum albumin (BSA), pH 7.4) containing 1.5

mg/ml papain and 2 mM DTT, and were incubated at 37°C for 30 min with shaking.

Then, the muscle strips were transferred to a constantly oxygenated incubation solution

containing 1.5 mg/ml collagenase type F and 1 mg/ml hyaluronidase type I-S, and were

incubated at 37°C an additional 20 min with shaking.

31

At the end of the digestion period, individual SMCs were obtained by gentle

titration followed by filtration through a 500 μm pore size sieve. Finally, cells were

collected by centrifugation at 1000 x g for 3 min and resuspended in fresh (enzyme-

free) incubation solution. The viability of freshly isolated SMCs after enzymatic

dispersion was always > 95% as tested by trypan blue exclusion. The SMC suspension

was deposited onto human placental collagen (type VI)-coated glass coverslips. Cells

were allowed to settle for 60 min at 37°C before NE uptake experiments.

3.3.4. Preparation of primary cultures of bronchial arterial SMCs

For immunochemical detection of a corticosterone binding site, bronchial arterial

SMCs were maintained for 3 days in DMEM supplemented with 10% FBS, 100 U/ml

penicillin, and 100 μg/ml streptomycin within a humidified atmosphere containing 5%

CO2 at 37°C. This was necessary since acutely dissociated cells did not sufficiently

adhere to the coverslips and were lost during the immunocytochemical procedure.

3.3.5. Caki-1 cell culture

To provide a positive control sample for RT-PCR optimization of EMT mRNA,

the human renal carcinoma-derived Caki-1 cell line was purchased from the American

Type Culture Collection (ATCC; Manassas, VA, USA). Caki-1 cells were cultured in

McCoy’s 5A medium (ATCC) supplemented with 10% FBS, 100 U/ml penicillin, and

100 μg/ml streptomycin within a humidified atmosphere containing 5% CO2 at 37°C.

Media were changed in every other day.

3.4. Selective permeabilization of the basolateral membrane of cells

grown at the ALI

The basolateral surface of the ALI culture of human bronchial epithelial cells was

exposed to the pore-forming agent Staphylococcus aureus alpha-toxin at 10,000 U·ml-1

concentration dissolved in solution 8 (Table 3-1) for 30 min at room temperature.

32

Solutions 8 and 9 were composed to reflect physiological intracellular K+, Na+ and Cl-

concentrations, however, we had to use 100 µM Ca2+ to avoid cell detachment (similar

to the experience of others (Ostedgaard et al., 1992)). For another set of experiments,

0.05% saponin was used basolaterally, as described in Results.

3.5. Measurement of CBF

Cells grown on coverslips in submerged culture were mounted at room

temperature onto the stage of a Nikon Eclipse E600FN (Nikon, Melville, NY, USA)

upright water-immersion lens microscope in an open (Warner Instrument RC-25F with

a working volume of 150 µl) or closed (Warner Instrument RC-21BR, working volume

of 260 µl) perfusion chamber. They were perfused constantly. Ciliated cells were

imaged with infrared differential interference contrast (DIC) optics with an optical gain

of 600x. For online CBF measurements, the light path was directed to a CCD video

camera (XC-7500 Sony) and a box of 3 x 3 pixels from the live, digitized, contrast-

enhanced video image was selected (where one pixel samples an area of 180 nm x 180

nm). The magnitude spectra from a fast Fourier transform (FFT) of each of the pixel's

intensity signals were computed online and displayed on the monitor for immediate

adjustments. The intensity signals were recorded and later used for analysis according to

published methods (Salathe & Bookman, 1999) using a sliding FFT window approach

(128 frames per FFT, sliding the FFT window through the data set by 100 frames at a

time), providing a frequency resolution of at least 0.23 Hz and a time resolution of ~3 s.

The individual FFT magnitude spectra were peak extracted for graphing (Salathe &

Bookman, 1999).

In order to measure CBF in cells grown at the air-liquid interface (but imaged

with the apical surface 'submerged'), we used a holding chamber for the Transwell

membranes allowing selective perfusion of the apical and basolateral sides of the cells.

Data acquisition and processing was identical to the one described for submerged

cultures.

33

3.6. Measurement of pHi

3.6.1. Fluorometric pHi measurements

Coverslips without a confluent monolayer (usually 1 day after plating) were

preferred for fluorescent measurement of pHi, because these ciliated cells could be more

easily calibrated for pHi using nigericin (see below). Coverslips were rinsed in solution

1 (Table 3-1) and loaded with 2.5 µM BCECF acetoxymethyl ester in solution 1 at 37˚C

for 15 - 30 min and again rinsed three times. For fluorescent measurements, a Lambda

DG4 excitation system (Sutter, Novato, CA, USA) was used with 10 nm wide excitation

filters centered on 495 and 440 nm (Chroma Technology Corp., Brattleboro, VT, USA).

'Ratio-tool' software from Isee Imaging (Raleigh, NC, USA) controlled the output of the

Lambda DG4. Ratiometric pH estimates were made by capturing the light (535 nm)

emitted from the cells for 200 ms through a 60x water immersion objective (Nikon Inc.)

and directing it to a cooled CCD camera (CoolSnap Hq, Photometrics, Tucson, AZ,

USA). Individual ciliated cells were identified as regions of interest (ROIs) and the

BCECF ratio of emission intensity after excitation at 495 and 440 nm was computed

within each ROI every 10 – 60 sec on a pixel-by-pixel basis (after background

fluorescence subtraction).

3.6.2. Calibration of pHi measurements

3.6.2.1 Preparation of the pH calibration curve

Nigericin was used to calibrate pHi measurements (Thomas et al., 1979). BCECF-

loaded cells were perfused with calibration solutions containing 15 µM nigericin and

130 mM KCl at pH 6.8, 7.2, 7.5, and 7.8, respectively (solutions 3, Table 3-1) while the

fluorescence ratio was measured. The ratio data were normalized to the ratio at pH 7.2

(normalized fluorescence ratio: NFR; Figure 3-2/A). Each ratio value was accepted

when the ratio reached a steady state level at a given calibration pH. Then, the

calibration curve was constructed by plotting the average NFR values from 47 cells (5

different organ donors) against the corresponding pH values. The correlation between

34

NFR and pH values was determined by linear regression analysis (r2 > 0.99, Figure

3-2/B) (Boyarsky et al., 1988; Osypiw et al., 1994; Paradiso, 1997; Evans et al., 2003).

3.6.2.2 pHi estimation

A one-point nigericin calibration was performed at the end of each experiment

exposing the cells to the calibration solution with a pH of 7.2 (Boyarsky et al., 1988).

NFR values were calculated by dividing the ratio data from the entire experiment by the

ratio obtained in calibration solution at pH 7.2. NFR values were transformed to pHi by

interpolation with the calibration plot.

Nigericin was delivered to the cells via separate tubing. After each experiment

with nigericin, the mounting chamber and the manifold were washed with 100% ethanol

and distilled water to avoid nigericin-induced changes in pHi in the following

experiments (Richmond & Vaughan-Jones, 1997; Bevensee et al., 1999).

Figure 3-2. Calibration of pHi measurements. (A) A record of normalized fluorescence ratio (NFR) from a single, submerged, BCECF-loaded ciliated cell. The cell was first perfused with standard Hepes-buffered solution (solution 1, Table 3-1) which was switched to solutions containing 130 mM K+ and 15 µM nigericin (solution 3, Table 3-1) buffered at the indicated pH. Each fluorescence ratio value (I495/I440) was divided by the ratio obtained at pH 7.2 (NFR). (B) pHi dependence of NFR. NFR data were obtained from 47 cells. Linear regression is shown with an r2 > 0.99.

35

3.6.3. Experimental procedures for pHi measurements

Three different methods were used to manipulate pHi: a) ammonium prepulse; b)

removal of CO2 from the extracellular medium; and c) permeabilization of the

basolateral membrane with the pore forming agent alpha-toxin (from Staphylococcus

aureus). For method a), bicarbonate-free solutions were used to avoid interference with

sAC. All coverslips and ALI cultures used in these experiments were washed with

bicarbonate free solution, mounted into an open chamber and exposed to ambient air for

at least 30 min before experiments to remove any remaining bicarbonate and CO2 from

the cells. Measurements in open chambers were performed during continuous perfusion

with the desired solution at a flow rate of 250 µl/min using a Harvard pump. After

changing from one solution to another, flow rate was increased to 1000 µl/min for one

minute to accelerate the full exchange of the bathing solution (NH4Cl prepulse was

applied for 2 minutes at 1000 µl/min). These changes in flow rates did not influence

CBF in control experiments, similar to our earlier findings (Lieb et al., 2002). For

method b), cells were loaded with BCECF in CO2/HCO3--buffered solution at 5%

ambient CO2 and then mounted into a closed chamber with delivery of all solutions via

a closed system. Cells were perfused at a constant flow rate of 1000 µl/min. Finally for

method c), ALI cultures were mounted into an open chamber with separate perfusion

ports for the apical and basolateral side, maintaining perfusion at 500 and 1000 µl/min

for the apical and basolateral side, respectively. Up to four cells per coverslip or ALI-

filter were measured in each experiment.

3.7. Measurement of [Ca2+]i

[Ca2+]i was measured with the same equipment used for pHi. Cells grown on

coverslips were loaded with 5 µM fura-2 acetoxymethyl ester in solution 1 (Table 3-1)

for 60 min at room temperature and were washed 3 times with solution 1. Cells were

excited through 10 nm wide excitation filters centered on 340 and 380 nm (Chroma

Technology Corp., Brattleboro, VT, USA), and the emitted light was captured at 535

nm. Fura-2 ratios were computed every 10-20 s, after background subtraction.

Calibration of the calcium signal was done with in vitro measurements as described

36

(Salathe & Bookman, 1995) according to Grynkiewicz (1985); however, we re-

estimated [Ca2+]i by adjusting fura-2’s Kd with changing pH (Lattanzio & Bartschat,

1991; Browning & Wilkins, 2002) as described below.

3.8. Simultaneous measurement of CBF and pHi or [Ca2+]i

By using a dual-image module and guiding the infrared signal for CBF

measurements to the XC-7500 Sony CCD camera while sending all fluorescence signals

(< 580 nm) to the cooled CCD camera, we were able to measure recordings of CBF and

fluorescence (i.e., pHi or [Ca2+]i) of the same single cell simultaneously.

3.9. mRNA-expression of sAC and catecholamine transporters

3.9.1. RT-PCR

Total RNA was extracted from human airway epithelial cells grown and

redifferentiated at the ALI, cultured Caki-1 cells and freshly isolated bronchial arterial

smooth muscle cells using the RNeasy Protect Mini Kit (Qiagen, Valencia, CA, USA).

Total RNA samples from human testis, brain, liver, and kidney were purchased from

Ambion (Austin, TX, USA). Extracted RNA samples were treated with DNase (DNase I

Amplification Grade; Life Technologies), precipitated with ethanol, and quantified

spectrophotometrically at 260 nm. Good quality of isolated RNA (28S to 18S rRNA

ratio > 1.75) was confirmed using an RNA 6000 LabChip Kit (Agilent Technologies,

Palo Alto, CA) and an Agilent 2100 Bioanalyzer (Agilent Technologies) provided by

the University of Miami DNA Microarray Facility. RNA (1 μg per sample) was used for

first strand cDNA synthesis with Superscript II RT (Life Technologies) using oligo-dT16

primers. For PCR amplification, oligonucleotide primers were designed based on the

published sequences of human sAC (GenBank accession number: NM_018417), NET

(GenBank accession number: NM_001043), OCT-1 (GenBank accession number:

NM_003057), OCT-2 (GenBank accession number: NM_003058), EMT (GenBank

accession number: NM_021977), and GAPDH (GenBank accession number:

37

NM_002046) cDNAs. PCR reactions were performed using Taq DNA polymerase (Life

Technologies). Optimized annealing temperatures and cycle numbers for each primer

pair are shown in Table 3-2.

Table 3-2: Oligonucleotide primers, annealing temperatures (Tm), cycle numbers, and product sizes for RT-PCR amplifications of sAC, catecholqmine transporter and GAPDH mRNAs.

Forward primer Reverse primer mRNA

Sequence Position Sequence Position Tm Cycles Size

sAC ctgagcagttggtggagatcctc 388-410 cagccagtcctatcttgactcgg 630-608 61°C 40 243 bp NET ggccacggtatggattgatgc 961-981 cctcccattgagctgtcaagg 1317-1297 60°C 36 357 bp

OCT-1 ggctggctacaccctaatcacag 759-781 agtccgtgaaccacaggtacatc 1177-1155 60°C 30 419 bp OCT-2 gtacaactggttcacgagctctg 1226-1248 cgccaagattcctaatgaatgtggg 1569-1545 60°C 30 344 bp EMT ctgggtggtccctgagtctcc 882-902 tcccaggcgcatgacaagtcc 1146-1126 61°C 26 265 bp

GAPDH cctgcaccaccaactgcttag 527-547 gcctgcttcaccaccttcttg 869-849 60°C 24 343 bp

RT-PCR products were electrophoresed on ethidium bromide-stained 2% or 3%

SeaKem agarose (BMA, Rockland, ME, USA) gels. Control reactions were performed

in the absence of RT to verify that the amplified products were from mRNA and not

from genomic DNA contamination. In the absence of RT, no PCR products were

observed. To confirm specific amplification, RT-PCR products were purified on a silica

spin columns (QiAquick PCR Purification Kit, Qiagen, Valencia, CA, USA), and

sequenced by the University of Miami DNA Core Laboratory. Sequences were

compared with the published cDNA sequences by PileUp (Wisconsin Package, GCG,

Madison, WI, USA).

3.9.2. Cloning of cDNA fragments of sAC variants

The sAC specific amplification resulted in multiple bands which were gel-purified

and ligated into pGEM-T Easy vector (Promega, Madison, WI, USA). Recombinant

plasmid vectors were introduced into DH5α E. coli (Invitrogen, Carlsbad, CA, USA).

DNA of the transformed cells was purified with Wizard Plus SV Minipreps Purification

System (Promega, Madison, WI, USA), restriction enzyme digested with EcoRI (New

England BioLabs, Ipswich, MA, USA), electrophoresed on 3% NuSieve GTG agarose

38

gel (BMA, Rockland, ME, USA) and sequenced by the University of Miami DNA Core

Laboratory.

3.10. Expression of sAC-protein

3.10.1. Western blotting for sAC

Protein extracts from ALI cultures of HAECs were obtained by lysis at 100 ◦C in

1% sodium dodecyl sulphate (SDS), 10 mM Tris pH 8.5, 0.1 M EDTA, followed by

sonication, and centrifugation 14,000 g, as described elsewhere (Forteza et al., 2005).

Protein extract was sent to the Department of Pharmacology of Joan and Sanford I.

Weill Medical College of Cornell University (New York, NY, USA), where

immunoblotting using sAC-specific monoclonal antibodies was kindly performed by L.

R. Levin and coworkers.

3.10.2. sAC-specific immunohystochemistry and immunocytochemistry

Human tracheal rings were immersed in 4% paraformaldehyde, embedded in

paraffin, cut into 3-µm sections, and processed by indirect immunofluorescence. After

deparaffinization, autofluorescence was reduced by incubating slides in 0.1% sodium

borohydride in PBS. Slides were subjected to antigen retrieval by incubating them in 10

mM citrate buffer (pH 6) for 15 min, were incubated in 0.3% H2O2 to quench

endogenous peroxidase activity and incubated with sAC-specific monoclonal mouse

antibodies (kindly provided by L. R. Levin and J. Buck, Cornell University, New York,

NY, USA) for 18 h at 4 C. Labeling was achieved using a horseradish peroxidase (HRP)

conjugated secondary goat anti-mouse antibody and developed with TSA 546

AlexaFluor (Molecular Probes, Eugene, OR, USA) according to the manufacturers

instructions. For visualizing the axonemes, acetylated anti-tubulin antibody (Molecular

Probes, Eugene, OR, USA) was used. A nonspecific monoclonal antibody served as the

nonimmune control. The emitted light was filtered and recorded with a cooled black and

white CCD camera (Quantix, Photometrics, Tucson, AZ). Fully differentiated human

airway epithelial cells grown at the ALI culture were fixed with 50% methanol/ 50%

39

acetone and underwent the same procedures as the tracheal tissue. Nuclei were

visualized with 4,6-diamidino-2-phenylindole (DAPI, Molecular Probes, Eugene, OR,

USA). Stained ALI cultures were imaged with confocal microscope. All

immunofluorescence pictures were pseudo-colored using Photoshop (Adobe).

3.11. Single-cell, fluorometric NE uptake measurements

3.11.1. NE uptake experiments

For NE uptake studies, human bronchial arterial SMCs were exposed to

incubation solution containing NE with or without different NE transporter inhibitors

(see below) at 37°C. To inhibit the intracellular NE-metabolizing enzymes, 500 μM

pargyline (MAO inhibitor) (Henseling, 1983) and 1 μM Ro-41-0960 (COMT inhibitor)

(Percy et al., 1999) were added into the incubation solution 30 min prior to the NE

uptake experiments. GSs, such as corticosterone, budesonide, and methylprednisolone,

were dissolved in ethanol and freshly diluted into the incubation solution just prior to

use. The final concentration of ethanol was 0.1%, a concentration with no significant

effect on NE uptake measurements as confirmed in control experiments using this

vehicle only.

3.11.2. NE uptake assay

At the end of the incubation period, SMCs were washed with ice-cold incubation

solution. Intracellular NE was visualized using a sucrose-potassium phosphate-

glyoxylic acid (SPG) method described for tissue slices (Torre & Surgeon, 1976; de la

Torre, 1980) and adapted by us for use in isolated vascular SMCs. Briefly, coverslips

with SMCs were washed with SPG solution (0.2 M sucrose, 236 mM KH2PO4, 1%

glyoxylic acid monohydrate, pH 7.4) at room temperature. After 5 min of air-drying, the

specimen was covered with a drop of light mineral oil. Then, the sample was sealed

with a coverslip and put into an oven at 95°C for 2.5 min.

40

To quantify fluorescence, a Nikon Eclipse E600FN microscope (Nikon, Melville,

NY, USA) with a Lambda DG-4 excitation system (Sutter Instruments, Novato, CA,

USA), a cooled CCD camera (Coolsnap HQ; Roper Scientific, Inc., NJ, USA), and Isee

software from ISee Imaging Systems (Raleigh, NC, USA) were used. Cells were

imaged at 600x with differential-interference-contrast (DIC) microscopy and individual

cells were identified as regions of interest. For quantification of the SPG fluorescence

(or intracellular NE concentration) in these cells (or regions of interest), a 10 nm wide

filter centered on 405 nm was used for excitation and the emission measured at > 455

nm using a long pass filter (emission maximum: 480 nm), integrating the signal for 1

second. The cooled CCD camera was always set to a predefined gain, which was held

constant throughout the experiments. SMCs were measured by selecting 5 well-

separated regions on each coverslip (5-10 cells per region).

Each single cell’s mean fluorescence intensity value (Fn), expressed in arbitrary

units, was corrected for background fluorescence by subtracting the mean Fn of SMCs

from the same tissue that had not been exposed to NE. Average NE uptake of each

experimental group was calculated using the mean normalized Fn of all cells. Since we

have shown that the intracellular fluorescence is nearly linear to the intracellular NE

concentration over a wide concentration range (see below), we chose to report here

mainly the fluorescence in arbitrary unit and only convert the values to NE

concentration for the Km determination of NE uptake.

3.12. Immunochemical detection of a plasma membrane binding site

for corticosterone in bronchial arterial SMC

Human bronchial arterial SMCs maintained on collagen-coated coverslips for 3

days were washed three times with PBS and incubated with 1 μM BSA, 1 μM

corticosterone-21-hemisuccinate-BSA, or 1 μM corticosterone-21-hemisuccinate-BSA

plus 100 μM corticosterone for 5 min at 37°C. Then the cells were fixed with 4%

paraformaldehyde buffered with PBS for 30 min at room temperature. Fixed cells were

washed with PBS containing 200 μg/ml goat IgG (blocking buffer) and incubated with

blocking buffer for 60 min at room temperature. After washing with blocking buffer,

41

cells were incubated with rabbit anti-BSA IgG primary antibody (10 μg/ml) (Molecular

Probes, Eugene, OR, USA) for 60 min at room temperature. After washing again with

blocking buffer, the cells were incubated with a tetramethylrhodamine isothiocyanate-

(TRITC) conjugated goat anti-rabbit IgG secondary antibody (20 μg/ml) for 60 min at

room temperature. After washing with PBS and mounting in permanent aqueous

mounting medium (Gel/Mount; Biomedia, Foster City, CA, USA), cells were visualized

on the Nikon Eclipse E600FN microscope described above and TRITC fluorescence

imaged using an appropriate excitation/emission filter set.

3.13. Statistical analysis

Two groups were compared using Student’s unpaired t test, while Tukey-Kramer

honestly significant difference test was used for comparison of more than two groups if

an analysis of variance indicated a significant difference between groups (JMP software,

SAS Institute Inc., Cary, NC, USA). A p < 0.05 was considered to be significant. Data

were expressed as mean ± SEM

Uptake experiments were carried out in triplicate with measurements on 25 to 50

cells each. For time-course analysis of NE uptake, the data were fit with nonlinear

regression methods using the Prism version 3.0a (GraphPad Software Inc., San Diego,

CA, USA) with the following equation: A(t) = kin/kout * (1 – e-kout * t), where A(t) is the

uptake of NE at the time t, kin and kout are the rate constants for inward and outward

transport, respectively, and t is incubation time. The variables for the fit were kin and

kout and, thus, were determined during the fitting procedure. Km was determined with

nonlinear regression fit (Prism) using the following equation: y = (Vmax * X)/(Km + x),

where y is uptake of x amount of NE. Again Vmax was a variable of the fitting procedure.

For IC50 calculations, the data were fit with a multisite inhibition model (Prism) using

the following equation: y = inhibitionmax + (inhibitionmin - inhibitionmax)/(1+10(logIC50 x

Hill slope)), where y is the effect of x amount of the inhibitor.

42

4. RESULTS

4.1. Regulation of human airway ciliary beat frequency by pHi

One of our basic aims was to investigate the expression and function of HCO3--

sensitive adenylyl cyclase, sAC in HAECs. As [HCO3-]i and pHi strictly correlate, first

we targeted to investigate how isolated pHi-changes influence CBF, a key issue for

further investigations of [HCO3-]i mediated CBF-effects.

4.1.1. The effect of changing pHi on CBF during ammonium prepulse

The ammonium prepulse technique was used to achieve a change in pHi of intact

cells without changing the pH of the bathing solution (pHo). Baseline CBF was 7.2 ± 0.2

Hz and baseline pH was 7.49 ± 0.02 (n = 63 cells from 7 different organ donors). When

the ammonium-free standard Hepes solution (solution 1, Table 3-1) was rapidly

switched to one containing 10 mM NH4Cl (10 mM NaCl was replaced by 10 mM

NH4Cl, solution 2, Table 3-1), CBF and pH increased at the same rate, considering the

time resolution of the measurements. CBF reached a maximum of 9.4 ± 0.2 Hz (31 ±

1% above baseline; p < 0.001) within 52 ± 7 s, whereas pH reached a maximum of 7.78

± 0.02 (p < 0.0001 compared to baseline) within 48 ± 6 s. Two minutes after the initial

change to ammonium chloride, the medium was switched back to the standard Hepes

solution. After removing extracellular NH4+, a rapid intracellular acidification took

place and CBF and pH rapidly decreased to 5.8 ± 0.2 Hz (19 ± 2% below original

baseline) and 7.24 ± 0.02, respectively. Figure 4-1/A and B shows two representative,

simultaneous recordings of CBF and pHi during and after ammonium-chloride

exposure. To better visualize the correlation between pHi and CBF, these two

parameters were plotted against each other in Figure 4-1/C-E.

43

Figure 4-1. CBF responses to changing pHi during ammonium prepulse. (A) Simultaneous CBF and pHi recording from a single, submerged, BCECF-loaded human tracheobronchial epithelial cell perfused with standard Hepes-buffered solution (solution 1, Table 3-1). Intracellular alkalization and acidification by transient exposure to 10 mM NH4Cl (solution 2, Table 3-1) result in pHi-coupled changes in CBF. During recovery from acid load, CBF again parallels pHi. (B) Traces from an experiment identical to panel A, but recorded from a different cell. Here, CBF fails to follow pHi during recovery from intracellular acid load. (C, D) Correlation between CBF and pHi plotted from the experiment shown in panel A and panel B, respectively. (E) Correlation between CBF and pHi plotted from 59 cells transiently exposed to 10 mM NH4Cl.

During the recovery from acid load, CBF either increased together with pHi

(Figure 4-1/A and 35 of 63 cells), or remained at a low level during the rest of the

experiment (Figure 4-1/B). In the 35 cells with recovering CBF, baseline CBF and pHi

were 7.5 ± 0.2 Hz and 7.45 ± 0.03, respectively, not significantly different from the

non-recovering 28 cells (6.9 ± 0.2 Hz and 7.53 ± 0.03, both p > 0.05). However, the

maximum pHi following ammonium load was significantly higher in the latter group

(7.84 ± 0.03) compared with the recovering cells (7.72 ± 0.03). The CBF peaks in both

groups after ammonium load were not significantly different (9.7 ± 0.3 Hz in the

recovering group versus 9.1 ± 0.3 Hz in the non-recovering group). During the

44

following acidification phase, pHi in non-recovering cells was found to be significantly

higher than in cells that recovered their CBF (7.27 ± 0.03 versus 7.20 ± 0.02, p < 0.05);

on the other hand, CBF at pHi nadir was significantly higher in the recovering group

(6.1 ± 0.2 Hz) compared with the non-recovering group (5.4 ± 0.2 Hz). Since the

decrease of CBF during intracellular acidification was not significantly different

between the two groups, it is possible that a too severe alkalization was responsible for

the failure of these cells to recover their CBF after an acid load. This notion was

supported by the fact that a lower pH could be reached in many other cells without any

ill effects on CBF. However, other factors could also play a role as in other groups of

cells, high pHi did not seem to influence the ability of CBF to recover after acidification

(see inhibitor experiments below).

4.1.2. The effect of changing pHi on CBF during removal of extracellular CO2

An additional way to change pHi without changing pHo was to remove

extracellular CO2 (Thomas, 1984; Willumsen & Boucher, 1992; Paradiso et al., 2003).

For this purpose, cells in closed chambers were continuously perfused with CO2/HCO3--

buffered solution, pHo = 7.4 (25 mM NaCl of the standard Hepes solution was replaced

with 25 mM NaHCO3 following equilibration with 5% CO2/95% O2, pH 7.4, no Hepes,

solution 7, Table 3-1). Baseline pHi in this group of cells was 7.24 ± 0.02 (n = 13 cells

from two donors), significantly lower than in cells bathed in Hepes-buffered, nominally

CO2/HCO3--free solution (7.49 ± 0.02, see above; p < 0.0001). Similarly, baseline CBF

was significantly lower in CO2/HCO3--buffered than in Hepes-buffered solution (6.1 ±

0.3 vs. 7.2 ± 0.2 Hz, p < 0.005). When the CO2/HCO3--buffered solution was switched

to a nominally CO2/HCO3--free buffer of the same pH (25 mM NaHCO3 was replaced

by equimolar Na-gluconate, buffered with 10 mM Hepes; solution 6, Table 3-1), a rapid

increase in pHi to 7.75 ± 0.03 was seen (p < 0.0001). When the CO2/HCO3--buffered

medium was reapplied and extracellular CO2 entered the cell forming carbonic acid, pHi

fell below the original baseline to 7.04 ± 0.02 (p < 0.0001 compared to both baseline

and peak pHi), followed by a slow pHi recovery. Similar to the ammonium prepulse

experiments, CBF followed the changes in pHi, increasing from a baseline of 6.1 ± 0.3

Hz to 10.3 ± 0.5 Hz (or 71 ± 6% above baseline; p < 0.0001) and falling back to 5.7 ±

45

0.3 Hz (7 ± 3% below original baseline; p < 0.0001 compared to the peak CBF, but not

significantly different from baseline). The time course of the intracellular alkalization

and the parallel CBF increase was again within the resolution of the pHi measurement

(10 s): pHi reached its maximum within 92 ± 6 sec and CBF within 106 ± 13 sec.

Figure 4-2/A shows a representative experiment.

Even though removal of external CO2 both increased CBF and pHi significantly

more than the alkalization phase of ammonium prepulse, the ratio of these two

parameters (ΔCBF/ΔpHi) did not differ between the two methods (8.4 ± 0.6 sec-1 (pH

unit)-1 versus 7.7 ± 0.4 sec-1 (pH unit)-1 in the CO2 and NH4Cl group, respectively; p >

0.05).

Figure 4-2. CBF responses to changing pHi during removal of external CO2. (A) Simultaneous CBF and pHi recording from a single, submerged, BCECF-loaded human tracheobronchial epithelial cell, mounted into a closed chamber and perfused with CO2/HCO3

--buffered solution (solution 7, Table 3-1) in exchange with CO2/HCO3

--free, Hepes-buffered solution (solution 6, Table 3-1). (B) Correlation between CBF and pHi plotted from the experiment shown in panel A. (C) Correlation between CBF and pHi plotted from 13 cells during and after removal of external CO2 identical to the experiment shown in panel A.

46

4.1.3. Inhibition and stimulation of PKA does not prevent the effect of pHi on CBF

Kinases, specifically cAMP-dependent kinase (PKA), are important regulators of

mammalian airway ciliary beating. Since the activity of kinase/phosphatase systems has

been reported to depend on pH (Cox & Taylor, 1995; Reddy et al., 1998a), H-7, a broad

based serine-threonine kinase inhibitor, was used to inhibit PKA. Cells were incubated

with 100 µM H-7 for at least 20 minutes (estimated Ki of H-7 for PKA, PKG and PKC

is 3.0 µM, 5.8 µM and 6.0 µM, respectively Hidaka et al., 1984) until a steady state

CBF was achieved. Baseline CBF and pHi in the H-7 group was 7.2 ± 0.3 Hz and 7.45 ±

0.08, respectively (n = 10 cells from one donor), not significantly different from date

and culture matched controls (6.4 ± 0.4 Hz, 7.56 ± 0.08 pH units, n = 6, both p > 0.05).

Following H-7 exposure, CBF invariably decreased but average CBF was not

significantly below original baseline after 20 minutes of H-7 exposure. Cells were

exposed to 10 mM NH4Cl (solution 2, Table 3-1) in the continuous presence of H-7.

During alkalization, pHi increased significantly to 7.82 ± 0.11 and 7.83 ± 0.07 in the H-

7 and control group, respectively (p > 0.05). CBF also increased significantly in both

groups: to 9.5 ± 0.4 Hz (or 49 ± 3% above H-7 baseline) in the H-7 group and to 8.7 ±

0.9 Hz (or 36 ± 6% above baseline) in the control group (p > 0.05). After removing

external ammonium chloride, pHi fell in both groups: to 7.30 ± 0.04 in H-7-exposed and

to 7.34 ± 0.06 in control cells (p > 0.05). Similarly, CBF decreased to the same extent in

the H-7 and control groups, falling to 6.1 ± 0.3 Hz (or 96 ± 2% of the H-7 baseline) and

to 5.8 ± 0.4 Hz (or to 91 ± 2% of the baseline; p >0.05), respectively. A representative

experiment is shown in Figure 4-3/A.

The efficacy of PKA inhibition by H-7 was confirmed in experiments using 1 µM

forskolin to stimulate CBF, where H-7 (100 µM) significantly attenuated forskolin-

induced stimulation of CBF. CBF increased by only 1.6 ± 0.2 Hz (or 31 ± 3 %) above

baseline in H-7 pre-treated cells compared to 4.8 ± 0.5 Hz (or 86 ± 11 %) in control

cells (all n = 6; p < 0.001 for both the absolute and percent values).

47

Figure 4-3. PKA is not involved in the pHi-mediated CBF changes. (A) Simultaneous CBF and pHi recording from a single, submerged, BCECF-loaded human tracheobronchial epithelial cell pre-treated and continuously perfused with 100 µM H-7 and transiently exposed to 10 mM NH4Cl. Neither the CBF nor the pHi responses differed from control cells (see Figure 4-1). (B) Simultaneous CBF and pHi recording from a single, submerged, BCECF-loaded human tracheobronchial epithelial cell continuously exposed to 10 µM forskolin followed by a transient exposure to 10 mM NH4Cl.

These data were confirmed with a more specific inhibitor of PKA, namely 10 µm

H-89 (Davies et al., 2000). The reported Ki values for PKA, CaM kinase II and PKC are

48 nM, 29.7mM, and 31.7mM, respectively. Cells were pre-incubated with H-89 for 20

min. As with H-7, H-89 did not influence the pH-mediated changes in CBF during

ammonium pre-pulse experiments (n = 4) compared with date- and culture-matched

control cells (n = 6). H-89 also inhibited the CBF increase upon exposure of cells to 10

µm forskolin (n = 7 for H-89/forskolin and n = 8 for forskolin control).

The possible role of PKA in mediating pHi-induced changes in CBF was also

tested by fully stimulating the enzyme with 10 µM forskolin prior to applying the

ammonium prepulse. Forskolin was continuously present during and after the

ammonium exposure. Baseline CBF and pHi in the forskolin group was 6.9 ± 0.4 Hz

48

and 7.61 ± 0.07, respectively (n = 15 cells from three donors), not significantly different

from the controls (6.4 ± 0.5 Hz, 7.52 ± 0.04 pH units, n = 14). Following forskolin

stimulation, CBF significantly increased by 42 ± 4% above baseline (up to 9.7 ± 0.5 Hz)

and pHi remained unchanged (7.63 ± 0.08). During the alkalizing phase of the

ammonium prepulse, pHi peaked at 7.99 ± 0.10 and 7.78 ± 0.03 in the forskolin and the

control group, respectively (p < 0.05 compared to the baseline in both groups, but not

significantly different from each other). In the forskolin group, peak CBF was 60 ± 6%

above the original baseline (absolute CBF value was 10.9 ± 0.6 Hz) and 13 ± 4% above

post-forskolin plateau. The percent change during alkalization of the control group cells

was 30 ± 3% above baseline (absolute CBF value: 8.3 ± 0.3 Hz, significantly lower than

peak CBF in the forskolin pre-treated cells, p < 0.001). These results show that even if

PKA is already stimulated, increasing pHi still further stimulates ciliary beating. During

acidification, pHi fell to 7.39 ± 0.07 in the forskolin pre-treated cells and to 7.26 ± 0.04

in the control cells (no significant difference between the two groups). As expected,

CBF in the forskolin stimulated cells was still 22 ± 4% above the original baseline

(absolute value: 8.4 ± 0.5 Hz) while the beating frequency in the control group was 32 ±

4% below baseline (absolute value: 4.3 ± 0.2 Hz; p < 0.0001; Figure 4-3/B).

4.1.4. Inhibition of protein phosphatases does not influence the effect of pHi on

CBF

To inhibit protein phosphatases, a combination of inhibitors was used: 10 µM

cyclosporin A (inhibitor of calcineurin, a type 2B protein phosphatase, at an IC50 = 5

nM, Cohen et al., 1989) and 1.5 µM okadaic acid (inhibitor of type 1 at an IC50 = 15 –

20 nM and type 2A protein phosphatase at an IC50 = 0.1 nM, Cohen et al., 1989).

Baseline CBF and pHi before application of the inhibitors was 6.7 ± 0.2 Hz and 7.46 ±

0.03, respectively (n = 11 cells from two donors), not significantly different from the

date and culture matched controls (7.0 ± 0.3 Hz, 7.57 ± 0.06 pH units, n = 10). Cells in

the phosphatase inhibitor group were perfused with the inhibitors for 17 min, resulting

in a statistically insignificant CBF decrease (6.0 ± 0.3 Hz) and unchanged pHi (7.47 ±

0.02). During the alkalization phase of the ammonium prepulse, both pHi and CBF

49

increased to the same level in both groups, significantly above baseline. Maximum pHi

was 7.86 ± 0.01 in inhibitor treated cells and 7.89 ± 0.06 in control cells (p > 0.05),

while maximum CBF was 8.3 ± 0.3 Hz (40 ± 4% above post-inhibitor plateau

frequency) in the inhibitor group and 9.4 ± 0.5 Hz (or 34 ± 3% above baseline) in the

control group (p > 0.05 for both percent and absolute changes). The nadir of pHi and

CBF (in percent decrease from baseline) during acidification did not differ between the

treated and control groups: pHi was 7.20 ± 0.03 and 7.27 ± 0.06, while CBF fell to 90 ±

3% of the CBF plateau reached after adding inhibitors and to 89 ± 3% of the baseline in

the two groups, respectively. A representative experiment is shown in Figure 4-4.

Figure 4-4. CBF responses to changing pHi during ammonium prepulse: lack of effect by phosphatase inhibition. Simultaneous CBF and pHi recording from a single, submerged, BCECF-loaded human tracheobronchial epithelial cell perfused with 10 µM cyclosporine A plus 1.5 µM okadaic acid and transiently exposed to 10 mM NH4Cl. Cyclosporine A plus okadaic acid did not change the CBF or pHi responses to NH4Cl compared to control cells.

4.1.5. pHi does not regulate CBF via changes in [Ca2+]i

[Ca2+]i plays a crucial role in regulating CBF (e.g., Salathe & Bookman, 1995;

Salathe et al., 1997; Salathe & Bookman, 1999; Ma et al., 2002; Zagoory et al., 2002).

It has been shown in several cell types that intracellular alkalization can increase, while

intracellular acidification can decrease [Ca2+]i via regulation of Ca2+-release from

intracellular stores (Thomas et al., 1979; Browning & Wilkins, 2002). We investigated

whether CBF regulation by pHi depends on [Ca2+]i.

First, [Ca2+]i and CBF from the same single cells were measured simultaneously

during ammonium prepulse. The estimated baseline [Ca2+]i of cells perfused with

standard Hepes-buffered solution (solution 1, Table 3-1) was 47.5 ± 10.3 nM, baseline

50

CBF was 8.0 ± 0.3 Hz (n = 18 from one donor). Two minutes after starting the

ammonium load (solution 2, Table 3-1), CBF increased as expected and significantly to

10.9 ± 0.4 Hz (37 ± 3% above the baseline). Estimated [Ca2+]i however, did not change

significantly (46.1 ± 7.6 nM). After the ammonium prepulse was completed and CBF

returned to baseline, P2Y receptors were stimulated with 10 µM ATP for one minute.

Exposure to ATP resulted in a significant increase in both estimated [Ca2+]i (peak: 234.3

± 36.4 nM) and CBF (peak: 12.6 ± 0.3 Hz or 61 ± 7% above baseline; see examples in

Figure 4-5/A and B). To estimate [Ca2+]i, constant Kd (400 nM, Browning & Wilkins,

2002), Rmax, Rmin, and β values were used according to Grynkiewicz (1985) using in

vitro calibration procedures as outlined in Materials and methods. However, the affinity

of fura-2 for Ca2+ has been reported to increase with increasing pH (i.e. Kd decreases at

alkaline pH). In addition, Rmax as well as β were found to be pH-dependent (Lattanzio &

Bartschat, 1991; Browning & Wilkins, 2002). Therefore, Rmax, Rmin, and β were

measured in vitro in the pH range of 7.0 – 8.0. The changes in Rmin and β were

negligible. The average maximum intracellular alkalization during ammonium prepulse

in our hands was ~ 0.30 pH units (from 7.50 to 7.80, see above). Taken our in vitro

estimations and an approximate 12.5% decrease in Kd (400 nM to 350 nM) based on the

study by Browning and Wilkins (2002), these changes would result in a 17% decreased

estimated [Ca2+]i. Thus, increases in [Ca2+]i are over-estimated and decreases are under-

estimated by the ratio data upon alkalization and therefore [Ca2+]i changes did not

explain the CBF increase upon rising pH.

51

Figure 4-5. pHi-mediated changes in CBF are not due to [Ca2+]i variations. (A) Simultaneous CBF and [Ca2+]i recording from a single, submerged, fura-2-loaded human tracheobronchial epithelial cell, transiently exposed to 10 mM NH4Cl. Ammonium pre-pulse had a negligible effect on estimated [Ca2+]i, even if corrected for the pH-related changes in Kd after NH4Cl exposure. Purinergic stimulation with 10 µM ATP, however, resulted in a transient increase of both estimated [Ca2+]i and CBF. (B) Experiment shown in panel A was repeated in cells loaded with BCECF instead of fura-2. NH4Cl exposure caused a significant change in pHi, while ATP did not. (C) Simultaneous CBF and pHi recording from a single, submerged, BCECF-loaded human tracheobronchial epithelial cell, transiently exposed to 1 µM thapsigargin and nominally calcium free solution (solution 4, Table 3-1) before transient exposure to 10 mM NH4Cl. pHi-mediated CBF changes were not different from control cells.

To further show that Ca2+ was not involved in the pHi-mediated changes in CBF,

intracellular Ca2+-stores were emptied with 1 µM thapsigargin, an inhibitor of the

endoplasmic reticulum Ca2+-ATPase (Thastrup et al., 1990). Upon thapsigargin

exposure, baseline CBF of 6.4 ± 0.6 Hz (n = 8 cells from three donors) quickly

increased and then declined until reaching a plateau at 8.2 ± 0.5 Hz or 34.5 ± 16.3%

above baseline (p > 0.05; Figure 4-5/C). Three minutes before an ammonium load,

extracellular Ca2+ was removed (solution 4, Table 3-1) and CBF decreased to 6.8 ± 0.6

Hz. Longer Ca2+-free medium exposure resulted in cell detachment. After 3 minutes in

calcium-free, EGTA-buffered solution (nominal 0 [Ca2+]o), cells were exposed to 10

mM NH4Cl (solution 5, Table 3-1) containing 1 µM thapsigargin, resulting in a 28 ± 3%

52

acceleration of ciliary beating above pre-NH4Cl baseline (8.6 ± 0.7 Hz), a value not

significantly different from the 39 ± 5% increase from baseline of 6.3 ± 0.5 Hz to 8.7 ±

0.6 Hz seen in date-matched control cells (n = 5) neither exposed to thapsigargin or

calcium-free extracellular solution. Intracellular acid load after NH4Cl removal

significantly decreased CBF in both groups. Here, however, CBF of cells in nominally

free calcium solutions decreased significantly more than CBF of control cells: CBF

decreased 22 ± 3% below pre-ammonium challenge baseline to 5.3 ± 0.5 Hz vs. 5 ± 2%

to 6.0 ± 0.5 Hz, respectively (p < 0.05 for percentage values; p > 0.05 for the absolute

values). Baseline pHi was 7.38 ± 0.04 before ammonium exposure in cells bathed in

nominally free calcium solutions vs. 7.48 ± 0.04 baseline in control cells (p > 0.05). pHi

peaks were the same in both groups: 7.77 ± 0.04 in nominally free calcium solutions vs.

7.77 ± 0.05 in controls (p > 0.05); pHi nadir was 7.20 ± 0.04 vs. 7.27 ± 0.1, respectively

(p > 0.05).

4.1.6. Basolaterally permeabilized cells grown at the ALI

To control the cytoplasmic environment and to test whether the changes in CBF

seen in intact cells were directly related to changes in pH, the basolateral membrane of

human tracheobronchial epithelial cells re-differentiated at the ALI was selectively

permeabilized with Staphylococcus aureus alpha-toxin as described in Materials and

methods. Staphylococcus aureus alpha-toxin makes the membrane permeable to

molecules smaller than 5 kD, including nucleotides (Bhakdi & Tranum-Jensen, 1991;

Ostedgaard et al., 1992; Walev et al., 1993; Jonas et al., 1994; Reddy & Quinton, 1994;

Bhakdi et al., 1996; Detimary et al., 1996; Watanabe & Takano-Ohmuro, 2002). To

evaluate the efficacy of membrane permeabilization, the basolateral (i.e., intracellular)

solution containing 5 mM MgATP and an ATP regenerating system (ARS; 50 U·ml-1

creatine phosphokinase, 10 mM creatine phosphate disodium salt, solution 8 at pH 7.2,

Table 3-1) (Kakuta et al., 1985) was exchanged by a solution without ATP and ARS

(solution 9, Table 3-1). Seven minutes after the removal of this system, CBF fell from a

baseline of 6.7 ± 0.5 Hz by 4.2 ± 0.4 Hz (or by 62 ± 3 %; p < 0.0001 for both absolute

and percentage changes, n = 10 cells from two donors). These results suggested that the

basolateral membrane was permeable to ATP, even though the cilia were able to

53

maintain a low beating frequency of 2.5 ± 0.2 Hz (Figure 4-6/A). CBF returned to the

original baseline when the ATP/ARS-containing basolateral solution was reapplied.

Baseline CBF of non-permeabilized cells was 11.6 ± 0.6 Hz (significantly higher than

that of permeabilized cells; p < 0.0001) and did not change upon removal of ATP/ARS

from the basolateral solution (10.4 ± 0.4 Hz, p > 0.05, n = 4; Figure 4-6/B).

When alpha-toxin was applied apically (in the same solution used for the

basolateral permeabilization; solution 8, Table 3-1) while the basolateral membrane was

perfused with the standard Hepes-buffered solution (solution 1, Table 3-1), removal of

apical ATP/ARS (by changing to solution 9, Table 3-1) did not change CBF

significantly. These results show that alpha toxin permeabilizes specifically the

basolateral membrane as previously reported in an epithelial cell line (Ostedgaard et al.,

1992).

Figure 4-6. Basolateral permeabilization of human tracheobronchial epithelial cells re-differentiated at the air–liquid interface with alpha-toxin. (A) The basolateral surface of human airway epithelial cells grown and re-differentiated on 3 µm pore-sized inserts was permeabilized as described in Materials and methods. Removal of ATP and the ATP-regenerating system (50 U ml.1 creatine phosphokinase, 10 mM creatine phosphate) caused a reversible CBF decrease. (B) When re-differentiated cells were not permeabilized, removal of ATP and the ATP-regenerating system did not change CBF significantly.

4.1.7. Correlation between pHi and CBF in permeabilized cells

After selective permeabilization of the basolateral membrane, the basal

compartment of the ALI chamber was perfused with solutions titrated to pH 6.8, 7.2, 7.6

and 8.0 (solution 8, Table 3-1; Figure 4-7). CBF was 3.9 ± 0.3 Hz, 5.7 ± 0.4 Hz, 7.0 ±

54

0.3 Hz, and 7.3 ± 0.3 Hz at pH 6.8, 7.2, 7.6, and 8.0, respectively (n = 18, each from

two different organ donors; Fig. 8B). All these values were significantly different from

each other except the ones measured at pH 7.6 and 8.0, suggesting that intracellular

alkalization had a ceiling effect on stimulating CBF in permeabilized cells.

Figure 4-7. Correlation between pHi and CBF in basolaterally permeabilized human tracheobronchial epithelial cells. (A) Representative measurement from a single re-differentiated cell permeabilized with alpha-toxin and basolaterally exposed to different solutions equilibrated at the shown pH. Increasing basolateral (i.e. intracellular) pH resulted in a CBF increase. (B) Data obtained as in panel A, but summarized for 18 measured cells. Shown are the mean ± SEM.

4.2. sAC expression and cellular distribution in HAECs

4.2.1. sAC mRNA expression

RNA extracted from HAECs grown at the ALI as well as commercially purchased

human testicular, brain, liver and kidney RNA were reverse transcripted into cDNA and

amplified by PCR using sAC specific primers shown in Table 3-2. Gel-electrophoresis

of PCR amplification products of testicular, airway epithelial and cerebral RNA

suggested three bands in the expected size (243 bps) range (Figure 4-8/B). The medium

sized band seemed to be the predominant in the lane of HAEC sample. PCR

amplification of liver cDNA seemed to produce two bands, while PCR of kidney cDNA

generated only one band. Subsequent cloning of RT-PCR products from HAEC RNA

and gel-electrophoresis of the restriction enzyme digested vectors confirmed the

presence of cDNA of three different sizes. Sequencing of the vector inserts revealed a

55

243 bp fragment of human testicular sAC cDNA (V1 Figure 4-8/D) and two variants: a

published sAC-variant (Reed et al., 1999; Reed et al., 2002; Geng et al., 2005) having

37 extra 5’ nucleotides to exon 5 derived from the intron sequence (V2, Figure 4-8/D),

and an unpublished variant containing 49 extra 5’ nucleotides (V3 Figure 4-8/D).

Vector inserts of four different clones containing the medium sized sAC fragment were

sequenced. Two of them were 100% identical to the published sAC-variant (Geng et al.,

2005); the third one contained an A instead of G in exon 5 (shown in green in Figure

4-8/D), while the fourth one missed a T in exon 4 and contained an A instead of G in

exon 5 (shown in red in Figure 4-8/D).

56

Figure 4-8. mRNA expression of soluble adenylyl cyclase (sAC) in human airway epithelial cells. (A) Schematic view of the coding region of human sAC with the location of catalytic domain 1 and 2 (black boxes with C1 and C2, respectively) and the forward (f) and reverse (r) primers used to amplify sAC-transcript from human airway epithelial cells. (B) RNA was extracted from human airway epithelial cells grown at the ALI (HAEC). Human testicular, brain, liver and kidney RNAs were purchased. Amplification products of RT-PCR using sAC-specific primers were visualized by gel-electrophoresis. (C) Cloning of RT-PCR products from human airway epithelial RNA and subsequent gel-electrophoresis of the restriction enzyme digested vectors confirmed the presence of cDNA of three different sizes. (D) Sequencing of the vector inserts revealed a 243 bps fragment of human sAC cDNA (V1) and two variants (V2 and V3). Boldfaced fragments represent the primers used for PCR-reaction, the extra 37 nucleotides in V2 and V3 are underlined and the insert of an additional 12 nucleotides in V3 is double underlined. Red and green letters in V2 indicate variable bases (see text). Exons are indicated by colored lines.

Mar

ker

Test

is

HA

EC

Bra

in

Live

r

Kid

ney

No

RT

Mar

ker

Mar

ker

sAC

(V1)

Mar

ker

sAC

(V2)

sAC

(V3)

310 271, 281

234 194

A

B C

sAC f (Exon 3)

r (Exon 5-6) C2 C1

Exon 3 Exon 4 h

V1 CTGAGCAGTTGGTGGAGATCCTCAACTACCACATAAGTGCAATAGTGGAGAAAGTGTTGA 60

V2 CTGAGCAGTTGGTGGAGATCCTCAACTACCACATAAGTGCAATAGTGGAGAAAGTGTTGA

V3 CTGAGCAGTTGGTGGAGATCCTCAACTACCACATAAGTGCAATAGTGGAGAAAGTGTTGA

Exon 4 h

V1 TTTTTGGAGGAGACATCCTGAAATTTGCAG------------------------------ 120

V2 TTTTTGGAGGAGACATCCTGAAATTTGCAG------------GGCAGGTCAAAAGCCCGC

V3 TTTTTGGAGGAGACATCCTGAAATTTGCAGATTGATCCCCGGGGCAGGTCAAAAGCCCGC

Exon 5 h

V1 -------------------GTGATGCACTGCTAGCCCTGTGGAGGGTGGAGCGAAAGCAG 180

V2 GGCATGTCTCTCTCTGAAGGTGATGCACTGCTAGCCCTGTGGAGGGTGGAGCGAAAGCAG

V3 GGCATGTCTCTCTCTGAAGGTGATGCACTGCTAGCCCTGTGGAGGGTGGAGCGAAAGCAG

Exon 5 h

V1 CTGAAAAACATTATCACAGTGGTAATTAAATGTAGCCTGGAGATCCATGGATTGTTTGAG 240

V2 CTGAAAAACATTATCACAGTGGTAATTAAATGTAGCCTGGAGATCCATGGATTGTTTGAG

V3 CTGAAAAACATTATCACAGTGGTAATTAAATGTAGCCTGGAGATCCATGGATTGTTTGAG

Exon 5 Exon 6 h

V1 ACCCAGGAGTGGGAAGAAGGCCTAGACATCCGAGTCAAGATAGGACTGGCTG 292

V2 ACCCAGGAGTGGGAAGAAGGCCTAGACATCCGAGTCAAGATAGGACTGGCTG

V3 ACCCAGGAGTGGGAAGAAGGCCTAGACATCCGAGTCAAGATAGGACTGGCTG

D

57

4.2.2. Expression and cellular distribution of sAC protein

Western blot analysis of whole cell extract of HAECs grown at the ALI using

sAC-specific monoclonal antibodies revealed a band close to the 50 kDa marker,

matching the band of sAC standard protein (Figure 4-9). A faint band at the same size

was also detected from the ciliary extract of HAECs (not shown). A ~50 kDa sAC

protein containing the amino-terminal catalytic domains C1 and C2 is supposed to be

the product of posttranslational cleavage of the full length (187 kDa) sAC protein and

has been reported in many human cell types (Chen et al., 2000; Sun et al., 2004; Zippin

et al., 2004; Han et al., 2005; Wang et al., 2005).

Figure 4-9. Western blot analysis of sAC expression in human airway epithelial cells. Whole cell protein extract from human airway epithelial cells (HAEC) cultured at the air-liquid interface was immunoblotted using anti-sAC monoclonal antibodies (right lane). Specific staining was detected at the expected size of truncated sAC protein (~50 kDa). As a control, standard sAC protein (sAC STD), a protein extract from cells overexpressing the truncated sAC-isoform, was running in the left lane. The molecular mass of the protein standard is indicated. (Western blot analysis was kindly performed by L. R. Levin and coworkers at Cornell University, New York, NY, USA.)

To further confirm the expression of sAC protein in HAECs and to identify its

distribution within the airway epithelium, human tracheal sections were stained with

sAC specific monoclonal mouse antibody and a secondary anti-mouse IgG antibody

labeled with Alexa 546 (red) (Figure 4-10). For visualizing the ciliary axonemes,

acetylated anti-tubulin antibody was used (green). Fluorescence microscopy

demonstrated that sAC is expressed in the ciliary cells along with the cilia.

50 kDa

75 kDa

25 kDa

58

Figure 4-10. Immunolocalization of soluble adenylyl cyclase (sAC) in human trachea. Human tracheal tissue sections were fixed with 4% paraformaldehyde and stained with monoclonal antibodies against sAC (secondary label was Alexa 546 – red channel, panel B) and against acetylated tubulin (secondary label was fluorescein – green channel, panel C). Panel A: bright field illumination, panel D: merged channels. Bar is 10 µm. sAC is expressed at the apex of superficial airway epithelial cells co-localized with the cilia.

Immunofluorescence study of HAECs cultured at the ALI was also performed in

order to construct confocal images for better intracellular localization of sAC. Using the

same sAC specific monoclonal antibody applied for the tracheal tissue, confocal

microscopy confirmed that sAC is expressed in cultured ciliated cells as well (Figure

4-11). The primary anti-sAC mouse antibody was visualized with Alexa 546-labeled

anti-mouse IgG (red). Cilia were stained with acetylated anti-tubulin antibodies (green),

nuclei were imaged with DAPI (blue). The merged image clearly proves that sAC is

located on the ciliary shaft co-localizing with the axonemal tubulin and in the apical

membrane (merged images, Figure 4-11; green + red = yellow). Staining with a non-

specific mouse IgG (control for sAC) completely eliminated the red channel

fluorescence (not shown).

C

B

D

A

59

Figure 4-11. Immunolocalization of soluble adenylyl cyclase (sAC) in human airway epithelial cells cultured at the air-liquid interface. Fully differentiated human ALI cultures were fixed with 50% methanol/50% acetone and stained with antibodies against acetylated tubulin (AcTb; secondary label was fluorescein – green channel) and against sAC (secondary label was Alexa 546 – red channel). Finally, the nuclei were stained with DAPI (blue channel). The merged channels are shown as well. Three levels obtained with confocal microscopy are shown (ciliary level, apical membrane level below cilia and cellular or basolateral level) together with a z-axis reconstructed image (z stack). Bar is 10 µm (cells apparently flattened during mounting procedure). sAC is expressed apically and along the axoneme.

4.2.3. Effect of cytosolic HCO3- on ciliary beating in HAECs

To demonstrate that sAC expressed in HAECs is functioning and involved in

CBF-regulation, we performed preliminary experiments to test the effect of cytosolic

HCO3- on CBF. In order to overcome the difficulties of controlling [HCO3

-]i by

externally applied bicarbonate without changing pHi (see Figure 4-2/A), the basolateral

60

surface of a polarized HAEC culture was partially permeabilized by the detergent

saponin (Kakuta et al., 1985). For this purpose, ALI culture of HAECs was mounted in

a closed chamber allowing selective perfusion of the apical and basolateral membrane

of the cells. The basolateral membrane was perfused with an ATP containing buffer

(similar to solution 8 containing 10 mM ATP, pH 7.2, Table 3-1) followed by an ATP-

free solution (solution 9, Table 3-1), containing 0.05% saponin. The apical compartment

was perfused with HCO3--free Hepes buffer (similar to solution 1, Table 3-1, pH

adjusted to 7.2) and CBF of twelve ciliated cells was recorded simultaneously. CBF of

two representative cells is shown in Figure 4-12.

Figure 4-12. Effect of intracellular HCO3

- on CBF of intact and basolaterally permeabilized HAECs. Polarized culture of HAECs grown at the ALI was mounted in a closed chamber. Apical and basolateral surface of the culture was perfused separately as indicated by the bars and CBF was measured continuously. CBF of two representative cells of the same experiment are shown. The basolateral membrane of the cells was exposed to 0.05% saponin as indicated in the absence of ATP (“no ATP”). CBF of one cell (blue plot) started to decrease upon saponin treatment indicating ATP loss through the permeabilized membrane (full arrow). Saponin was then removed permanently in order to avoid cellular death. After re-applying basolateral ATP (as indicated by the end of “no ATP” bar), CBF of permeabilized cell suddenly increased while the intact cell (red plot) did not respond on basolateral ATP (empty arrows). Bilateral CO2/HCO3

- inhibited CBF of intact cell because of intracellular acidification, while stimulates ciliary beating of permeabilized cell possibly through activation of sAC (black triangles).

In order to avoid saponin-related toxicity, the basolateral saponin was temporarily

removed several times. As soon as CBF of at least one cell started to decrease indicating

cellular ATP-efflux because of membrane permeabilization (blue plot at the full arrow

in Figure 4-12), saponin was removed permanently from the basolateral perfusate. Re-

application of ATP to the basolateral membrane resulted in increasing CBF, further

61

confirming that attenuation of ciliary beating was the consequence of ATP loss. On the

contrary, those cells which remained intact (indicated by the absence of CBF change on

saponin) did not respond on basolateral ATP (red plot, Figure 4-12). Then, both the

apical and basolateral surfaces of the cells were exposed to CO2/HCO3--containing

buffers of the same pH (apical perfusate: solution 7 from Table 3-1, pH adjusted to 7.2;

basolateral perfusate: solution 8 from Table 3-1, 25 mM K-gluconate replaced by 25

mM KHCO3, equilibrated with 5% CO2, pH 7.2,). As expected based on the experiment

shown in Figure 4-2/A, intact cells responded on CO2/HCO3--containing buffers with

slowing ciliary beating because of intracellular acidification (CO2 freely crosses the

plasmamembrane and forms carbonic acid inside the cell, while HCO3- mainly remained

out of the cell). On the contrary, CO2/HCO3- augmented CBF of permeabilized cell in

the same culture, possibly via sAC-activation.

4.3. NE transporter mRNA expression profile in human airway

epithelial cells

Because non-neuronal cells may express various and/or multiple transport systems

for NE, RT-PCR reactions were initiated to examine the presence of neuronal (NET

(Pacholczyk et al., 1991)) and non-neuronal (OCT-1 (Grundemann et al., 1994), OCT-2

(Okuda et al., 1996), and EMT) transporter mRNAs in HAECs grown and

redifferentiated at the ALI, and freshly isolated bronchial arterial smooth muscle cells.

Based on the published expression data, human brain, liver, kidney, and Caki-1 cell line

mRNA samples were used to optimize the RT-PCR reactions for NET, OCT-1, OCT-2,

and EMT mRNAs, respectively. Gel electrophoresis of the RT-PCR products for these

samples showed bands of expected sizes (Figure 4-13). Gel-purification and sequence

analysis confirmed that these amplicons were fragments of the corresponding NE

transporter cDNAs. The isolated fragments contained only exon sequences (from at

least 3 different exons for each) confirming the amplification of mRNA rather than

genomic sequences (Grundemann & Schomig, 2000).

62

Figure 4-13. RT-PCR analysis of NE transporter mRNA expression. RT-PCR products using the primers and conditions listed in Table 3-2 were electrophoresed on ethidium bromide-stained 2% agarose gels. Gel-purification and DNA sequencing was used to confirm specific amplification of NET (357 bp), OCT-1 (419 bp), OCT-2 (344 bp), EMT (265 bp), and GAPDH (343 bp) cDNAs. Staring RNA was from human Brain, Liver, Kidney, Caki-1 cells, bronchial arterial smooth muscle (BASM), and airway epithelial cells (AEC) cultured at the air-liquid interface.

Both HAECs and human bronchial arterial smooth muscle cells expressed OCT-1

and EMT. Interestingly, HAECs also expressed NET mRNA, similarly to the kidney

RNA sample. Brain and kidney expressed mRNA for every neuronal and non-neuronal

catecholamine transporter, whereas Caki-1 cells expressed only mRNA for every non-

neuronal transporter (Figure 4-13).

4.4. Pharmacological characterization of NE uptake

4.4.1. NE uptake characteristics in bronchial arterial SMCs

To study the pharmacologic characteristics of OCT-1 and/or EMT mediated NE

transport and to determine whether it is blockable by GSs, we used human bronchial

arterial smooth muscle cells as a model, expressing both OCT-1 and EMT. Since rabbit

aortic arterial SMCs showed steroid-sensitive NE uptake in prior experiments (Horvath

et al., 2001) and since RT-PCR analysis indicated that EMT and OCT-1 mRNAs are

63

expressed in human bronchial arterial SMCs, NE uptake was measured in these cells in

the absence or presence of 10 μM corticosterone. This concentration was chosen to

inhibit EMT but not other OCT-1 (and OCT-2) in a significant amount (Martel et al.,

1999; Hayer-Zillgen et al., 2002); see also below. The amount of uptake blockable by

10 μM corticosterone was defined as EMT-mediated.

To show first that NE uptake into human bronchial arterial SMCs is a time-

dependent phenomenon, cells were incubated in 50 μM NE with and without 10 μM

corticosterone for 5, 15, 30, 45, and 90 min. NE uptake was detectable after 5 min and

increased in a time-dependent fashion for 45 min (Figure 4-14A). The concentration-

dependence was evaluated by incubating the cells in 25, 50, 250, and 1,000 μM NE with

and without 10 μM corticosterone for 5 min. EMT-mediated uptake seemed to saturate

towards 1,000 μM NE (Figure 4-14B). The calculated Km for NE uptake was

approximately 240 µM. This is close to the Km value of 245 µM calculated for NE

uptake into rabbit aortic SMCs (see above), a process likely mediated by EMT

(Schomig & Schonfeld, 1990).

Figure 4-14. Time- and concentration-dependence of NE uptake by human bronchial arterial SMCs. (A) Cells were exposed to 50 μM NE-containing incubation medium for different time periods. Shown is the EMT-mediated (solid circle) and not EMT-mediated (open circle) uptake of NE. EMT-mediated uptake was defined as the amount of uptake that was blockable by 10 μM corticosterone. (B) EMT-mediated NE uptake rates (v) were calculated from the 5-min time-point and plotted against NE concentrations. F is fluorescence in arbitrary units. Lines were fitted to the experimental data using nonlinear regression methods. Shown are means ± SEM for triplicate experiments with n of cells = 25-50 each.

64

The NE uptake inhibition by 10 µM corticosterone as well as the calculated Km

for NE uptake, suggested that EMT played the major part in cellular NE uptake. To

support this hypothesis, the pharmacological inhibitor profile of NE uptake was further

investigated. In these experiments, SMCs were exposed to 50 μM NE for 5 min without

or with 1 μM desipramine (inhibits only NET with a Ki of 4 nM (Buck & Amara,

1995)), 1 μM corticosterone (inhibits OCT-1, OCT-2, and EMT with the IC50 values of

21.7 μM, 34.2 μM, and 0.29 μM, respectively (Schomig & Schonfeld, 1990;

Grundemann et al., 1997; Martel et al., 1999)), or 1 μM O-methyl-isoprenaline (inhibits

OCT-2 and EMT with the IC50 values of 580 μM and 4.38 μM, respectively, but not

OCT-1 (Schomig & Schonfeld, 1990; Martel et al., 1999; Hayer-Zillgen et al., 2002)).

Control experiments showed that these inhibitors did not affect the fluorometric assay at

the concentrations used (data not shown in figure). Desipramine did not decrease NE

uptake significantly (the decrease was only 4.2 ± 5.7% in triplicate experiments with n

of cells = 25-50 each; p > 0.05 versus control), whereas corticosterone and O-methyl-

isoprenaline inhibited NE uptake by 63.2 ± 6.9% and 57.8 ± 2.9%, respectively (in

triplicate experiments with n of cells = 25-50 each; p < 0.05 versus control for both)

(Figure 4-15). Together with NE transport kinetics data (see above), these inhibitor

characteristics suggest that a significant amount of NE uptake is mediated by EMT.

Figure 4-15. The effects of NE transporter inhibitors on NE uptake by human bronchial arterial SMCs. Cells were exposed to 50 μM NE for 5 min without (control) or with 1 μM desipramine (DESI) to inhibit NET, 1 μM corticosterone (CORT) to inhibit EMT, or 1 μM O-methyl-isoprenaline (OMI) to inhibit EMT. F is fluorescence in arbitrary units. Shown are means ± SEM for triplicate experiments with n of cells = 25-50 each. *p < 0.05 versus control.

4.4.2. Inhibition of NE uptake by GSs

To test whether GSs used in the clinical practice are also able to inhibit cellular

NE uptake, we investigated the effects of budesonide and methylprednisolone. Similarly

65

to corticosterone, budesonide and methylprednisolone inhibited NE uptake after 5 min

of incubation (Figure 4-16). The rank order for inhibitory potency was corticosterone >

budesonide > methylprednisolone. The calculated apparent IC50 values for these

compounds were 0.12 μM (close to the reported IC50 of 0.29 µM for EMT in other

cells), 0.89 μM, and 5.56 μM, respectively.

Figure 4-16. Inhibition of NE uptake by GSs in human bronchial arterial SMCs. Cells were exposed to 50 μM NE for 5 min in the presence of corticosterone, budesonide, or methylprednisolone. Data are shown as a percentage of NE uptake measured in the absence of inhibitors. Shown are means ± SEM for triplicate experiments with n of cells = 25-50 for each. The calculated IC50 values for corticosterone, budesonide, and methyl-prednisolone were 0.12 μM, 0.89 μM, and 5.56 μM, respectively.

4.4.3. EMT inhibition by corticosterone is a nongenomic effect

Evidence have been provided from our laboratory that NE uptake into rabbit

aortic smooth muscle cells is mediated by a nongenomic action of GSs independent on

transcription, protein synthesis, and membrane penetration of the drug (Horvath et al.,

2001). Here we repeated these experiments with human bronchial arterial SMCs. In

addition, we also examined the acute reversibility of the corticosterone effect and its

dependence on the classical, intracellular GS receptor.

To investigate reversibility, human bronchial arterial SMCs were exposed to 1

μM corticosterone for 5 min, which blocked NE uptake by about 65% in our

experiments (see above). Then the cells were transferred into a corticosterone-free

medium containing 50 μM NE for uptake measurements. There was no significant

difference in NE uptake between SMCs pretreated with corticosterone and control cells

not exposed to corticosterone (17.1 ± 2.2 versus 21.1 ± 0.9 arbitrary units; in triplicate

66

experiments with n of cells = 25-50 each; p > 0.05) (Figure 4-17A). Thus, the inhibition

of NE uptake by corticosterone is reversible after removal of the GS.

Figure 4-17. Corticosterone inhibits NE uptake into human bronchial arterial SMCs via a nongenomic mechanism. (A) Cells were exposed to 50 μM NE for 5 min without (control) or with 1 μM corticosterone (CORT). Additionally, cells were pre-treated with 1 μM corticosterone for 5 min but exposed to NE in corticosterone-free solutions (CORT⇒NE). (B) Cells were exposed to 50 μM NE without (control) or with 10 μM RU486 (RU) plus 1 μM corticosterone, 100 μM actinomycin D (ACT) plus 1 μM corticosterone, or 10 μM cycloheximide (CYC) plus 1 μM corticosterone. (C) Cells were exposed to 50 μM NE without (control) or with 1 μM corticosterone-BSA conjugate (CORT:BSA). F is fluorescence in arbitrary units. Shown are means ± SEM for triplicate experiments with n of cells = 25-50 for each. *p < 0.05 versus control.

To show that classical genomic pathway (i.e., binding to cytoplasmic receptors

thereby initiating changes in transcription and protein synthesis of target genes) was not

involved in corticosterone's rapid action, either 10 μM RU486 (a cytoplasmic GS

receptor antagonist), 100 μM actinomycin D (transcription inhibitor), or 10 μM

cycloheximide (protein synthesis inhibitor) was added to the incubation medium 30 min

prior to the addition of 50 μM NE and 1 μM corticosterone. Control experiments

showed that either RU486, actinomycin D, or cycloheximide did not inhibit NE uptake

at the concentrations used (data not shown in figure). Corticosterone inhibited NE

uptake by 66.7 ± 12.4%, 62.1 ± 6.8%, and 69.4 ± 11.6% in the presence of RU486,

actinomycin D, or cycloheximide (in triplicate experiments with n of cells = 25-50 each;

p < 0.05 versus control for all, p > 0.05 versus corticosterone-treated) (Figure 4-17B).

Thus, the NE uptake inhibition by corticosterone does neither depend on the

cytoplasmic GS receptor nor on changes in transcription or translation.

67

To investigate whether corticosterone acts at the plasma membrane,

corticosterone was prevented from entering the cell by conjugation to the membrane-

impermeant carrier protein BSA (Zheng & Ramirez, 1997, 1999). Corticosterone-21-

hemisuccinate-BSA (1 µM) decreased NE uptake into SMCs by 57.7 ± 16.3% (in

triplicate experiments with n of cells = 25-50 each; p < 0.05 versus control, p > 0.05

versus corticosterone-treated) (Figure 4-17C). Since membrane-impermeant

corticosterone also inhibited NE uptake into human bronchial arterial SMCs, we

concluded that the acute effects of GSs are likely mediated through a GS binding site at

the plasma membrane.

Inasmuch as the activity of the membrane-impermeant corticosterone-BSA

conjugate suggested that corticosterone acted at a site located at or in the plasma

membrane, we looked for further evidence of a membrane-binding site for

corticosterone in human bronchial arterial SMC. To immunocytochemically visualize

the binding of membrane-impermeant corticosterone-BSA conjugate to the plasma

membrane, rabbit IgG antibodies against BSA were used in combination with TRITC

labeled goat anti-rabbit IgG secondary antibodies (Figure 4-18). Fluorescent labeling

was absent in control cells that were incubated in 1 μM BSA-containing medium for 5

min (Figure 4-18A and B), whereas cells incubated in 1 μM corticosterone-21-

hemisuccinate-BSA for 5 min were specifically labeled (Figure 4-18C and D).

Inasmuch as the conjugated corticosterone was necessary for BSA binding, this

experiment demonstrated a binding site for corticosterone in the cell membrane.

Competition with 100 µM corticosterone completely inhibited binding of

corticosterone-21-hemisuccinate-BSA to the cells (Figure 4-18E and F), indicating that

corticosterone binding to the plasma membrane was specific.

68

Figure 4-18. Immunochemical demonstration of a specific plasma membrane binding site for corticosterone in human bronchial arterial SMC. Cells were incubated with 1 μM BSA (A-B), 1 μM corticosterone-BSA (C-D), or 1 μM corticosterone-BSA plus 100 μM corticosterone (E-F). Rabbit anti-BSA primary and a tetramethylrhodamine isothiocyanate-labeled goat anti-rabbit IgG secondary antibody were used for immunocytochemistry. A, C and E: differential interference contrast microscopy; B, D and F: fluorescence (tetramethylrhodamine isothiocyanate) microscopy. Original magnifi-cation: x600.

69

5. DISCUSSION

5.1. Regulation of human airway ciliary beat frequency by pHi

Previous studies showed that extracellular alkaline solutions up to pH 9 – 10.5

had no effect on mammalian airway CBF, while extracellular acidic solutions attenuated

ciliary beating (van de Donk et al., 1980; Luk & Dulfano, 1983; Clary-Meinesz et al.,

1998a). However, the relationship between intracellular pH and CBF has been unclear.

Here we demonstrate that relatively small changes in pHi result in significant changes in

ciliary beating of human tracheobronchial epithelial cells. The observed changes in

ciliary beating are large enough to alter a more macroscopic physiological parameter,

i.e. mucociliary clearance: Seybold et al. (1990), for instance, found that a 16% increase

in CBF can lead to a 56% increase in mucociliary transport velocity in isolated whole

tracheas.

Our experiments suggest that changing pHi may directly act on the ciliary motile

machinery, possibly the outer dynein arm. The exact mechanism of how pHi changes

CBF remains unclear. One possibility is a change in the charge of histidine, possibly

affecting dynein ATPase or a dynein light chain that influences dynein ATPase activity.

To our knowledge, no data are available from the literature on the former hypothesis.

However, a dynein light chain conformation change has been reported to depend on pH-

induced changes in histidine ionization (Barbar et al., 2001). This dynein light chain,

LC8, has been originally found on outer dynein arms (Piperno & Luck, 1979), has been

reported to be important for flagellar beating and can be found as a dimer or monomer,

depending on pH-induced histidine ionization (Barbar et al., 2001). Thus, although

completely speculative, histidine charge changes on the outer dynein arm could be

important in pH-dependent modulation of CBF.

The baseline pHi of human airway tracheobronchial ciliated cells submerged in

nominally CO2/HCO3--free solutions (pH adjusted to 7.4) at room temperature was

surprisingly high at 7.49 ± 0.02. Poulsen et al. (1996), however, reported a similarly

high pHi (7.41 ± 0.09) in bovine tracheal epithelial cells bathed in CO2/HCO3--free

buffers. Similar pHi values were reported at 7.44 ± 0.01 in rat cardiac myocytes (Evans

70

et al., 2003), at 7.40 ± 0.02 in rat distal colon (Dagher et al., 1994), at 7.42 ± 0.02 in rat

ileum villus cells (Dagher et al., 1997), and at 7.67 ± 0.05 in rat ileum crypt cells

(Dagher et al., 1997). On the other hand, pHi of human nasal airway epithelial cells

bathed in CO2/HCO3--free medium reported by another group was considerably less at

7.08 – 7.16 (Willumsen & Boucher, 1992; Paradiso, 1997; Paradiso et al., 2003). The

reasons for these differences are not clear, but could be due to calibration issues,

differences in experimental temperatures (Roos & Boron, 1981), disparate cell types

and culturing methods (polarized cell culture). Our results using CO2/HCO3--buffered

solutions, however, agree with the published data: baseline pHi was 7.24 ± 0.02 in our

experiments, 0.24 pH units lower than in nominally CO2/HCO3--free solution (Paradiso

et al., 2003). The pHi changes in response to ammonium prepulse were similar to those

reported by others (Boyarsky et al., 1988; Singh et al., 1995; Paradiso, 1997; Ramirez et

al., 2000; Brett et al., 2002). Although both the basolateral and apical membrane was

available for NH4Cl in our submerged culture experiments, the shape of the pHi-plot

depicted in Figure 4-1/A and B suggested that the apical membrane responses were

predominant (Willumsen & Boucher, 1992; Boron et al., 1994; Singh et al., 1995;

Paradiso, 1997).

Our experiments in intact human tracheobronchial epithelial cells revealed a close

kinetic relationship between pHi and CBF in the examined pH-range of 7.0 – 7.8

(Figure 4-1, Figure 4-2). The changes occurred coincidentally at the beginning of the

alkalization and acidification phase of ammonium prepulse or external CO2-removal,

i.e. within the time resolution of the CBF measurements (3 sec) and pH recordings (10 -

20 sec).

However, in some experiments, CBF was slow to follow pHi during the pHi-

recovery from intracellular acidification or even remained low (Figure 4-1/B),

suggesting a prolonged or possibly irreversible, toxic effect of low pHi on cilia.

The mode of pHi-action on CBF is not completely clear, and the possibility with

histidine charge changes discussed above remains purely speculative. However, several

possibilities could be excluded. First, activation of sAC is unlikely, since all coverslips

and ALI cultures in this study, except those used for the CO2 removal experiments, were

perfused with CO2/HCO3--free solutions and exposed to ambient air for at least 30 min

71

before experimentation. Therefore, the intracellular [HCO3-] is expected to approach

zero in these cells and changes in pHi should not have been able to change sAC activity

via its HCO3-–sensitivity. Furthermore, the use of protein kinase inhibitors did not alter

the CBF response on pHi changes.

Protein kinase/phosphatase systems could also be plausible targets of pHi changes

since the catalytic efficiency of PKA is optimal at near neutral pH and is inhibited at

acidic pH (Cox & Taylor, 1995). Acidic pH not only inhibits PKA but also activates

phosphatases to de-phosphorylate PKA targets, while alkaline pH has the opposite

effect on both PKA and phosphatases (Reddy et al., 1998a). However, neither inhibition

of protein kinases with H-7, PKA with H-89 nor pre-stimulation of PKA by the

transmembrane adenylyl cyclase activator forskolin could attenuate CBF responses to

changing pHi. The same concentrations of the kinase inhibitors H-7 and H-89

significantly attenuated the CBF-stimulatory effect of forskolin, showing the efficacy of

these agents. The use of H-7 as a broad inhibitor also excluded other kinases relevant

for CBF (e.g., PKC). Similarly, pre-treatment with a mixture of two phosphatase

inhibitors failed to show any significant difference in the coupling of CBF and pHi

(Figure 4-3, Figure 4-4). Although it is more difficult to test the efficacy of

phosphatase inhibition, the combination of two inhibitors at the concentrations used (at

100, 3000 and 2000 times the estimated IC50 for phosphatase 1, 2A and 2B,

respectively) was likely sufficient to inhibit these enzymes.

Another important mediator of CBF regulation, [Ca2+]i, was also reported to be

influenced by pHi in several cell types (Thomas et al., 1979; Browning & Wilkins,

2002). However, we could not modify the coupling of pHi and CBF by emptying

internal Ca2+-stores with thapsigargin and inhibiting the influx of external Ca2+ using

nominally Ca2+-free bathing solutions (Figure 4-5/C). Simultaneous measurements of

[Ca2+]i and CBF during ammonium prepulse failed to show significant changes in

estimated [Ca2+]i when CBF changed (Figure 4-5/A), even when the estimated [Ca2+]i

was corrected for changes in the Kd of fura-2 due to pH (Lattanzio & Bartschat, 1991;

Browning & Wilkins, 2002). Although measurement of the fura-2 fluorescence of the

whole cell does not exclude the possibility of a localized rise in [Ca2+] in a subcellular

compartment, this possibility is unlikely since the absence of calcium in intracellular

stores as well as in the extracellular space did not change the results. The

72

physicochemical effect of increasing pH to decrease the concentration of ionized

calcium cannot account for the rise in CBF during alkalization as the opposite, namely a

decrease in CBF, would have been expected.

Our experiments on permeabilized cells (Figure 4-6, Figure 4-7) further confirm

that CBF is regulated by pHi and rule out the possibility that CBF changes during

ammonium prepulse were coupled to changes in extra- or intracellular NH4+ and/or NH3

concentrations rather than to changes in pHi. That ciliary beating was not completely

abolished in our permeabilized cells after 7 min perfusion with ATP and ARS-free

bathing solution could be due to the presence of non-diffusible pools of nucleotides

(Malaisse & Sener, 1987; Aprille, 1988; Detimary et al., 1996) and by residual

production of ATP close to cilia.

Together, these data suggest that phosphorylation/dephosphorylation events and

changes in [Ca2+]i are not responsible for the pHi–mediated changes in CBF and support

a direct action of pHi on axonemal proteins. Studies performed on demembranated

mammalian spermatozoa are of special interest in this regard because cilia and sperm

flagella share considerable ultrastructural similarities as well as some similarities in

their beat regulation, e.g. by PKA (for review see, Urner & Sakkas, 2003). Most

published experiments on demembranated spermatozoa demonstrate that mild

alkalization augments flagellar beat frequency (FBF) of spermatozoa of different

species (Gibbons & Gibbons, 1972; Brokaw & Kamiya, 1987; Keskes et al., 1998a).

One of these studies (Keskes et al., 1998a) also showed that human spermatozoa that

lack outer dynein arms failed to increase beat frequency during alkalization suggesting

that outer dynein arms, that mainly determine the frequency of ciliary beating (Brokaw

& Kamiya, 1987), might be directly involved in the response to changing pHi. Whether

the effects of pH on spermatozoa are mediated through the same mechanisms, remains

unclear since some of the pH effects in spermatozoa could be due to the activation of

sAC.

Our results show that intracellular alkalization results in faster ciliary beating,

while intracellular acidification attenuates CBF in human tracheobronchial epithelial

cells. Variations in pHi of airway epithelia may occur in vivo in response to shifting

luminal CO2 concentrations from 5% to 0.02% during a full breathing cycle (Willumsen

73

& Boucher, 1992). It is intriguing to speculate that a possible pHi change during the

breathing cycle might influence CBF in the large airways. If it occurs, it would result in

a faster ciliary beating during inspiration. Furthermore, airway diseases associated with

acidification of the surface liquid and possibly epithelium (e.g. asthma) could depress

ciliary activity, possibly contributing to the decrease in mucociliary clearance seen in

these diseases.

5.2. Expression of sAC in human airway epithelial cells

Our RT-PCR, immunoblot and immunostaining studies have shown the first time

that sAC is expressed in normal ciliated HAECs, both in tracheal tissue and cell culture.

Recently, Calu-3 cells, a human adenocarcinoma cell line with airway serous cell

properties, have also been shown to express sAC (Sun et al., 2004; Wang et al., 2005).

Our immunohistochemical (Figure 4-10) and immunocytochemical (Figure 4-11)

studies clearly demonstrate that sAC is expressed in the apical membrane and in the

cilia of HAECs. Within the cilia, sAC is co-localized with ciliary tubulin.

Our RT-PCR studies revealed cDNA fragments of three different sizes. The

shortest fragment was 100% identical to position 388-630 of human sAC cDNA (gi

21614540, NM 018417). Four clones of the medium sized fragment were sequenced,

two of which were 100% identical to a recently published sAC variant containing 37

extra 5’-nucleotides to exon 5 derived from the intron sequence (Geng et al., 2005).

However, two other clones showed variability in one base and one of them also missed

a T (Figure 4-8/D). We have also found a heretofore unpublished sAC transcript

containing an additional 12 bps of intron sequence, in addition to the 37 bps of the

already published sAC-variant. In addition to these sAC-transcript variants, a sAC

variant missing the entire exon 5 has also been described recently (Geng et al., 2005).

The reverse primer we used overlapped exon 5 and 6, therefore this short sAC-transcript

missing exon 5 might have escaped our notice. It is not clear whether all these variant

transcripts actually give rise to distinct polypeptides (Geng et al., 2005).

The preliminary experiment shown in Figure 4-12 suggests that increasing

[HCO3-]i results in faster ciliary beating in HAEC. In these experiments we used the

74

detergent saponin, instead of the pore forming agent Staphylococcus aureus alpha-toxin,

in order to achieve a better permeability of the basolateral membrane. Another advance

of saponin is that, as we found, cells become permeabilized randomly following a short

exposure to this detergent. This feature allowed us to perform simultaneous control

experiment using cells with intact basolateral membrane from the same culture.

However, we were also faced to some technical difficulties. Permeabilization by

saponin developed suddenly and overexposure of permeabilized cells to saponin often

resulted in cellular death. Monitoring of CBF in the absence of basolateral ATP during

saponin-exposure helped us to recognize immediately when basolateral membrane had

become permeable (indicated by falling CBF resulting from leaking ATP) and thus, to

withdraw saponin right away.

The opposite CBF-response on CO2/HCO3--containing solution in permeabilized

and intact cells suggests a HCO3-- rather than a pHi-effect; the increase in CBF in

permeabilized cells following exposure to CO2/HCO3- cannot be attributed to

intracellular alkalization, because basolateral pH was constant (kept at 7.2) and the

basolateral membrane was supposed to be freely permeable to both CO2 and HCO3-.

Indeed, if any change in pHi occurred, one can expect a slight cytosolic acidification

because of apical CO2-load without any significant apical HCO3--influx. However,

further studies, e.g. experiments using specific sAC- and PKA-inhibitors are needed to

demonstrate that HCO3--related increase in CBF results from sAC-activation and

cAMP-production.

Although here we have shown that sAC is expressed in the cilia of human ciliated

airway epithelial cells and our preliminary results on basolaterally permeabilized cells

loaded with CO2/HCO3- suggest that sAC is involved in the regulation of CBF, the

physiological role of sAC in these cells remains to be defined. One possible function of

sAC is to maintain a certain concentration of cAMP in the ciliary shaft. As mentioned in

section 1.3.4, rising [Ca2+]i has no effect on CBF in the absence of cAMP (Ma et al.,

2002). sAC, as an enzyme which is regulated independently of G-protein coupled

receptors but can be activated by certain cytosolic ions, might be a constitutive source

of cAMP in the ciliary compartment. Patch-clamp experiments reported by Ma et al.

(Ma et al., 2002) supports this hypothesis: in the absence of cAMP in the pipette

solution (i.e., cytosolic solution), CBF did not differ in cells perfused with a pipette

75

solution containing less than 1 nM Ca2+ compared to those cells which were perfused

with 100 nM or even 500 nM of Ca2+. Importantly, the pipette solutions in these

experiments did not contain any ion which is a known activator of sAC. Furthermore, an

apparent contradiction between these patch-clamp experiments and an older study of

basolaterally permeabilized canine tracheal epithelium (Kakuta et al., 1985), where

CBF was shown to be regulated by cytosolic Ca2+ in the absence of externally applied

cAMP could also be explained by this hypothesis: basolateral (i.e., intracellular)

solution used in the latter experiments contained SO3--ion, which is a weak activator of

sAC in vitro (Chen et al., 2000). Further studies from our laboratory not presented here

have shown that several types of tmACs, namely type 1 (tmAC1), tmAC3 and tmAC8

are also expressed in HAECs (Sutto et al., 2005). However, immunocytochemistry has

shown that none of them is colocalized with ciliary tubulin (figure not shown) further

supporting the theory that cAMP produced by sAC has a special role in the regulation of

ciliary beating.

Speculatively, sAC might also function as a buffer of pHi-related CBF alterations.

As we have shown, cellular CO2-load results in a fall in CBF because of cytosolic

acidification. Cellular CO2-load will shift the equation (1) to the left:

(1) H+ + HCO3- ⇔ CO2 + H2O.

Therefore, increasing [HCO3-]i might produce cAMP through the activation of

sAC, counterbalancing the attenuation of CBF upon increasing H+-concentration (i.e.,

acidification). In contrast to CO2-load, acidification caused by H+-load will result in a

decrease in [HCO3-]i as equation (1) will be shifted to the right; therefore sAC activation

by HCO3- will not be expected. However, a latest report has demonstrated that two

(randomly selected) prokaryotic adenylyl cyclases, Slr1991 and CyaB1, are actually

stimulated by dissolved CO2 and not by HCO3- (Hammer et al., 2006). If this result

could be extrapolated to human sAC, CBF decrease upon cellular H+-load could also be

opposed by sAC-related cAMP-production, however, our experiments using ammonium

prepulse showed that there was no significant difference in pHi-related CBF-transients

when protein kinase inhibitors were used (but those experiments were performed in the

virtual absence of CO2 and in open chamber).

76

5.3. Expression and putative function of catecholamine transporters in

human airway epithelial cells

RT-PCR using specific primers for four different catecholamine transporters

demonstrated the expression of OCT-1, EMT and NET in HAECs. Freshly isolated

human bronchial arterial smooth muscle cells expressed the same extraneuronal NE

transporters, i.e. OCT-1 and EMT, therefore we used this model for characterizing NE

uptake assay, as described previously (Horvath et al., 2001; Horvath et al., 2002). NE

uptake was mainly mediated by EMT, being sensitive to the inhibitors corticosterone

and O-methyl-isoprenaline (OMI), specific blockers of EMT but not OCT-1 and OCT-2

(Martel et al., 1999; Hayer-Zillgen et al., 2002). NE uptake was acutely blockable by

various GSs including budesonide and methylprednisolone. The inhibitory action of

GSs was rapidly reversible, and mediated by a non-genomic pathway possibly through a

specific GS binding site in the plasma membrane.

The physiologic function of catecholamine transporters expressed in HAECs is

not completely understood. We hypothesize that they may have a role in inactivating

hydrophilic β-adrenoceptor agonist drugs (e.g., salbutamol, formoterol) by mediating

their cellular uptake into the airway epithelial cells. In this regard, the rapid inhibitory

effect by GSs on this transport process might be of importance providing additional

rationale for combined inhaled β2-agonist/GS therapy in obstructive lung disease

(Figure 4-15 and Figure 4-16). Physiological mechanisms presumably underlying the

favorable effects of this combinatory treatment include increasing β2-receptor

expression (Mak et al., 1995a), restoring G-protein/β2-receptor coupling (Mak et al.,

1995b), and inhibition of β2-receptor downregulation (Hadcock et al., 1989) induced by

inhaled GSs. Conversely, β2-agonists prime the glucocorticoid receptor and effect its

nuclear localization by modulating its phosphorylation (Eickelberg et al., 1999). The

mutual potentiation of GSs’ and β2-agonists’ action through these mechanisms involves

gene transcription and takes hours to manifest itself. Our results presented here are in

keeping with an earlier report of a rabbit model from our laboratory (Horvath et al.,

2001) and provide evidence of a rapid, nongenomic inhibitory action of GSs on EMT-

mediated cellular NE uptake. This GS-mediated effect could increase the local

77

concentration of β2-agonists at β2-adrenergic receptor sites and hence potentiate β-

agonists’ effect on airway epithelium and mucociliary activity.

However, several questions remain to be answered. First, our NE uptake

experiments were carried out in a human bronchial arterial smooth muscle cell model

expressing OCT-1 and EMT. Therefore, our data on catecholamine transport and the

potency of GSs to inhibit this transport are to be confirmed in HAECs, which seem to

express NET as well, besides OCTs. Steroid inhibition of NET has been reported

(Horishita et al., 2002; Toyohira et al., 2003), therefore it can be predicted that GSs will

inhibit catecholamine uptake in HAECs as well, similarly to vascular smooth muscle

cells.

The second issue to be considered is whether β2-agonists used in the clinical

practice are substrates for and thus functionally inactivated by the corresponding

catecholamine transporters, similarly to NE. Therefore, further uptake experiments are

needed using these drugs.

Third, the potential inhibition of β2-agonist uptake by airway epithelial cells

could theoretically decrease the inhaled drug’s availability in deeper tissue layers such

as bronchial smooth muscle cells, the main target of β2-adrenergic agonist drugs.

Although there have no clinical data been reported supporting such a disadvantageous

effect of the combinatory inhaled β2-agonist/GS therapy, additional in vitro (e.g.,

transport studies on polarized epithelial cell culture) and in vivo studies are necessary to

clarify this issue.

A most recent study by Lips et al. has shown the expression of all three isoforms

of OCTs in HAECs (Lips et al., 2005). The reason for the discrepancy between this

report and our results regarding OCT-2 expression is not clear and might be explained

by methodological differences (e.g., mRNA extraction from cultured and

redifferentiated vs. abraded epithelial cells). The remaining NE uptake after blocking

EMT by corticosterone (Figure 4-14) could suggest the presence of functional OCT-2

or even OCT-1, which transporters are less sensitive to corticosterone (Hayer-Zillgen et

al., 2002). However, the experiment depicted in Figure 4-16 makes this assumption less

likely since concentrations of corticosterone > 1 µM did not decrease uptake into the

cells as would have been expected if OCT-2 or OCT-1 were to play a role, given the

78

IC50 of their inhibition by corticosterone, which is 34.2 µM and 21.7 µM, respectively

(Hayer-Zillgen et al., 2002). Alternatively, the remaining NE uptake after inhibition

could result from nonspecific uptake, possibly related to the measurement technique, or

less likely due to binding of NE to receptors on the cell surface.

Interestingly, besides the extraneuronal NE transporters (OCT-1 and EMT),

cultured HAECs seem to express the neuronal NE transporter (NET) mRNA as well.

Furthermore, total RNA sample from human kidney (purchased from Ambion, Austin,

TX, USA), also contained NET mRNA. NET has been known to be localized mainly at

sympathetic synapses (Koepsell et al., 2003). Besides being expressed in neurons and

adrenal neuroendocrine (chromaffine) cells, NET has been reported in some

extraneuronal tissues, namely in placenta and pulmonary capillary endothelial cells

(Eisenhofer, 2001; Bottalico et al., 2004). The known principal function of NET is

deactivating catecholamines. At the adrenergic synaptic cleft, NET participates in

terminating the activation of postsynaptic receptors; in pulmonary capillaries, NET

clears circulating catecholamines from the bloodstream; placental NET is supposed to

serve as a protective mechanism preventing vasoconstriction in the placental vascular

bed securing a stable blood flow to the fetus (Catravas & Gillis, 1983; Bryan-Lluka et

al., 1992; Westwood et al., 1996; Eisenhofer, 2001; Koepsell et al., 2003). However, to

our knowledge, neither in airway epithelial cells nor in kidney has NET expression been

reported thus far. Although contamination with neuronal mRNA might be possible in

the total RNA sample from human kidney, it seems unlikely in the case of HAECs,

since there is no evidence for adrenergic neurons in the surface epithelium of the human

bronchi (Partanen et al., 1982; Wanner et al., 1996; Belvisi, 2002). However, further

experiments, including immunofluorescence and functional studies are necessary to

confirm this result.

Perhaps the most significant implication of OCTs in airway epithelium is their

ability to mediate cellular ACh release, an attribute of the newly discovered

autocrine/paracrine non-neuronal cholinergic systems (Wessler & Kirkpatrick, 2001;

Wessler et al., 2003). Such an autocrine cholinergic regulation has been suggested in

many tissues (Wessler et al., 2003). An earlier functional study carried out in human

placenta, a useful model of non-neuronal cholinergic systems, suggested that ACh was

released by OCT-1 and EMT (Wessler et al., 2001). A most recent report by Lips et al.

79

mentioned above (Lips et al., 2005) has provided evidence that all three isoforms of

OCTs are expressed in the ciliated cells of human airway, are located in the luminal

membrane of ciliated cells, and two of them, OCT-1 and OCT-2 (but not EMT) mediate

ACh uptake and efflux when expressed in Xenopus laevis oocytes. Moreover,

budesonide (but not beclomethasone) inhibited OCT-2 mediated ACh efflux (Lips et al.,

2005). Human bronchial epithelium possesses all the components essential for a

functioning non-neuronal cholinergic system (Proskocil et al., 2004; Lips et al., 2005),

the physiological role of which, however, has not yet been elucidated. Several data

suggest that non-neuronal ACh release may contribute to CBF regulation in the airways

(as discussed in paragraph 1.2.3.2). Certainly, the bronchial non-neuronal cholinergic

system is a new, promising field of research in airway pathophysiology.

Taking these data together further conclusions can be drawn. An expression

pattern of OCTs, β-adrenergic receptors and NET in airway epithelial cells can make

one speculate that an extraneuronal adrenergic autocrine/paracrine system, similarly to

the non-neuronal cholinergic system, exists in the airway epithelium. This hypothesis,

although clearly speculative, could be supported by certain facts. First, OCTs, as bi-

directional transporters, are able to mediate not only uptake but also efflux of

catecholamines (Koepsell et al., 2003; Ciarimboli & Schlatter, 2005). Considering that

epinephrine is more potent stimulator of β-adrenergic receptors than NE, it is

noteworthy that, in terms of substrate specificity, both OCT-1 and EMT prefer

epinephrine over NE (Martel et al., 1999; Eisenhofer, 2001). Second, although the

expression of β-adrenergic receptors in HAEC has long been known, the physiological

source of their ligand has not been identified thus far. It seems very likely that

circulating catecholamines are unable to reach these receptors, at least those ones

expressed in the apical plasmamembrane (Wong et al., 1988a), and neuronal activation

is also impossible because bronchial surface epithelium seems to lack adrenergic

innervation (Partanen et al., 1982; Wanner et al., 1996). Looking for a physiological

source of β-adrenergic receptor ligand in the airway lumen, inflammatory cells appear

to be possible candidates, inasmuch they synthesize and secrete catecholamines

(Spengler et al., 1994; Musso et al., 1996; Bergquist & Silberring, 1998). Although this

hypothesis might be reasonable, lymphocytes seem to synthesize primarily NE (Musso

et al., 1996; Bergquist & Silberring, 1998; Musso et al., 1998), which is less potent

80

activator of β2-adrenergic receptors than epinephrine. The other possibility is that

airway epithelial cells release epinephrine, which would not be unprecedented:

epithelial cells in the skin, namely keratinocytes do synthesize and release both NE and

epinephrine thereby regulating keratinocyte differentiation in an autocrine, and

melanogenesis in melanocytes in a paracrine way (Schallreuter et al., 1992; Schallreuter

et al., 1995; Gillbro et al., 2004). In contrast to the known autocrine systems in HAEC

(i.e., the purinergic and the non-neuronal cholinergic system), which primarily regulate

[Ca2+]i, the extraneuronal β-adrenergic system would be an autocrine regulator of

intracellular cAMP, maybe in orchestration with sAC, perhaps in different subcellular

compartments. However, all these speculations should be confirmed by proving that

HAECs possess the armament for autocrine catecholaminergic regulation, i.e. that they

express the enzymes involved in the biosynthesis and degradation of catecholamines. If

this autocrine machinery existed in airway epithelium, NET and/or OCTs might be

responsible for catecholamine ‘reuptake’ into HAECs.

81

6. CONCLUSIONS

Our experiments presented here provide novel information on the regulation of

ciliary beating in HAECs.

1. Here we have shown the first time that intracellular acidification attenuates, while

intracellular alkalization augments ciliary beating in HAECs.

2. Phosphorylation/dephosphorylation events and [Ca2+]i changes do not mediate pHi-

induced CBF changes. pHi might directly act on the ciliary motile machinery.

3. RT-PCR, Western blotting, immunohisto- and immunocytochemical studies

revealed that ciliated HAECs express sAC, a recently cloned unique adenylyl

cyclase, which is regulated by certain cytosolic ions like HCO3- and Ca2+, but

independent of pHi-changes and G-protein regulation.

4. Immunofluorescence studies in tracheal tissue and cultured HAECs have shown

that sAC is located in the cilia and in the apical membrane of ciliated HAECs. Our

preliminary functional studies suggest that sAC may be involved in the regulation of

CBF.

5. We have also demonstrated that HAECs express mRNA for extraneuronal (OCT1,

EMT) and neuronal (NET) catecholamine transporters.

6. Using a model of airway vascular smooth muscle cell expressing OCT1 and EMT,

we have shown that NE uptake is acutely and reversibly blockable by a non-genomic

action of various GSs including budesonide and methylprednisolone.

82

SUMMARY

BACKGROUND. Ciliary function has a pivotal role in airway mucus clearance,

but the regulation of ciliary beat frequency (CBF) is still not completely understood. In

these studies we focused on two issues. (i) In human airway epithelial cells (HAEC),

transmembrane adenylyl cyclases (tmAC) are the only known source of cAMP. Soluble

adenylyl cyclase (sAC), a recently cloned enzyme, regulates flagellar motility in sperm.

In contrast to tmACs, sAC is insensitive to G-protein activation, but is stimulated by

HCO3-, independently of H+. Here we examined the expression and cellular distribution

of sAC in HAECs. Because functional experiments of sAC are complicated by the fact

that intracellular HCO3- concentration and intracellular pH (pHi) correlate, and neither

has ever been examined for ciliary effects, we also studied how pHi itself influences

CBF in HAECs. (ii) β-adrenergic agonists are the most important ciliostimulant drugs

of airway epithelium. Organic cation transporters (OCT) mediate non-neuronal

norepinephrine (NE) uptake and thus, inactivate NE at adrenergic receptors. NE uptake

by certain OCTs has been reported to be inhibited by glucocorticoids (GSs), thereby

increasing NE concentration at receptor sites. We hypothesized that inhaled β-

adrenergic agonists could also be potentiated by GSs inhibiting their uptake into

HAECs, and thus enhancing ciliostimulation. Therefore we tested the expression of

OCTs in HAECs and examined the characteristics, including GS-inhibition, of NE

transport.

RESULTS. (i) We have shown that sAC (both mRNA and protein) is expressed in

HAECs. Confocal microscopy localized sAC in the ciliary shaft, co-localizing with

axonemal tubulin. Preliminary functional studies have suggested that cytosolic HCO3-

stimulates CBF in HAECs. We also demonstrated that relatively small changes in pHi

result in significant changes in CBF in HAECs (increasing pHi increases CBF and vice

versa). Changing pHi may directly act on the ciliary motile machinery, as the

involvement of the kinase/phosphatase system, Ca2+ and sAC has been excluded. (ii)

mRNA for certain OCTs (OCT-1 and EMT) as well as neuronal epinephrine transporter

(NET) are expressed in HAECs. NE uptake of airway vascular smooth muscle cells

expressing both OCT-1 and EMT was acutely inhibited by GSs in a non-genomic way.

83

Further experiments are needed to define the role of catecholamine transporters

expressed by HAECs.

84

ÖSSZEFOGLALÁS

A légúti mucociliáris clearance fenntartásában a ciliumok működésének

kulcsszerepe van, azonban a ciliumok csapási frekvenciája (CBF) szabályozásának sok

részlete még tisztázatlan. Jelen vizsgálataink két kérdés tanulmányozását célozták. (i)

Az emberi légúti hámsejtek (HAEC) jelenleg ismert ciklikus AMP-t (cAMP) termelő

enzimei az ún. transzmembrán adenilciklázok (tmAC) csoportjába tartoznak. Egy

újonnan klónozott enzim, a szolubilis adenilcikláz (sAC) részt vesz a spermiumok

ostormozgásának szabályozásában. A sAC a tmAC-októl elkülönülő csoportot képez:

szemben a tmAC-okkal, működését nem befolyásolja a G-protein, viszont aktiválja a

HCO3--ion (függetlenül a pH-tól). Jelen kísérleteinkben egyrészt a sAC HAEC-ekben

való expresszióját és sejten belüli lokalizációját vizsgáltuk. Mivel a sAC funkcionális

vizsgálatát nehezíti az a tény, hogy a sejten belüli HCO3--koncentráció ([HCO3

-]i) és pH

(pHi) összefüggnek, és mind a [HCO3-]i, mind a pHi CBF-ra gyakorolt hatása

ismeretlen, jelen tanulmányban azt is megvizsgáltuk, hogy maga a pHi változása hogyan

hat a CBF-ra ezekben a sejtekben. (ii) A β-agonisták a legfontosabb ismert

ciliostimuláns gyógyszerek a légúti hám esetében. Az organikus kation transporterek

(OCT) felelősek a nem-neuronális sejtekbe történő noradrenalin (NE) felvételért és

ezáltal részt vesznek a NE inaktiválásában. A glukokortikoszteroidok (GS) gátolják a

bizonyos OCT-eken keresztül történő NE-felvételt és így növelhetik a NE

koncentrációját az adrenerg receptorok környezetében. Hasonló mechanizmust

feltételezve az inhalált β-agonisták és a HAEC-ek esetében, megvizsgáltuk az OCT-ek

expresszióját a HAEC-ekben és a NE transzport OCT-ek általi transzportjának

jellemzőit és a GS-ok gátló hatását is.

EREDMÉNYEK. (i) Kimutattuk mind a sAC mRNS, mind a sAC fehérje jelenlétét a

HAEC-ekben. Konfokális mikroszkóp segítségével a sAC a csillószőrös HAEC-ek

felszíni plazmamembránjában és magában a ciliumokban volt látható. A csillószőrökben

a sAC a tubulinnal azonos lokalizációban jelent meg. Funkcionális vizsgálataink kezdeti

eredményei arra utalnak, hogy a sejten belüli HCO3- serkenti a CBF-t ezekben a

sejtekben. Azt is kimutattuk, hogy relatíve kis pHi változás jelentős CBF változást

eredményez a HAEC-ekben (emelkedő pHi növeli a CBF-t és vica versa). A pHi

változás valószínűleg közvetlenül a ciliaris proteineken keresztül fejti ki ezt a hatását,

85

mivel a kináz/foszfatáz rendszer, a citoplazmatikus Ca2+ és a sAC szerepét kísérleteink

kizárták. (ii) Egyes OCT fehérjéket (OCT-1 és EMT), csakúgy, mint a neuronális

adrenalin transportert (NET) kódoló mRNS kimutatható volt a HAEC-ekben. A NE-

felvételt akutan, nem-genomiális mechanizmussal blokkolták a GS-ok mind az OCT-1,

mind az EMT mRNS-ét expresszáló légúti vascularis simaizomsejtekben. További

vizsgálatok szükségesek a katekolamin transzporterek HAEC-ekben betöltött élettani

szerepének tisztázásához.

86

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LIST OF PUBLICATIONS

Publications relevant to theses

Articles

1. Sutto Z., Conner G.E., Salathe M.: Regulation of human airway ciliary beat frequency

by intracellular pH. J. Physiol. 560:519-32, 2004. IF = 4.346

2. Horvath G., Sutto Z., Torbati A., Conner G.E., Salathe M., Wanner A.:

Norepinephrine transport by the extraneuronal monoamine transporter in human

bronchial arterial smooth muscle cells. Am. J. Physiol. Lung Cell. Mol. Physiol.

285:L829-37, 2003. IF = 3.735

Abstracts

3. Sutto Z., Nlend M.C., Conner G.E., Fregein N., Salathe M.: Polarized expression of

Ca2+-sensitive adenylyl cyclases in human airway epithelial cells. Proc. Am. Thorac.

Soc. 2: A221, 2005.

4. Sutto Z., Conner G.E., Salathe M.: Evidence for direct regulation of ciliary beat

frequency by intracellular pH. Am. J. Respir. Crit. Care. Med. 169: A673, 2004.

5. Nlend M.C., Sutto Z., Horvath G., Conner G.E., Salathe M.: Expression of calcium-

sensitive adenylyl cyclases isoforms in human airway epithelial cells. Am. J. Respir.

Crit. Care. Med. 169: A675, 2004.

6. Sutto Z., Conner G.E., Salathe M.: Measurement of ciliary beat frequency on

permeabilized epithelial cells: correlation between intracellular pH and ciliary

beating. FASEB 18: A1053, 2004.

7. Sutto Z., Conner G.E., Salathe M.: Intracellular pH regulates ciliary beat frequency

in human airway epithelial cell. Am. J. Respir. Crit. Care. Med. 167: A398, 2003.

8. Horvath G., Sutto Z., Salathe M., Conner G.E., Wanner A.: Acute inhibition of the

extraneuronal catecholamine transporter by glucocorticosteroids in human bronchial

arteries. Am. J. Respir. Crit. Care. Med. 167: A1006, 2003.

112

9. Sutto Z., Conner G.E., Salathe M.: Bicarbonate regulates ciliary beat frequency in

human airway epithelial cell. Mol. Biol. Cell. 13: 191a, 2002.

10. Horvath G., Sutto Z., Salathe M., Conner G.E., Wanner A.: Nongenomic action of

corticosterone on norepinephrine uptake by human bronchial arterial smooth muscle

cells. Mol Biol Cell 13: 273a, 2002.

Other publications

Articles

1. Sutto Z.: Differential diagnosis of cough. Cough relief. Praxis 15(11): 11-19, 2006.

[Hungarian]

2. Sutto Z., Magyar P.: Tuberculosis and pregnancy. Med. Thor. 52: 195-9, 1999.

[Hungarian]

3. Sutto Z., Bartfai Z., Lantos A., Major T. jr., Vadasz G., Varnai Zs., Zsiray M.:

Human polyspecific intravenous immunoglobulin therapy in Wegener’s

granulomatosis. Med. Thor. 52: 134-8, 1999. [Hungarian]

4. Sutto Z.: New perspectives in the diagnosis and treatment of Wegener’s

granulomatosis. Orvoskepzes 73: 99-103, 1998. [Hungarian]

5. Vajda E., Sutto Z., Zsiray M., Appel J., Lantos A., Kardos K.: Malignant

mesothelioma of the pleura. Orv. Hetil. 137(5): 233-8, 1996. [Hungarian]

6. Nagy L., Sutto Z., Tolnay E., Terek K., Orosz M., Szentpaly O.: Eosinophil

secretory products in acute severe asthma. Orv. Hetil. 137(3): 121-4, 1996.

[Hungarian]

7. Sutto Z.: Environmental control of allergens. Haziorvos Tovabbkepzo Szemle 1:

186-8, 1996. [Hungarian]

8. Sutto Z., Nagy L.: Pulmonary edema due to heroin abuse. Med. Thor. 47: 341-7,

1994. [Hungarian]

113

9. Nagy L., Orosz M., Sutto Z.: Inflammatory mediators of allergy in upper respiratory

tract allergy. Orv. Hetil. 134:743-4, 1993. [Hungarian]

10. Nagy L., Sutto Z.: Neuroendocrine peptides in the course of asthma bronchiale.

Orvoskepzes 67: 264-8, 1992. [Hungarian]

Book Chapters

11. Tarjan E., Muller V., Orosz Zs., Sutto Z., Magyar P.: Pneumonia. In: Test questions

and explanations in the field of pulmonology. Ed. Magyar P., Lantos A. Medicina,

Budapest, Hungary, 2005., pp 134-146. [Hungarian]

12. Lantos A., Magyar Pal, Sutto Z.: Mycotic lung diseases. In: Test questions and

explanations in the field of pulmonology. Ed. Magyar P., Lantos A. Medicina,

Budapest, 2005., pp 173-180. [Hungarian]

13. Magyar P., Bohacs A., Sutto Z., Tamasi L., Vajda E.: Diffuse interstitial lung

diseases, eosinophilic lung diseases. In: Test questions and explanations in the field

of pulmonology. Ed. Magyar P., Lantos A. Medicina, Budapest, 2005., pp 219-232.

[Hungarian]

14. Lantos A., Appel J., Bohacs A., Bartfai Z., Magyar P., Rozgonyi Zs., Sutto Z.,

Tarjan E., Vastag E., Vajda E., Vardy Visi K.: Test questions related to case reports.

In: Test questions and explanations in the field of pulmonology. Ed. Magyar P.,

Lantos A. Medicina, Budapest, 2005., pp 341-410. [Hungarian]

15. Zsiray M., Sutto Z.: Interstitial lung diseases of unknown origin. In: Pulmonology.

Ed. Magyar P., Hutas I., Vastag E. Medicina, Budapest, 1998., pp 538-557.

[Hungarian]

16. Sutto Z.: Inflammatory diseases of the upper respiratory tract. In: Pulmonology. Ed.

Magyar P., Hutas I., Vastag E. Medicina, Budapest, 2002., pp 262-265. [Hungarian]

17. Orosz M., Szentpaly O., Bartfai Z., Nagy L., Sutto Z., Temesi G., Tolnay E.:

Airway diseases induced by grain dust and flour. In: Environmental pollutants and

the respiratory tract. Vol. 7. Ed. Szabo T., Miriszlai E. Heviz, Hungary 1997.

[Hungarian]

114

Abstracts

18. Szondy K., Ostoros Gy., Sutto Z., Lantos A., Magyar P.: Gemcitabine plus cisplatin

treatment for advanced NSCLC. Lung Cancer 25 (Suppl.1): S17, 1999.

19. Orosz M., Szentpaly O., Bartfai Z., Sutto Z., Nagy L.: Occupational allergic airways

diseases among health care workers. Med. Thor. 51: S24, 1998. [Hungarian]

20. Horvath G., Vajda E., Sutto Z., Vastag E., Magyar P.: Bronchodilating responses to

inhaled 20 μg, 80 μg and 120 μg ipratropium bromide followed by 400 μg fenoterol

in stable COPD, comparison with fenoterol alone. Eur. Respir. J. Abstracts Book

39S, 1998.

21. Varnai Zs., Lantos A., Sutto Z., Major T. jr.: Artificial ventilation during

bronchofiberoscopy in mechanically ventilated patients. Med. Thor. 49: S50, 1996.

[Hungarian]

22. Lantos A., Major T. jr., Sutto Z., Tarjan E., Vajda E., Varnai Zs.: Talc pleurodesis

for the treatment of secondary spontaneous pneumothorax with persistent air leak.

Med. Thor. 49: S51, 1996. [Hungarian]

23. Nagy L., Sutto Z., Tolnay E., Terek K., Orosz M., Szentpaly O.: Eosinophil

secretory products in acut severe asthma. Allergy, 51 (30): S62, 1996.

24. Lantos A., Bartfai Z., Sutto Z., Szondy K., Tarjan E., Varnai Zs., Zsiray M.:

Chemical pleurodesis with talc or tetracycline derivatives of malignant pleural

effusions Onkologia (Suppl.): 18, 1995.

25. Nagy L., Orosz M., Tolnay E., Sutto Z.: Eosinophil secretory products (ECP, EXP)

in acute severe asthma. Med. Thor. 1994. 47: S9, 1994.

115

ACKNOWLEDGEMENTS

The author wishes to express his sincere gratitude to all individuals at the Division

of Pulmonary and Critical Care Medicine, University of Miami School of Medicine

(UM) and the Department of Respiratory Diseases, Semmelweis University (SU) who

have contributed to these studies with special thanks to:

Matthias Salathe, M.D. (UM)

Gábor Horváth, M.D., Ph.D. (SU, UM)

Prof. Pál Magyar, M.D., D.Sc. (SU)

Marie-Christine Nlend, Ph.D. (UM)

Gregory E. Conner, Ph.D. (UM)

Prof. Adam Wanner, M.D. (UM)

Andreas Schmid, M.D. (UM)

Nathalie Schmid (UM)

Attila Kiss, M.D., Ph.D. (SU)

Erika Vajda, M.D. (SU)

Kinga Emődi, M.D. (SU)

These studies were completed in the asthma research laboratories of the Division

of Pulmonary and Critical Care Medicine at the University of Miami School of

Medicine (Miami, FL, USA) in 2002-2004.

This work was supported by grants from the American Heart Association (to Z.S. and

G.H.) and the National Institute of Health research grants (to M.S., G.C. and A.W.).