JPET Fast Forward. Published on May 14, 2009 as …jpet.aspetjournals.org/content/jpet/early/2009/05/14/... · 4, 10 glucose, 2 glutamine, and 20 HEPES adjusted to pH 7.4 with NaOH),

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Title Page

Histamine H1 Receptor Induces Cytosolic Calcium Increase and Aquaporin

Translocation in Human Salivary Gland Cells

Ji-Hyun Kim, Seong-Hae Park, Young Wha Moon, Sungmin Hwang, Donghoon Kim, Su-

Hyun Jo, Seog Bae Oh, Joong Soo Kim, Jeong Won Jahng, Jong-Ho Lee, Sung Joong Lee,

Se-Young Choi, and Kyungpyo Park

Department of Physiology (J.-H.K., S.-H.P., S.H., D.K., S.B.O., J.S.K., S.J.L., S.-Y.C.,

K.P.)

and Department of Oral & Maxillofacial Surgery (J.W.J, J.-H.L),

Dental Research Institute, Seoul National University School of Dentistry, Seoul 110-749,

Korea.

Department of Biology, The Catholic University of Korea College of Medicine, Seoul, 137-

040, Korea (Y.W.M).

Department of Physiology, Kangwon National University School of Medicine, Chuncheon

200-701, Korea (J.-H.K., S.-H.J.).

JPET Fast Forward. Published on May 14, 2009 as DOI:10.1124/jpet.109.153023

Copyright 2009 by the American Society for Pharmacology and Experimental Therapeutics.

This article has not been copyedited and formatted. The final version may differ from this version.JPET Fast Forward. Published on May 14, 2009 as DOI: 10.1124/jpet.109.153023

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Running Title: Histamine H1 receptor in human salivary gland cells

Address correspondence to:

Se-Young Choi, Ph.D.

Department of Physiology, Seoul National University School of Dentistry,

28 Yeongun, Jongno, Seoul, 110-749, Korea

Phone: 82-2-740-8650, Fax: 82-2-762-5107, E-mail: [email protected]

Number of text pages : 38

Number of tables : 0

Number of figures : 7

Number of references : 40

Words in Abstract : 237

Words in Introduction : 542

Words in Discussion : 1477

Nonstandard abbreviations used:

AQP-5, Aquaporin-5; [Ca2+]i, Cytosolic free Ca2+ concentration; Fura-2-AM, Fura-2-

acetoxymethyl ester; HRP, horseradish peroxidase; GFP, green fluorescent protein.


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Abstract

One of the common side effects of anti-histamine medicines is xerostomia (dry mouth). The

current consensus is that anti-histamine-induced xerostomia comes from an anti-muscarinic

effect. While the effect of anti-histamines on salivary secretion is both obvious and

significant, the cellular mechanism is still unclear due to a lack of knowledge of histamine

signaling in human salivary glands. Here we have studied histamine receptors and the effect

of anti-histamines on human submandibular acinar cells. In primary cultured human

submandibular gland and a HSG cell line, histamine increased the intracellular Ca2+

concentration. The histamine-induced [Ca2+]i increase was inhibited by histamine H1

receptor-specific antagonists, and the expression of the functional histamine H1 receptor

was confirmed by RT-PCR. Interestingly, histamine pretreatment did not inhibit a

subsequential carbachol-induced [Ca2+]i rise without ‘heterologous desensitization’.

Chlorpheniramine inhibited a carbachol-induced [Ca2+]i increase at a 100-fold greater

concentration than histamine receptor antagonism, whereas astemizole and cetrizine

showed more than 1000-fold difference, which explains in part the xerostomia-inducing

potency among the anti-histamines. Notably, histamine resulted in translocation of

aquaporin-5 to the plasma membrane in human submandibular gland cells and green

fluorescent protein (GFP)-tagged aquaporin-5 expressing HSG cells. We found that

histidine decarboxylase and the histamine H1 receptor are broadly distributed in

submandibular gland cells, whereas choline acetyltransferase is localized only at the

parasympathetic terminals. Our results suggest that human salivary gland cells express


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histamine H1 receptors and histamine synthesizing enzymes, revealing the cellular

mechanism of anti-histamine-induced xerostomia.


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Introduction

The salivary gland is an exocrine gland that secretes saliva (Turner and Sugiya, 2002). The

saliva secretion requires the coordinated activity of membrane transporters, receptors, and

ion channels. Close communication between the salivary gland and neurons significantly

modulates salivary secretion. Neuronal modulation of the salivary gland is mediated by

neurotransmitter release from neurons and receptor activation in salivary gland cells. The

most important neuronal modulation is muscarinic signaling from parasympathetic neurons

(Turner and Sugiya, 2002; Li et al., 2006). In salivary gland acinar cells, acetylcholine

activates muscarinic M3 receptors, the Gq/11 protein and phospholipase C, increasing

cytosolic Ca2+ levels, which is the most critical signal molecule for salivary secretion

(Ambudkar, 2000). Animals lacking muscarinic M3 receptors (Matsui et al., 2000) or the

IP3 receptor (Futatsugi et al. 2005) show severely decreased salivation.

Xerostomia (dry mouth) is a pathological condition characterized by decreased salivary

secretion (Guggenheimer and Moore, 2003). Xerostomia usually results in the sensation of

a burning mouth, halitosis, and oral infection. Prolonged xerostomia induces rampant caries,

erosion and, finally, tooth loss. Saliva is especially important in host defense of the oral

cavity, thus loss of the salivary fluid membrane can increase the amount of pathological

bacteria, such as Streptococcus, and induce oral disease. For these reasons, modulation of

salivary secretion has been intensively studied. A series of neuronal inputs, which are


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triggered through activation of the autonomic nervous system by external stimulation (acid

or mechanical), delicately modulate salivary gland function.

Xerostomia can result from many conditions with salivary gland dysfunction, including

radiation therapy of salivary glands or Sjögren’s syndrome: an autoimmune disease

characterized by decreased salivary and lacrimal secretion. However, xerostomia is most

frequently experienced as a side effect of medication (Sreebny and Schwartz, 1997;

Guggenheimer and Moore, 2003). A series of extrinsic medicines decrease salivary

secretion without any obvious pathological changes in the salivary gland. Currently,

muscarinergic inhibitors are one of the few xerostomia-causing extrinsic medicines for

which the mechanism is fully understood. Anticholinergic drugs decrease salivation and

induce xerostomia by modulating receptor-mediated Ca2+ signaling in salivary gland cells.

The xerostomia-inducing mechanisms of many other medicines are unclear. One example is

anti-histamine medication; classically the histamine H1 receptor inhibitors. Frequently

prescribed for minor allergic reactions or the cold-like symptoms of allergic rhinitus, anti-

histamines are reported to cause xerostomia (Elad et al., 2006); however, the exact

mechanism for an anti-histamine-induced decrease in salivary secretion is still unknown.

The biological action of histamines is mediated by their binding to histamine receptors,

classified as H1, H2, H3, and H4 (Parsons and Ganellin, 2006). Even though the

xerostomia inducing effect of anti-histamines strongly suggests that histaminergic Ca2+

signaling is functional in human salivary glands, there is no solid evidence for the


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expression of any histamine receptors. The current consensus is that anti-histamine-induced

xerostomia comes from an anti-muscarinic effect; indeed, a number of studies have shown

that first-generation histamine H1 receptor antagonists possess anti-muscarinic activity

(Yasuda and Yasuda, 1999; Liu et al., 2006).

We hypothesized that the xerostomic effects of anti-histamines are mostly due to histamine

H1 receptor blocking. To test this hypothesis, we sought to elucidate the action of

histamine and anti-histamines in the human salivary glands. Here we report intracellular

Ca2+ increase and subsequent aquaporin translocation by histamine H1 receptor activation

in human submandibular glands.


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Methods

Materials

Histamine, chlorpheniramine, astemizole, ranitidine, thioperamide, carbachol,

diaminobenzidine, paraformaldehyde were purchased from Sigma (St. Louis, MO, USA).

Fura-2-acetoxymethyl ester (Fura-2-AM) was obtained from Molecular Probes (Eugene,

OR, USA). Aquaporin-5 (AQP-5) antibody and secondary donkey anti-goat IgG-

horseradish peroxidase (HRP) were purchased from Santa Cruz Biotechnology (Santa Cruz,

CA, USA). Anti-rabbit Cy3 and normal donkey serum were obtained from Jackson

ImmunoResearch Lab (West Grove, PA, USA). Biotinylated anti-Goat IgG, and Vectastain

Elite ABC Kit were purchased from Vector Laboratories (Burlingame, CA, USA). Anti-

rabbit human histidine decarboxylase antibodies, and anti-rabbit human histamine receptor

H1 antibodies were purchased from Progen Biotechnik GmBH (Heidelberg, Germany) and

Acris Antibodies GmBH (Herford, Germany), respectively. Goat anti-choline

acetyltransferase (anti-ChAT) antibodies were purchased from Chemicon International Inc.

(Temecula, CA, USA). Collagenase P was purchased from Roche Molecular Biochemicals.

Modified Eagle’s Medium, bovine calf serum, and penicillin/streptomycin were purchased

from GIBCO (Grand Island, NY, USA). [3H]cAMP was purchased from NEN (Boston,

MA, USA). Lipofectamine 2000 was obtained from Invitrogen (Carlsbad, CA, USA).

Polyethylene glycol 400 diesterate was purchased from Polyscience (Warrington, PA,

USA).


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Cell Preparation

The dissociation of human submandibular salivary gland cells was performed as previously

described (Choi et al., 2006). Pieces of human submandibular glands were surgically

removed from 12 oral cancer patients, who had provided informed consent. The patients

included males and females ranging from 38 to 69 yrs old. The glands did not contain

atypical cells when assessed histologically. The tissues were quickly removed and finely

minced in a physiological salt solution (containing [in mM]: 135 NaCl, 5.4 KCl, 1.2 CaCl2,

0.8 MgSO4, 0.33 NaH2PO4, 0.4 KH2PO4, 10 glucose, 2 glutamine, and 20 HEPES adjusted

to pH 7.4 with NaOH), supplemented with 10 mM sodium pyruvate, 0.02% trypsin

inhibitor, and 0.1% bovine serum albumin. The cells were then digested in the same

solution containing collagenase P (0.3 mg/7.5 ml) at 37°C for 75 min with continuous

agitation. The prepared acinar cells were re-suspended in physiological salt solution

containing 0.1% bovine serum albumin. This study was performed according to the

guidelines for experimental procedures found in the Declaration of Helsinki, the World

Medical Association, and this study was approved by the Institutional Review Board

(CRI06002) of Seoul National University Dental Hospital. The HSG cell line was grown in

Modified Eagle’s Medium supplemented with 10% (v/v) heat-inactivated bovine calf serum

and 1% (v/v) penicillin (5000 U/mL) + streptomycin (5000 µg/mL) solution. The cells were

cultured in a humidified atmosphere of 95% air and 5% CO2. The culture medium was

changed every 2 days, and the cells were subcultured weekly.


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[Ca2+]i Measurement.

Cytosolic free Ca2+ concentration ([Ca2+]i) was determined with the fluorescent Ca2+

indicator fura-2/AM as previously described (Choi et al., 2001). Briefly, a cell suspension

was incubated with fresh Modified Eagle’s Medium medium containing fura-2/AM (4 µM)

for 40 min at 37°C with continuous stirring. The cells were then washed with a HEPES-

buffered solution (140 mM NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 10 mM glucose,

and 10 mM HEPES, pH adjusted to 7.4 with NaOH) or Locke's solution (NaCl, 154 mM;

KCl, 5.6 mM; MgCl2, 1.2 mM; CaCl2, 2.2 mM; HEPES, 5.0 mM; glucose, 10 mM, pH 7.4

with NaOH) and left at room temperature until use. Sulfinpyrazone (250 µM) was added to

all solutions to prevent dye leakage. For the dissociated human salivary gland cells were

transferred to a poly-L-lysine-coated 25 mm coverslips for cell attachment for 40 min in

incubator, and the coverslips were mounted onto the chamber. Changes in fluorescence

ratio were monitored by Ca2+ imaging machine with MetaFlour software (Molecular

Devices, Sunnyvale, CA, USA) with dual excitation at 340 and 380 nm and emission at 500

nm. Experiments with suspended HSG cells were performed with the

spectrofluorophotometer. In this case, calibration of the fluorescent signal in terms of

[Ca2+]i was performed using the following equation:

[Ca2+]i = KD[(R - Rmin)/(Rmax - R)](Sf2/Sb2)

where R is the ratio of fluorescence emitted by excitation at 340 and 380 nm. Sf2 and Sb2

are the proportionality coefficients at 380 nm excitation of Ca2+-free fura-2 and Ca2+-

saturated fura-2, respectively. In order to obtain Rmin, the fluorescence ratios were

measured after adding 4 mM EGTA, 30 mM Trizma base, and 0.1% Triton X-100 to the


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cell suspension. To obtain Rmax, the cell suspension was then treated with CaCl2 at a final

concentration of 4 mM, and the fluorescence ratio measured.

[3H]cAMP Measurement.

The cAMP concentration in the cells was determined by a [3H]cAMP competition assay for

binding to the cAMP binding protein (Choi et al., 2001). To determine cAMP production,

cells were harvested and washed with Locke’s solution (154 mM NaCl, 5.6 mM KCl, 1.2

mM MgCl2, 2.2 mM CaCl2, 5 mM HEPES, 10 mM glucose, pH 7.4). The cells were then

preincubated with Locke’s solution containing 1 mM isobutylmethyl xanthine (IBMX) to

inhibit phosphodiesterase. IBMX was also added to the stimulating buffer with agonists.

After stimulation for 15 min at 37oC, the reaction was terminated by addition of twice the

volume ice-cold absolute ethanol. Cells were then incubated for 2 hr at -20 oC to extract the

cAMP. The cells in ethanol were centrifuged for 10 min at 4 oC at 10,000 x g. The

supernatant was evaporated and the residues dissolved in TE buffer (0.2 ml Tris-HCl, 4

mM EDTA, pH 7.5). Fifty μl sample solution was employed in the cAMP assay. This assay

is based on competition between [3H]cAMP and unlabelled cAMP in the sample for a crude

cAMP-binding protein prepared from bovine adrenal gland. Free [3H]cAMP is adsorbed by

charcoal and removed by centrifugation, and bound [3H]cAMP in the supernatant is

determined by liquid scintillation counting. Each unknown sample was incubated with 50

μl [3H]-labeled cAMP (5 μCi) and 100 μl binding protein for 2 hr at 4oC. The protein-

bound cAMP and unbound cAMP were separated by adsorbing free cAMP onto charcoal

(100 μl), followed by centrifugation at 12,000 x g at 4oC. Two hundred μl supernatant was


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placed in an Eppendorf tube containing 1.2 ml scintillation cocktail to measure the

radioactivity. The cAMP concentration in the sample was determined based on a standard

curve and expressed as pmol/cell number.

Reverse Transcription Polymerase Chain Reaction (RT-PCR)

Total RNA was isolated from primary culture human submandibular cells. The cDNA was

synthesized from 3 μg of total RNA by incubating the RNA for 1 hr at 37 °C in a reaction

mixture containing 0.5 μg of oligo (dT)15, 0.5 mM dNTP mix, 1X first-strand buffer,

RNase inhibitor (20 units), 5 mM dithiothreitol, and Moloney Murine Leukemia Virus

reverse transcriptase (200 units). Histamine receptor mRNA expression was measured

using the following PCR reaction conditions (Matsubara et al., 2005): initial denaturing at

94 °C for 3min, denaturing at 94 °C for 30 sec, annealing at 60 °C for 30 sec, and extension

at 72 °C for 30 sec. A total of 35 reaction cycles were performed. The amplified DNA

products were separated by electrophoresis on a 1.5 % agarose gel. PCR Primers used for

RT-PCR analysis of human salivary gland tissues were as follows: Human GAPDH, 5’-

ACC ACA GTC CAT GCC ATC AC-3’ and 5’-TCC ACC ACC CTG TTG CTG TA-3’;

human H1 receptor, 5’-GAC TGT GTA GCC GTC AAC CGG A-3’ and 5’-TGA AGG

CTG CCA TGA TAA AAC C-3’; human H2 receptor, 5’- TCG TGT CCT TGG CTA TCA

C-3’ and 5’-CTT TGC TGG TCT CGT TCC T-3’; human H3 receptor, 5’-TCA GCT ACG

ACC GCT TCC TGT CGG TCA C-3’ and 5’-TTG AGT GAG CGC GGC CTC TCA GTG

CCC C-3’; human H4 receptor, 5’-GAA TTG TCT GGC TGG ATT AAT TTG CTA ATT

TG-3’ and 5’-AAG AAT GAT GTG ATG GCA AGG ATG TAC C-3’.


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Construction of AQP-5 and Western blotting

Briefly, total RNA was isolated from primary culture human submandibular salivary gland

cells and full-length cDNA extracted from it was amplified with reverse-transcriptase PCR

(RT-PCR). PCR utilized 5’-GAATTCTATGAAGAAGGAGGTGTGCTCC as the

upstream primer and 5’-GGTACCGCGGGTGGTCAGCTCCATGGT as the downstream

primer (introduced EcoRI and KpnI restriction site are underlined). The 798 bp human

AQP-5 gene was then cloned into pEGFP-C1. HSG cells were transiently transfected with

the AQP-5 gene using LipofectAMINE 2000 (Invitrogen). AQP-5 transfected HSG cells

were grown in a 60-mm dish and incubated with carbachol or histamine for 15 min at 37°C.

The apical membrane fraction was obtained as previously described (Ishikawa et al., 2000).

After electrophoresis in an SDS-PAGE (12 %) gel, the protein was transferred to a

nitrocellulose membrane. After blocking, the membrane was incubated with anti-AQP-5 as

the primary antibody. The membrane was then washed and incubated with donkey anti-goat

IgG-HRP. The membrane was finally subjected to an electrochemiluminescence assay.

Immunofluorescence and Confocal Microscopy

GFP-tagged-AQP-5 transfected cells were plated on glass coverslips before experiments

were performed. Cells were treated with carbachol and histamine for 15 min. Cells were

then visualized for fluorescence using a differential interference fluorescence microscope

(Zeiss).


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Immunohistochemistry

Human submandibular gland tissue samples were fixed with 4 % paraformaldehyde in 0.1

M phosphate buffer (pH 7.2) for 4-5 h, and then embedded in Polyethylene glycol 400

diesterate wax after dehydration in graded alcohols. Eight μm thick tissue sections were

obtained using a microtome (Leica RM 2135), deposited on gelatin-coated slides, and air-

dried. After de-waxing with xylene followed by hydration, the tissue sections were treated

with 1.5 % hydrogen peroxide for 30 min and washed twice for 15 min in 0.1 M sodium

phosphate buffered saline (PBS). Tissue sections were treated with normal donkey serum

(1:50 dilution) for 30 min and incubated with anti-rabbit human histidine decarboxylase

antibodies (1:100 dilution), anti-rabbit human histamine receptor H1 antibodies (1:100

dilution) or goat anti-choline acetyltransferase antibodies (1:500 dilution) overnight. After

washing twice in PBS, the sections incubated with anti-histidine decarboxylase or anti-

histamine H1 receptor antibodies were treated with α-rabbit Cy3 (1:200 dilution for anti-

histidine decarboxylase, 1:1000 dilution for anti-histamine H1 receptor) for 1 h, and then

observed under confocal laser scanning microscope (Carl Zeiss, LSM510 META). Sections

were incubated with anti-ChAT antibodies were treated with biotinylated anti-Goat IgG

(1:100 dilution) for 1 h, then bound secondary antibodies were amplified with the

Vectastain Elite ABC Kit. Antibody complexes were visualized with 0.05 %

diaminobenzidine for 3 min. Sections were washed twice with PBS, counterstained with

hematoxylin, dehydrated, and coverslipped with permount.


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Data Analysis

All quantitative data are expressed as mean ± SEM. We calculated the half-maximal

effective concentration (EC50) and half-maximal inhibitory concentration (IC50) with the

Microcal Origin program. Differences were determined by one-way analysis of variance

and considered to be significant only when P < 0.05.


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Results

Functional Histamine H1 Receptor in Human Salivary Gland Cells and HSG Cells

We examined histamine-induced cellular responses in human submandibular gland cells.

Throughout our experiments, we used primary cultured or dissociated human

submandibular gland cells, as well as the HSG cell line due to the limited supply of human

normal submandibular tissue. Histamine (1 µM and 100 µM) triggered a [Ca2+]i rise in

fura-2-loaded primary cultured human submandibular gland cells (Fig. 1A) and HSG cells

(Fig. 1C). Histamine elevated [Ca2+]i in a concentration-dependent manner proportional to

the histamine concentration between 10-7 M and 10-4 M (supramaximal concentration). The

half-maximal effective concentrations (EC50) were 38.5 ± 3.0 µM in primary cultured

human submandibular gland cells (Fig. 1C) and 11.8 ± 0.6 µM in HSG cells (Fig. 1D). It

reached a peak level within a few seconds and fell back to the basal level in 200 sec.

To identify the histamine receptor subtype expressed in HSG cells, subtype-specific

antagonists were tested. It is generally accepted that chlorpheniramine, cetrizine,

astemizole, and loratadine inhibit the histamine H1 receptor, ranitidine blocks the histamine

H2, and thioperamide inhibits H3 and H4 receptors (Liu et al., 2001; Parsons and Ganellin,

2006). Chlorpheniramine inhibited the histamine-induced [Ca2+]i increase (Fig. 2A) in a

concentration-dependent manner with an IC50 of 128 ± 16 nM (Fig. 2B). Cetrizine also

inhibited the histamine-evoked Ca2+ response (Fig. 2C) in a concentration-dependent

manner with an IC50 of 141 ± 17 nM (Fig. 2D). Astemizole and loratadine also inhibited

histamine-induced [Ca2+]i increase with IC50 of 73.6 ± 4.5 nM (Fig. 2E-2F) and 64.2 ± 5.3


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nM (Fig. 2G-2H), respectively. Ranitidine (30 μM) and thioperamide (1 μM) did not affect

the histamine-induced [Ca2+]i increase (Fig. 3A), despite application at concentrations

known to be sufficient for receptor-specific blockade. We monitored histamine-induced

cAMP production to investigate the existence of adenylyl cyclase-linked histamine

receptors (Kim et al., 2006). Forskolin (direct adenylyl cyclase activator) and isoproterenol

(β-adrenergic agonist) successfully increased cAMP levels; however, histamine failed to

elevate cAMP in HSG cells (Fig. 3B). To detect histamine receptor subtype mRNA

expression in the human submandibular gland, we designed specific primers based on

histamine receptor subtype sequences. The specificity of the primers was confirmed by

sequencing the gene products and by amplifying full-length human H1, H2, H3, and H4

receptor cDNAs. Only the H1 receptor was expressed in the dissociated human salivary

gland cells (Fig. 3C) and HSG cells (Fig. 3D).

Histamine Receptor Signaling Is Independent from Muscarinic Receptor Signaling

Increases in cytosolic Ca2+ play a critical role in saliva secretion (Ambudkar, 2000).

Intracellular Ca2+ activates a series of membrane transporters, ion channels, and aquaporins

in salivary gland acinar cells, which transport water and electrolytes to make primary saliva.

Salivary gland cells have cytosolic Ca2+-mobilizing muscarinic M3 receptors in their

plasma membrane and receive signals from parasympathetic neurons (Nakamura et al.,

2004). We confirmed these inputs by showing a carbachol-induced [Ca2+]i increase (Fig. 4).

We examined the possibility that histamine and muscarinic receptors affect each other in

terms of Ca2+ signaling. Interestingly, pretreatment with histamine did not dramatically


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decrease the carbachol-evoked [Ca2+]i increase in the dissociated human submandibular

gland cells (Fig. 4A). Similar results were seen in HSG cells (Fig. 4B). Also pretreatment

with carbachol did not inhibit the histamine-evoked [Ca2+]i increase (Fig. 4C). These data

suggest that histamine and muscarinic receptors function independently of one another with

no overlapping Ca2+ signaling pathway. We compared the carbachol- and histamine-evoked

[Ca2+]i increases at the subcellular level, but could not find a difference in localized [Ca2+]i

increase (Supplementary Fig. 1).

Interestingly, anti-histamine-induced xerostomia has been explained by blocking

muscarinic rather than histamine receptors in human salivary glands (Sreebny and Schwartz,

1997; Guggenheimer and Moore, 2003). To clarify this possibility, we tested the effect of

histamine H1 inhibitors on the carbachol-induced [Ca2+]i increase in HSG cells.

Chlorpheniramine - one of the first-generation anti-histamines - inhibited the carbachol-

induced [Ca2+]i increase at high concentrations (Fig. 5A). Inhibition occurred in a

concentration-dependent manner with an IC50 of 43.9 ± 9.2 μM: 100-fold greater than the

IC50 at histamine H1 receptors (Fig. 5B). We next monitored the carbachol-induced [Ca2+]i

response in the presence of three second-generation anti-histamines. Cetrizine (300 μM, Fig.

5C) and astemizole (1 μM, Fig. 5E) did not affect, or only mildly inhibited, the muscarinic

Ca2+ response. Relatively loratadine (1 μM) showed more potent inhibition (Fig. 4G).

Astemizole and cetrizine at higher concentrations showed a continuous increase in

cytosolic Ca2+ level without any receptor-mediated responses (data not shown). Their


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predicted IC50 at muscarinic receptors were >100 μM and therefore 1000-fold higher than

the inhibitory concentration at the histamine H1 receptor.

Histamine H1 Receptor Induces the Translocation of Aquaporin-5

Aquaporin-5 (AQP-5) is an important channel protein that regulates water movement in the

salivary gland (Delporte and Steinfeld, 2006). To determine the role of histamine H1

receptors in salivary secretion, we examined whether histamine causes GFP-tagged AQP-5

to translocate to the plasma membrane in transfected HSG cells. Cells treated with

carbachol showed higher fluorescence at the plasma membrane than in the cytosol (Fig.

6Ab, 6Ae), whereas cells treated with vehicle showed similar fluorescence intensities in

both the cytosol and plasma membrane (Fig. 6Aa, 6Ad). Notably, we identified that

histamine also caused higher fluorescence intensity in the plasma membrane (Fig. 6Ac,

6Af). These results were confirmed using western blotting for AQP-5 in the apical plasma

membrane and intracellular membranes of HSG and primary cultured human

submandibular gland cells. Both carbachol and histamine caused AQP-5 to translocate to

the apical plasma membrane in HSG cells (Fig. 6B) and human submandibular glands (Fig.

6C). These results suggest that the histamine H1 receptor may also mediate AQP-5

translocation to the plasma membrane in human submandibular glands.

Histamine H1 Receptors and Histamine Histidine Decarboxylase are Colocalized in

Human Salivary Gland Cells


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In order to elucidate the site(s) of histamine secretion and action, we labeled histamine H1

receptors with an H1 receptor-specific antibody. As shown in Fig 7, the histamine H1

receptor was seen distributed across most of the human submandibular gland including the

acinar and ducts. Interestingly, histidine decarboxylase (histamine synthesizing enzyme)

showed a similar distribution to labeled H1 receptors. In contrast, the distribution of choline

acetyltransferase (acetylcholine synthesizing enzyme; Anderson and Garrett, 1994) -

although highly localized - was distinct from the H1 receptor and histidine decarboxylase

distributions. The difference between histidine decarboxylase and choline acetyltransferase

distributions suggests that histamine is not secreted from the acetylcholine-releasing

parasympathetic terminals but from the salivary gland itself; therefore, histamine is likely

to act in an autocrine manner.


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Discussion

In this study, we have shown for the first time that histamine H1 receptors mediate Ca2+

signaling in human submandibular gland cells and we suggest that the histamine H1

receptor signaling is a novel target mechanism in addition to the classical mechanism of

anti-histamine-induced xerostomia. We report that (a) the histamine H1 receptor, expressed

exclusively in human submandibular gland cells, increases cytosolic Ca2+ without cAMP

production, (b) histamine stimulation results in aquaporin translocation, and (c) histamine

receptors coexist with histidine decarboxylase in gland cells.

Histamine has been reported to modulate salivary flow in humans and dogs (Shimizu and

Taira, 1980); however, there has been no evidence concerning histamine H1 receptors in

human salivary glands, and the currently accepted hypothesis is that anti-histamine-induced

xerostomia is due to muscarinic receptor antagonism. This hypothesis was generally

applied to classical anti-histamines (for instance, chlorpheniramine and promethazine)

which have been shown to inhibit muscarinic receptors (Yasuda and Yasuda, 1999; Liu et

al., 2006; Brown and Eckberg, 1997; Shelton and McCarthy, 2000). However, recently

developed ‘second-generation’ anti-histamines, such as astemizole and fexofenadine, also

induce xerostomia (Wilson et al., 1987; Sreebny and Schwartz, 1997; Elad et al., 2006),

despite their extremely low affinity for muscarinic receptors (Laduron et al., 1982; Liu et

al., 2006). These observations suggest that xerostomia is not solely related to muscarinic

receptor inhibition.


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To address this question, we attempted to make clear the relationship between muscarinic

and histaminergic signaling in human submandibular gland cells. First, we checked for

crosstalk between the histamine and muscarinic receptors. If salivary gland cells express

histamine receptors, both histamine and muscarinic signaling may modulate cytosolic

[Ca2+]i and salivary secretion. In general, ‘heterologous desensitization’ refers to a decrease

in signaling of a certain receptor by another neurotransmitter that shares the same

downstream signal transduction pathway. In our report, the carbachol-induced [Ca2+]i

increase was not affected by histamine-induced [Ca2+]i increase, thereby ruling out

heterologous desensitization between muscarinic- and histamine-mediated Ca2+ signaling

(Fig. 4). These results suggest that histamine can independently induce Ca2+ signaling, and

possibly induce salivary secretion, in the absence of muscarinic stimulation.

Second, we tested the affinity of anti-histamines for muscarinic receptors. We monitored

the effect of chlorpheniramine - a typical first-generation anti-histamine - on muscarinic

signaling. As in a previous report (Shelton and McCarthy, 2000), we confirmed that

chlorpheniramine significantly inhibited the carbachol-induced [Ca2+]i increase (Fig. 5).

The inhibitory effect, however, was obtained with a very high chlorpheniramine

concentration: about 300-fold greater than the effective concentration against histamine H1

receptors. In clinical applications, the chlorpheniramine serum level reaches approximately

700 nM; this concentration is enough to inhibit histamine H1 receptors, but would hardly

be effective at muscarinic M3 receptors (Fig. 5B). We suggest that the salivary histamine


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H1 receptor is a more probable and feasible candidate for anti-histamine-induced

xerostomia. Most first-generation histamine H1 receptor antagonists exhibit serious

xerostomia, whereas some of the second-generation histamine H1 receptor antagonists

show only a mild xerostomic side effect. Even though our results successfully showed a

primary xerostomic inducing mechanism of anti-histamine, we could not clearly answer the

reason for the difference in the xerostomic potency among H1 receptor antagonists.

Theoretically, there should be no difference between the two generations of anti-histamines

if histamine signaling is the only xerostomic mechanism at play. Conversely, if muscarinic

signaling is the only xerostomic target for the antihistamines, then it would be unlikely for

the second-generation histamine H1 receptors blockers (such as astemizole) to cause

xerostomia. According to our results in human salivary glands, neither hypothesis is

completely true. We found a relatively larger difference between inhibitory concentration

for histamine receptors and muscarinic receptors for astemizole, loratadine, and cetrizine

than the inhibitory concentrations recorded for chlorpheniramine. We found a large

difference between inhibitory concentration for histamine receptors and muscarinic

receptors for astemizole and cetrizine. The IC50 values for chlorpheniramine, on the other

hand, revealed a relatively closer affinity for both receptors. From this we can conclude that

the difference in xerostomia induction potency between first- and second-generation

histamine H1 receptor antagonists may be ‘in part’ due to their different affinities for

muscarinic receptors. In fact, we believe that the difference in xerostomic potency might

not only be caused by inhibition of muscarinic signaling, but may also be due to the


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multiple effects of blocking histamine H1 receptors, such as histaminergic signaling

inhibition, sedation, and so on.

Third, we performed experiments to determine whether histamine activates cytosolic

machinery related to salivary secretion in a manner similar to muscarinic receptors. In the

salivary gland, AQP-5, one of thirteen mammalian AQPs, modulates water secretion to

produce primary saliva (Ishikawa et al., 2005). Sjögren’s syndrome is characterized by a

xerostomia that results from defective AQP-5 translocation to the luminal side (Tsubota et

al., 2001). Translocation of AQP protein due to Ca2+ signaling is therefore a critical step in

fluid secretion. In our experiments, histamine stimulation resulted in translocation of AQP-

5 to the luminal membrane, similar to the effect of carbachol (Fig. 6). It must be

emphasized that altered AQP-5 translocation provides only indirect evidence for a role in

salivary gland water secretion in vivo; however, an in vitro approach using human cells is

preferable to the treatment of humans with histamine receptor agonists, which would

inevitably induce severe allergy and hypersensitivity responses. Our results suggest that

both histamine and parasympathetic acetylcholine can potentially control salivation.

Among histamine receptors, H2 receptor activation is known to produce cAMP and H1

receptors are linked to Ca2+ mobilization. In rat parotid tissues, histamine is reported to

stimulate H2 receptors and induce amylase secretion (Eguchi et al., 1998). Salivary

secretion in cats and rats is inhibited by histamine H2 receptor antagonists (Stanovnik and

Erjavec, 1983; Sim and Ang, 1985). Adenylyl cyclase-linked G protein-coupled receptor


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activation serves an important role in salivary protein secretion via intracellular cAMP

production (Yamada et al., 2006). In other tissues, such as collecting duct of kidney, the

increased cAMP could feasibly translocate AQP and trigger water secretion (Valenti et al,

2005). Our results in human submandibular gland cells, however, do not support the

existence of histamine H2 receptors (Fig 3); a finding perhaps explained by species

differences. Previous studies have shown differences in the composition of saliva between

humans and other species, as well as the membrane transporter profiles of saliva

components (Bardow et al., 2000). Due to the difficulties in fitting animal models to human

physiology for certain features of salivary function, our results serve to enhance our

understanding of the human salivation mechanism and enable better xerostomia treatment.

We next examined the histamine source and target to determine the origin of salivary

modulation. During inflammation, histamine may be produced by factors circulating in the

blood stream, for example platelets. In ‘normal’ conditions, however, neurons are likely to

be the principal source of histamine (Haas and Panula, 2003). Histamine was reported to be

released together with acetylcholine from cardiac parasympathetic neurons, which co-stain

for histidine decarboxylase and choline acetyltransferase (Singh et al., 1999). In order to

address histamine secretion from parasympathetic terminals, we compared the choline

acetyltransferase and histidine decarboxylase distributions in the human submandibular

gland. Our results showed comparable distributions of histidine decarboxylase and

histamine H1 receptor, but no overlap with choline acetyltransferase. These results strongly

suggest that the parasympathetic terminals in human submandibular glands are unlikely to


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release histamine. Instead, histamine may be release from the salivary gland itself and act in

an autocrine manner.

Acetylcholine is secreted from parasympathetic terminals and diffuses about 100 nm before

stimulating receptors on the salivary acinar cell membrane (Konttinen et al., 2006).

Sympathetic neurons, with limited axon terminals to modulate a target organ, possess

varicosities to increase the number of synaptic contacts (Konttinen et al., 1996); however,

such varicosities are relatively less developed in parasympathetic neurons (Arvidsson et al.,

1997). In general, synthesized histamine is stored in cytosolic vesicles and secreted by

intracellular Ca2+-triggered vesicular membrane fusion (Puri and Roche, 2008). Salivary

gland cells also have the machinery for secretory vesicle fusion (Wang et al. 2007). We

suggest the hypothesis that cholinergic release from parasympathetic terminals activates

muscarinic Ca2+ signaling in salivary gland cells. This triggers the release of histamine,

which activates neighboring cells expressing H1 receptors. The cascade will spread the

signal over the entire glands. Although further study is necessary to confirm this hypothesis,

our data provides a possible role for histamine in the propagation of parasympathetic

signals to the salivary gland in spite of its localized and limited cholinergic input.

Currently, the significance of H1 receptor expression to overall human salivary fluid

secretion is unknown and the possibility of other interactions between anti-histamines and

muscarinic signaling still remain. However, taken together, our results show that histamine

is a possible secretory signal, and suggest that anti-histamines directly inhibit histamine H1


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receptors in human submandibular glands. As salivary Ca2+ signaling is correlated with

saliva secretion, these results may contribute to understanding the cellular mechanism of

anti-histamine-evoked xerostomia.


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Footnotes

This work was supported by the Korea Science & Engineering Foundation grant [R13-

2008-008-01001-0] through the Oromaxillofacial Dysfunction Research Center for the

Elderly at Seoul National University.

J.-H.K., S.-H.P. contributed equally to this work.

We thank Dr. Alexander Davies for his English correction of this manuscript.


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Legends for Figures

Fig. 1. The effect of histamine on human submandibular gland cells and HSG cells.

(A) Fura-2-loaded dissociated human submandibular gland cells were treated with 1 μM

(gray trace) or 100 μM histamine (black trace), and changes in F340/F380 were monitored.

Typical Ca2+ transients from more than three separate experiments are presented. (B)

Dissociated human submandibular gland cells were challenged with the indicated histamine

concentrations. The peak heights of changes in the fluorescence ratio were monitored. Each

point represents mean ± SEM. (C) Fura-2-loaded HSG cells were treated with 1 μM (gray)

or 100 μM histamine (black), and the increase in cytosolic Ca2+ concentration ([Ca2+]i) was

monitored. Typical Ca2+ transients from more than four separate experiments are presented.

The results were reproducible. (D) HSG cells were challenged with the indicated histamine

concentrations. The peak height of Ca2+ elevation was monitored. Each point represents

mean ± SEM. All results were reproducible.

Fig. 2. Histamine H1 receptor antagonists inhibit the histamine-induced [Ca2+]i

increase in HSG cells. Fura-2-loaded cells were treated with 100 μM histamine with (gray

trace) or without (black trace) the preincubation of 10 μM chlorpheniramine (A), 1 μM

cetrizine (C) 300 nM astemizole (E), or 300 nM loratadine (G). Typical Ca2+ transients

from more than five separate experiments are presented. Cells were preincubated with the

indicated concentrations of chlorpheniramine (B), cetrizine (D), astemizole (F), or

loratadine (H), and subsequently treated with 100 μM histamine. The peak height of Ca2+


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elevation was monitored. Each point represents mean ± SEM. The experiments were

performed four times independently. All results were reproducible.

Fig. 3. The histamine H1 receptor is expressed in HSG cells. (A) Fura-2-loaded cells

were preincubated with 1 μM chlorpheniramine (CPR), 1 μM astemizole (AST), 30 μM

ranitidine (RAN), and 1 μM thioperamide (THIO) and subsequently treated with 100 μM

histamine. The peak height of histamine-induced Ca2+ elevation was monitored. Each point

represents mean ± SEM. The experiments were performed three or more times

independently. (B) Cells were stimulated with 3 μM forskolin, 100 μM histamine, or 1 μM

isoproterenol in the presence of 1 mM IBMX for 15 minutes; cAMP production was then

monitored. The cAMP levels were measured as described in Materials and Methods. The

relative cAMP productions are depicted as percent of forskolin-induced cAMP production

with the mean ± SEM of triplicate assays. The data are representative of three separate

experiments. (C) RNA extracted from HSG cells was reverse-transcribed into cDNA and

amplification reactions were performed using histamine H1, H2, H3, and H4 receptor

specific primers and Pfu polymerases described in Materials and Methods. GAPDH was

used as an internal loading control. All results were reproducible. ** P < 0.01.

Fig. 4. Histamine and muscarine receptors do not share Ca2+ signaling in HSG

cells. (A) Fura-2-loaded primary cultured human submandibular gland cells treated

with 100-μM histamine with (black trace) or without (gray trace) the pretreatment of

100 μM carbachol. (B) Fura-2-loaded HSG cells were treated 100 μM carbachol with


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(black trace) or without (gray trace) the pretreatment of 100μM histamine. (C) Fura-2-

loaded HSG cells were treated 100 μM histamine with (black trace) or without (gray

trace) the pretreatment of 100 μM carbachol. All presented results are typical Ca2+

transients from three or more separate experiments.

Fig. 5. Histamine H1 receptor antagonists inhibit the carbachol-induced [Ca2+]i

increase at high concentrations in HSG cells. Fura-2-loaded cells were treated with 100

μM carbachol with (gray trace) or without (black trace) the preincubation of 10 μM

chlorpheniramine (A), 300 μM cetrizine (C) 1 μM astemizole (E), or 1 μM loratadine (G).

Typical Ca2+ transients from three or more separate experiments are presented. Cells were

preincubated with the indicated concentrations of chlorpheniramine (B), cetrizine (D),

astemizole (F), or loratadine (H), and subsequently treated with 100 μM carbachol. The

peak height of Ca2+ elevation was monitored. Each point represents mean ± SEM. Dotted

gray lines represent the effect of chlorpheniramine on ‘histamine’-induced [Ca2+]i increase

for comparison. The experiments were performed more than three times independently. All

results were reproducible.

Fig. 6. Effect of carbachol and histamine on AQP-5 expression in the intracellular

membrane and apical membrane fractions of HSG cells and primary cultured human

submandibular gland cells. (A) GFP-tagged-AQP-5 transfected HSG cells were

preincubated in the absence (Aa) or presence of 100 μM carbachol (Ab) and 100 μM

histamine (Ac) for 15 min. Spatial quantification (Ad, Ae, Af) was performed along a path


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across the plasma membrane, indicated by a white line in (Aa), (Ab) and (Ac), respectively.

Yellow Scale bar, 20 µm. (B, C) Intracellular membrane and apical membrane of AQP-5

transfected HSG cells (B) and primary cultured human submandibular glands (C) incubated

in carbachol and histamine for 15 min were subjected to immunoblot with AQP-5 antibody.

ICM: intracellular membrane, APM: apical membrane.

Fig. 7. The histamine H1 receptor and histidine decarboxylase in human

submandibular glands. Human submandibular gland tissue samples were fixed, embedded,

de-waxed, and incubated with anti-rabbit human histidine decarboylase antibodies, anti-

rabbit human histamine receptor H1 antibodies, or goat anti-choline acetyltransferase

antibodies. (A) Negative control (x100), (B) histidine decarboxylase (x100), (C) histamine

H1 receptor (x100), (D) histidine decarboxylase (x400), (E) histamine H1 receptor (x400),

(F, G) choline acetyltransferase (x400).


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300

500100 sec

1 μM100 μM

100

[Ca2+

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Histamine

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CarbacholHistamine

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