Degradation of submandibular gland AQP5 by parasympathetic denervation of chorda tympani and its...

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The degradation of submandibular gland AQP5 by parasympathetic denervation

of chorda tympani and its recovery by cevimeline,

an M3 muscarinic receptor agonist§

Xuefei Li,1,2 Ahmad Azlina,3 Mileva Ratko Karabasil,3 Nunuk Purwanti,3 Takahiro Hasegawa,

Chenjuan Yao, Tetsuya Akamatsu, and Kazuo Hosoi4

Department of Molecular Oral Physiology, Institute of Health Biosciences,

The University of Tokushima Graduate School,

3-Kuramoto-cho, Tokushima-shi,

Tokushima, 770-8504 Japan

Running title: Autonomic regulation of salivary gland AQP5

Address correspondence to: Prof. Kazuo Hosoi, Department of Molecular Oral Physiology, Institute of

Health Biosciences, The University of Tokushima Graduate School, 3-15-18, Kuramoto-cho,

Tokushima-shi, Tokushima 770-8504 Japan

Phone: +81-88-633-7323; Fax: +81-88-633-7324; E-mail: [email protected]

of 46Articles in PresS. Am J Physiol Gastrointest Liver Physiol (May 1, 2008). doi:10.1152/ajpgi.00359.2007

Copyright © 2008 by the American Physiological Society.

mailto:[email protected]

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Footnotes:

§This work was supported in part by grants-in-aid for scientific research (Nos. 13671940 and 13671941)

from the Ministry of Education, Culture, Sports, Science, and Technology of Japan and by a grant from the

Japan China Medical Association.

1This work was submitted to The Graduate School of Dentistry, The University of Tokushima, as a part of

a dissertation for the Doctorate of Philosophy Degree.

2Supported by a scholarship from Rotary Yoneyama Memorial Foundation, Inc. Present address:

International Division, Otsuka Pharmaceutical Factory, Inc, 115 Tateiwa, Muya-cho, Naruto, Tokushima

772-8601, Japan

3Supported by a scholarship from the Ministry of Education, Culture, Sports, Science, and Technology of

Japan.

4To whom correspondence should be addressed: Department of Molecular Oral Physiology, Institute of

Health Biosciences, The University of Tokushima Graduate School, 3-18-15 Kuramoto-cho,

Tokushima-shi, Tokushima 770-8504, Japan. Tel: +81-88-633-7323; Fax: +81-88-633-7324; E-mail:

[email protected].

5Abbreviations used : AQP, aquaporin; AMC, 7-amino-4-methylcoumarin; CSPD, disodium

3-(4-methoxyspiro{1,2-dioxetane-3,2’-(5-chloro) tricyclo [3.3.1.13,7] decan}-4-yl) phenyl phosphate;

CSTD, cervical superior trunk denervation; CTD, chorda tympani denervation; DIG, digoxigenin; DMSO,

dimethyl sulfoxide; DTT, dithiothreitol; ECL, enhanced chemical luminescence; EDTA,

ethylenediaminetetraacetic acid; EGTA, O, O'-Bis(2-aminoethyl)ethylene-glycol-N, N, N’,

N’,-tetraacetic acid; HBSS, Hanks’ balanced salt solution; HEPES,

N-2-hydroxyethylpiperazine-N’-ethanesulfonic acid; MocAc-, methoxy coumarin-4-yl acetyl-; MOPS,

3-(N-morpholino)propanesulfonic acid; PBS, phosphate-buffered saline; PMSF, phenylmethylsulfonyl

fluoride; RT-PCR, reverse transcriptase-polymerase chain reaction; SDS-PAGE, sodium

dodecylsulfate-polyacrylamide gel electrophoresis; SMG, submandibular gland

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Abstract

By chorda tympani denervation (CTD; parasympathectomy), the aquaporin 5 (AQP5) but not AQP1

protein level in the rat submandibular gland (SMG) was significantly decreased, dropping to 37% of that

of the contralateral gland at 4 weeks. The protein levels of AQP5 and AQP1 were not significantly affected

by denervation of the cervical sympathetic trunk (CSTD; sympathectomy). Administration of cevimeline

hydrochloride, an M3 muscarinic receptor agonist (10 mg/kg for 7 days, p.o.), but not pilocarpine (0.3

mg/kg for 7 days, p.o.), recovered the AQP5 protein level reduced by CTD, and increased the AQP1

protein level above the control one. The mRNA level of AQP5 was scarcely affected by CTD and

cevimeline hydrochloride administration. Administration of chloroquine (50 mg/kg for 7 days, p.o.), a

denaturant of lysosomes, increased the AQP5 protein level reduced by CTD. An extract obtained from the

submandibular lysosomal fraction degraded the AQP5 protein in the total membrane fraction in vitro.

These results suggest the possible regulation of the AQP5 protein level in the SMG by the parasympathetic

nerves/M3 muscarinic receptor agonist and imply the involvement of lysosomal enzymes but not a

transcriptional mechanism in this regulation.

Key words: aquaporin-5, parasympathectomy, cevimeline, M3 muscarinic receptor agonist, salivary gland

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Introduction

Aquaporins (AQPs) are the family of water channel proteins distributed widely in nature, being found

in organisms from bacteria to animals. To date, 13 AQPs have been identified in various tissues of

mammals (1, 12-14, 17, 23). These proteins exist in the cell membrane and selectively transport water or

water and solutes such as glycerol and urea through the central pore of each AQP monomer in response to

osmotic gradients (2, 11, 16, 20, 39, 41, 51). AQPs have been divided into 2 major sub-groups based on

their permeability properties: AQP0, 1, 2, 4, 5, 6, and 8 are classically defined as water-selective channels,

whereas AQP3, 7, 9, and 10 are known as aquaglyceroporins (15), which are channels with less selectivity

and are permeated by water, glycerol, and other small solutes. Based on differences in physiological

function, each member of the AQP family appears to be distributed in particular tissues. For example,

AQP1 is a constitutively active channel (40) and has been identified in multiple tissues; whereas AQP2

expression is restricted to renal collecting duct cells and cells of the endolymphatic sac of the inner ear

(24), where it is regulated by vasopressin (10, 11, 36). AQP5 exists in cells of the submandibular, parotid,

sublingual, and lacrimal glands, as well as in the trachea, eye, distal lung, and duodenum but not in the

kidney or brain (37, 38, 41). Ma et al. (30) reported that AQP5 plays an important role in the secretion of

salivary fluid; i.e., pilocarpine-stimulated salivary fluid secretion in AQP5-knockout mice was reduced by

more than 60% compared with that by wild-type mice. Similar results were also found for AQP5 mutant

rats (35).

The hormonal regulation of AQPs has been well studied; e.g., AQP2 trafficking in the kidney collecting

tubular cells is triggered by short-term exposure to vasopressin, and expression of this aquaporin is

up-regulated by long-term exposure to this hormone (33, 42). On the other hand, the participation of the

autonomic nervous system in the regulation of AQP expression has not been studied much. The salivary

glands are innervated by both sympathetic and parasympathetic nerves (7). The submandibular (SMG) and

sublingual glands are innervated by the parasympathetic nerve derived from the superior salivatory nucleus

of the medulla oblongata, whereas the parasympathetic innervation of the parotid glands is from the

inferior salivatory nucleus. The sympathetic nerves that innervate the major salivary glands are from the

superior cervical ganglion, which lies in front of the second and third cervical vertebrae. In general,

parasympathetic stimulation induces a larger amount of salivary secretion than the sympathetic one. It is,

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therefore, of interest to examine how the expression and/or trafficking of AQPs are regulated by the

autonomic nervous system.

In the present study, we examined the effects of sympathectomy, parasympathectomy, and the

combination of parasympathectomy and administration of cevimeline hydrochloride on the AQP5

expression in the rat SMG. We found that the level of AQP5 protein in the SMG was positively regulated

by the parasympathetic nervous system and that such regulation was based on the alteration of activity of a

protein degradation pathway involving a lysosomal enzyme(s).

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Materials and Methods

Reagents

Anti-AQP5, anti-AQP1, anti-Na+,K+-ATPase α-subunit, and anti-kallikrein (anti-mK1) anti-sera were

prepared in our laboratory (25, 34, 45, 54). Aprotinin, agarose S, chloroquine diphosphate, hyaluronidase,

phenylmethylsulfonyl fluoride (PMSF), and pilocarpine hydrochloride were from Wako Pure Chemical

Industries., Ltd. (Osaka, Japan). Bio-Rad protein assay kit was obtained from Bio-Rad Laboratories

(Hercules, CA). Complete EDTA-free protease inhibitor cocktail tablets and digoxigenin

(DIG)-luminescent detection kit for nucleic acids were from Roche Diagnostics (Basel, Switzerland).

Cevimeline hydrochloride was a generous gift from Daiichi Pharmaceutical Co., Ltd. (Tokyo, Japan). The

DNA ladder size makers (100 bp) were from New England Biolabs (Beverly, MA). Dimethyl sulfoxide

(DMSO), ethylenediaminetetraacetic acid (EDTA), Tri-reagent, collagenase, and propidium iodide were

purchased from Sigma Chemical Co. (St. Louis, MO). MOCAc-Pro-Leu-Gly,

MOCAc-Gly-Lys-Pro-Ile-Leu-Phe-Phe-Arg-Leu-Lys(Dnp)-D-Arg-NH2,

MOCAc-Gly-Ser-Pro-Ala-Phe-Leu-Ala-Lys(Dnp)-D-Arg-NH2, Suc-Leu-Leu-Val-Tyr-MCA,

Z-Arg-Arg-MCA, and 7-Amino-4-methylcoumarin (AMC) were from the Peptide Institute (Osaka, Japan).

Enhanced chemical luminescence (ECL)-detection kit and Percoll were obtained from GE Healthcare

(Buckinghamshire, UK). Leupeptin, pepstatin A, and 3-morpholinopropanesulfonic acid (MOPS) were

from Dojindo Molecular Technologies, Inc. (Kumamoto, Japan); and SuperScriptTM One-step RT-PCR

with Platinum® Taq system came from Invitrogen Corporation (Carlsbad, CA). One-step SYBR® RT-PCR

Buffer III, TaKaRa Ex Taq® HS, and PrimeSciptTM RT enzyme Mix II were purchased from Takara Bio Inc.

(Tokyo, Japan). Fuji RX X-ray film was procured from Fuji Film Co., Ltd. (Kanagawa, Japan).

Animals and operation

Seven-week-old male Sprague-Dawley (SD) rats, weighing 170 to 190 g, were purchased from SLC Co.,

Ltd. (Shizuoka, Japan). Since 20-30% of the rats obtained from the breeder have a mutation in their AQP5

gene, resulting in lower membrane expression of its gene product (34), we selected the rats expressing the

normal amount of AQP5, as described previously (35), and mated them in our animal facility. They were

kept in our animal facility with free access to water and laboratory chow. Illumination in the animal house

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was controlled such that it was lighted during the period of 06:00-18:00. Cervical sympathetic trunk

denervation (CSTD) or chorda tympani denervation (CTD) was performed under anesthesia with sodium

pentobarbital (50 mg/kg BW i.p.) as described previously (55). The front cervical skin of a group of rats

was incised, and they were regarded as a sham-operated control for CSTD rats. To some denervated rats,

cevimeline hydrochloride (10 mg/kg BW p.o.), pilocarpine (0.3 mg/kg BW p.o.) or chloroquine

diphosphate (50 mg/kg) was administered every day for 7 days starting from the 15th day after the

operation. The non-operated animals were given water or the above drugs for the same period of time. All

reagents were dissolved in sterile distilled water at a concentration so that 0.1ml/100g body weight of the

reagents would be given. The exact amount of the reagents was given after the body weight of each animal

was measured. On the 22nd day after the operation the animals were killed, and their SMGs were

immediately dissected.

The tissues were processed for use in the assays to determine AQP protein and mRNA levels and for

immunohistochemistry. The SMG tissue was also used to prepare acinar cells and lysosomal fraction,

which is described below. For the time-course study and the study to examine the effect of denervation

(Figs. 1, 2, and 5), unilateral CTD (left side) was performed while in the experiment where effects of

agonists/inhibitors (chloroquine) was examined (Figs. 3, 4, 6, 7, and 9), bilateral CTD was performed. This

is because the contralateral gland would become a good control in experiments to examine the effects of

denervation. On the other hand, in experiment where effects of agonists/chloroquine were examined,

bilateral CTD was carried out in order to avoid the possible and unnecessary effects on the experimental

gland from contralateral gland when unilateral CTD was performed.

The protocol applied for the present animal experiment was approved by the Institutional Review Board

of the Animal Committee of the University of Tokushima.

Isolation of acinar cells by Percoll gradient centrifugation

The SMGs from control, CTD, and cevimeline hydrochloride-administered CTD rats were cut into

small pieces and digested at 37°C for 15 min in Mg2+-free HBSS containing 0.03% collagenase, 250 IU/ml

hyaluronidase, 0.2 mM CaCl2 (2 ml/gland) under constant shaking at strokes of 100 cycles/min. The

digested tissue was pipetted 10 times with a 5-ml pipette having a 2-3 mm opening. The tissue sample was

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mixed with 5 ml of Ca2+, Mg2+-free cold HBSS and stood for 10 min to allow sedimentation of the

organoid. The supernatant was removed, and the organoid was washed as described above and stood again

for 10 min for sedimentation. Two ml of diluted organoid (4 ml/gland) was placed on top of a pre-chilled

isotonic 65% Percoll solution in Ca2+, Mg2+-free HBSS (8 ml), which was prepared as described by

Vincent and Nadeau (50), and centrifuged at 500 x g for 30 min in a refrigerated centrifuge (HITACHI

05PR-22; Hitachi Koki Co., Ltd. Tokyo, Japan). The 2 layers were separated; and the lower one containing

acini was recovered, washed, and subjected to enzyme assay and Western blotting of Na+, K+-ATPase

α-subunit. The purity of isolated acini as well as that of ducts was confirmed by phase-contrast microscopy

and by Western blotting for kallikrein.

Preparation of the total membrane fractions and Western blotting

Tissue specimens were dissected as described above and homogenized in 9 volumes (w/v) of ice-cold

homogenization buffer (5 mM HEPES buffer, pH 7.5, containing 50 mM mannitol, 10 mM MgCl2, 1 mM

PMSF, 1 µg/ml aprotinin, 2 µg/ml pepstatin A, 2 µg/ml leupeptin, and 1 tablet of complete EDTA-free

protease inhibitor cocktail per 25 ml of buffer) by using a 15-ml taper-type glass mortar fitted with a

Teflon pestle (Wheaton Science Products, Millville, NJ). The homogenate was filtered through a Nylon

mesh (150 mesh), and the nuclear fraction was removed by centrifugation at 600 x g for 10 min at 4°C.

The supernatant thus obtained was centrifuged at 105,000 x g for 1 h at 4°C to obtain the pelleted total

membrane fraction, which was next suspended in the same homogenization buffer. The protein

concentration in each sample was measured by using a Bio-Rad protein assay kit with bovine serum

albumin as the standard (22).

For analysis of AQP proteins, samples were incubated at 37°C for 30 min in a sample-loading buffer

(0.0625 M Tris-HCl buffer, pH 6.8, containing 2% SDS, 10% glycerol, 0.001% BPB, and 5%

2-mercaptoethanol), and subjected to SDS-PAGE; i.e., protein samples (5 µg each) and standard molecular

mass protein markers were electrophoretically resolved in 12% polyacrylamide slab gels by using a

Mini-Protean II device (Bio-Rad, Richmond, CA). After electrophoresis, the proteins separated on the gel

were transferred onto a nitrocellulose filter by means of a Bio-Rad Mini Trans Blot apparatus according to

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Towbin et al. (46). The blotted filter was blocked with 3% skim milk, probed with anti-AQP5 or

anti-AQP1 antisera (both 3,000-times diluted in blocking solution), and then incubated with 3000-times

diluted peroxidase-labeled anti-rabbit IgG. Anti-AQP5 and anti-AQP1 antibodies preabsorbed with the

respective C-terminal peptide at a final concentration of 40 µg peptide/ml of 3,000-times diluted antiserum

solution were also reacted with the blotted filter in place of non-preabsorbed antiserum to verify the

specificity of the band. All filters were treated with ECL chemical luminescence reagents, and exposed to

X-ray films. The bands appeared in X-ray film were digitalized and quantified by NIHImage software. The

bands in one experiment and separated into more than two gels were normalized by the intensity of the

bands of the same 2-3 samples run in each gel.

Western blotting of Na+, K+-ATPase α-subunits and kallikrein were performed as described previously

(25, 54). Na+, K+-ATPase α-subunit assay was run with 10 µg of sample protein, which was treated with

sampling buffer at 37°C for 30 min, resolved by 7.5% polyacrylamide electrophoresis, and probed with

300,000-times diluted antiserum for Na+, K+-ATPase α-subunits. For the kallikrein assay, 1 µg protein was

incubated with sampling buffer at 95°C for 5 min, resolved by 12% polyacrylamide electrophoresis, and

probed with 100,000-times diluted anti-kallikrein antiserum. Other processes were the same as those

described above.

Isolation of the lysosomal fraction, extraction of lysosomal enzymes, and AQP5 degradation assay in vitro

The SMG lysosomal fraction was prepared according to Tsuji et al. (48), with a slight modification; i.e.,

the SMG was homogenized in 5 volumes of 0.25 M sucrose containing 10 mM Tris-HCl and centrifuged at

1,200 x g at 10 min. The supernatant was recovered and centrifuged at 17,000 x g for 20 min. The pellet

rich in lysosomes was suspended in 10 mM sodium phosphate buffer (pH 6.0), frozen and thawed once,

and homogenized in 10 volumes of 10 mM sodium phosphate buffer (pH 6.0). The pellet thus treated was

centrifuged at 17,000 x g, and the supernatant was recovered. The recovery and purification of lysosomes

by this procedure were 47% and 4-fold, respectively, on the basis of the pepstatin-A-sensitive cathepsin D

activity.

The protein concentration of the lysosomal extract was measured, and the AQP5- degrading activity of

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the extract was then assayed. The degradation of AQP5 by this lysosomal extract was assayed as follows:

the total membrane fraction prepared as described in the previous section was mixed at 2 °C with the

lysosomal extract in a reaction mixture consisting of 16 mM acetate buffer (pH 4.0) containing 33 mM

NaCl (final concentration). The mixture was then incubated at 37 °C for 1 h, after which the reaction was

stopped by adding sampling buffer for Western blotting. Western blotting for AQP5 was then carried out as

described above.

Measurement of cathepsin D activity

Samples for the measurement of cathepsin D activity were prepared as described by Baskin-Bey et al.

(5) with slight modifications. The SMG tissue was removed from animals anesthetized with sodium

pentobarbital (50 mg/kg BW, i.p.) and cut into small pieces. The tissues were placed into a Teflon glass

homogenizer and homogenized at 4°C with 9 volumes (w/v) of buffer consisting of 70 mM sucrose, 220

mM mannitol, 1 mM EGTA, and 10 mM HEPES (pH 7.4). The homogenate was centrifuged at 600 x g for

15 min at 4°C to remove the nuclei and cell debris. The supernatant was repeatedly frozen-and-thawed 3

times (-80°C and RT) to allow lysis of the lysosome. The cell fraction separated by Percoll was also treated

in the same way.

Cathepsin activities were measured as described by Yasuda et al. (56). Fluorogenic peptide substrates

MOCAc-Gly-Lys-Pro-Ile-Leu-Phe-Phe-Arg-Leu-Lys-(Dnp)-D-Arg-NH2,

MOCAc-Gly-Ser-Pro-Ala-Phe-Leu-Ala-Lys(Dnp)-D-Arg-NH2, and Z-Arg-Arg-MCA were used for

measurements of cathepsin B, cathepsin D plus E, and cathepsin E activities, respectively. Cathepsin D

activity was calculated by subtracting the cathepsin E activity from the cathepsin D plus E activity. All of

the substrates were dissolved in DMSO at a 1 mM concentration and stored at -20°C until used. The

reaction mixture consisted of 986 µl of buffer (50 mM AcONa, pH 4.0, containing 100 mM NaCl), 4 µl of

1 mM substrate solution, and 10 µl of the sample solution in a total volume of 1 ml. The changes in

fluorescence intensity produced by substrate cleavage were measured by timed scanning at an emission

wavelength of 393 nm with excitation at 328 nm (for cathepsins D plus E and E), and at an emission of

460 nm with excitation at 380 nm (for cathepsin B), by using a Fluorescence Spectrophotometer F-4500

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(Hitachi, Tokyo, Japan) operating at room temperature. The fluorescence of 0.1 µM MOCAc-Pro-Leu-Gly

solution and 10 µM AMC solution in DMSO were measured as a standard to compute moles of substrate

hydrolyzed. The spontaneous degradation during measurements before addition of enzyme samples was

below the detectable level for all cathepsin assays and proteasome assays described below.

Measurement of proteasome activity

Samples for the measurement of proteasome activity were prepared as described by Bardag-Gorce et al.

(4). The tissue was homogenized in 9 volumes (w/v) of buffer comprising 50 mM Tris-HCl (pH 7.5), 10%

glycerol, 5 mM EDTA, 1 mM EGTA, 1 mM DTT, 1 mM PMSF, and 2.5 µM pepstatin-A at 4°C. The

homogenate was centrifuged at 105,000 x g for 1 h at 4°C, and the supernatant obtained was used for

protein and enzyme assays. The chymotrypsin-like proteosomal activity was measured as described by

Tsukahara et al. (49). The reaction mixture contained 980 µl of 50 mM Tris-HCl (pH 7.5), 10 µl of 10 mM

Suc-Leu-Leu-Val-Tyr-MCA as a substrate, and 10 µl of sample solution in a total volume of 1 ml. The

fluorescence emission of AMC was measured by timed scanning at the wavelength of 460 nm with

excitation at 380 nm at room temperature. AMC solution (100 µM) in the reaction buffer was used as a

standard to compute moles of substrate hydrolyzed.

Preparation of total RNA and reverse transcriptase-polymerase chain reaction (RT-PCR)

Tissues for RNA preparation were quickly processed by using Tri-reagent according to the

manufacturer’s protocol. The RNA concentration was measured by use of a NanoDropND-1000

(NanoDrop Technologies, Inc. Wilmington, DE). By running RT-PCR described below, each RNA sample

thus prepared was verified that they were not contaminated with substantial amount of genomic DNA, and

subjected to real-time PCR also. RT-PCR amplification was performed by using the SuperScriptTM One

Step RT-PCR system in a thermal cycler (TaKaRa Thermal Cycler MP, model TP 3000, Shiga, Japan). To a

final volume of 25 µl, the following components were mixed on ice: 12.5 µl of 2x reaction mixture, 5 pmol

of each primer, 0.5 µl of RT/Taq mix, and 0.5 µg of template RNA. The RT reaction (cDNA synthesis) was

carried out at 45°C for 30 min. The reaction mixture was then incubated at 94°C for 2 min to inactivate the

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enzyme and denature the RNA/cDNA hybrid. The DNA amplification by PCR was next performed for 30

cycles, each cycle consisting of denaturation at 94°C for 15 sec, primer annealing at 55°C for 30 sec, and

extension at 72°C for 1.5 min.

In the present study, the primers used were prepared according to the published sequences (13, 22,): i.e.,

5’-CCCCCAAGGCACCATGAAAAA-3’ (sense) and 5’-TCACGAATCTCTGAGGTCTG-3’ (antisense)

for AQP5, and 5’-ATGGCCAGCGAAATCAAGAAG-3’ (sense) and

5’-TTTGGGCTTCATCTCCACCC-3’ (antisense) for AQP1. The product sizes were 1074 and 813 bp,

respectively. No PCR product other than these bands appeared in the present experiment. A 2-µl volume of

the RT-PCR product was size-fractionated in 3% agarose in TAE buffer, and stained with ethidium

bromide (2 µg/ml) by a standard procedure. The gel was placed in ChemiImager (Alpha Innotech Co., San

Leandro, CA), and photographs were taken by a digital camera.

Real-time PCR

The real-time RT-PCR was performed by using a real-time thermal cycler (Takara Thermal Cycler Dice

Real time System, Shiga, Japan). To a final volume of 25 µl, the following components were mixed on ice:

12.5 µl of 2x One-step SYBR® RT-PCR Buffer III, 2.5 units of TaKaRa Ex Taq® HS, 0.5 µl of

PrimeScriptTM RT enzyme Mix II, 5 pmoles of each primer, 0.1 ng of template RNA, and RNase-free

distilled H2O to make a total volume 25 µl. The RT reaction (cDNA synthesis) was carried out at 42°C for

5 min. The reaction mixture was then incubated at 95°C for 10 sec to inactivate the enzyme and denature

the RNA/cDNA hybrid. The DNA amplification by PCR was next performed for 40 cycles, each cycle

consisting of denaturation at 95°C for 5 sec, primer annealing and extension at 60°C for 30 sec. Primer sets

used for quantification were as follow: 5-TGGCCACCCTCATCTTCGT-3 (sense) and

5-GATGTGGCCACCACTCACAG-3 (antisense) for AQP5, 5-AGGCTTCAATTACCCACTGGA-3

(sense) and 5-TTTGGGCCAGAGTAGCGATG-3 (antisense) for AQP1, and

5-ACCCTAAGGCCAACCGTGAA-3 (sense) and 5-ACAGCCTGGATGGCTACGTACA-3

(antisense) for β-actin. The product sizes were 149, 98, and 85 bp, respectively. The PCR product was

subjected to dissociation conditions, i.e., to confirm that it had a single component. The relative mRNA

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amount for AQP5 and AQP1 was calculated and normalized based on the amount of β-actin.

Northern blotting

Twenty micrograms of total RNA prepared as described above was ethanol-precipitated and dissolved

in 20 µl of freshly prepared sample buffer composed of 1x MOPS solution (20 mM MOPS, 5 mM sodium

acetate, 1 mM EDTA; pH 7.0), 50% formamide, 6.4% formaldehyde, and 5.3% autoclaved glycerol. The

RNA sample was heated at 95°C for 2 min and then rapidly cooled on ice, after which 10 µl (10 µg) of it

was applied to a 1% agarose gel containing 2% formaldehyde. The RNA separated by electrophoresis was

transferred onto a Nylon filter, Hybond-N+, by the standard procedure. The filter was pre-hybridized in

hybridization buffer without probe at 42°C for at least 3 h, and then hybridized at the same temperature for

more than 15 h in hybridization buffer composed of 5x SSPE (1x SSPE = 150 mM NaCl, 10 mM sodium

phosphate, and 1 mM EDTA; pH 7.4), 50% formamide, 2.5x Denhardt’s solution, 10% SDS, 0.2 mg/ml

salmon sperm DNA, 10% dextran sulfate, and 50 ng/ml DIG-labeled cDNA probe specific for AQP5 (see

below for preparation of the probe). The filter was washed with 2x SSPE at room temperature for 5 min, at

42°C for 20 min, and then with 1x SSPE at 42°C twice each for 20 min. It was further washed with

washing buffer for 5 min and then incubated for 1 h at room temperature in 1% blocking solution supplied

in the DIG Luminescence Detection Kit. The filter was then reacted at room temperature for 1 h with

10,000 times-diluted alkaline phosphatase-conjugated anti-DIG-Fab in 1% blocking solution, and then

washed in washing buffer twice, each time for 15 min. It was equilibrated with buffer III, which was

composed of 0.1 M Tris-HCl (pH 9.5), 0.1 M NaCl, and 50 mM MgCl2, and subsequently incubated with

100 times-diluted CSPD in buffer III for 5 min. The excess liquid was blotted onto Whatman 3 MM filter

paper; and the filter was placed in a hybridization bag, incubated at 37°C for 15 min in an air oven, and

finally exposed to Fuji RX-X ray film.

Synthesis of DIG-labeled cDNA probes

The DIG-labeled cDNA probe was prepared by PCR using DIG-labeling Mix and AQP5 cDNA as a

template. An aliquot of 2.5 µl PCR DIG-labeling Mix (10x), 5 pmol of each primer, 2 U of Taq DNA

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polymerase, 50 pg of template AQP5 cDNA, and 1x PCR buffer (supplied with Ex TaqTM polymerase,

Takara) were mixed in a total volume of 25 µl to make the reaction mixture. The sequences of sense and

antisense primers, as well as the PCR thermal condition used in this labeling experiment, were the same as

those described above (see “Preparation of total RNA and reverse transcriptase-polymerase chain reaction

(RT-PCR)”). Complementary DNA of AQP5 used for template DNA was prepared from total RNA by

RT-PCR as described in the same section above, purified by electrophoresis, and recovered by use of a

Qiagen spin column 50 (QIAGEN GmbH, Germany).

Immunohistochemistry

For AQP5 immunohistochemistry, rats were anesthetized, and a fixative (3% paraformaldehyde and 0.1%

glutaraldehyde in 50 mM Na-Pi buffer, pH 7.4) was injected via the left ventricle and circulated throughout

the body for 5 min. The SMG tissues were dissected, cut into 5-mm3 cubes, and immersed in the same

fixative at 4°C for 2 h. The fixed tissue was washed at 4°C overnight in PBS (pH 7.4) containing 6.8%

sucrose. The specimens were then embedded in Tissue-Tec Oct compound and rapidly frozen in liquid

nitrogen. Frozen sections of 5-µm thickness were cut and fixed further in ethanol at -20°C for 1 min. The

sections were washed in PBS and blocked with 1.5% goat serum in PBS, and immunoreacted with

1,000-times diluted rabbit anti-AQP5 antiserum (primary antibody). After having been washed with PBS,

the sections were reacted with 200 times-diluted FITC-conjugated affinity purified goat anti rabbit IgG

(H+L) (second antibody), and washed with PBS. For control staining, sections were incubated with the

same concentration of antibody preabsorbed with the peptide used as the immunogen (40 µg/ml). All

sections were next incubated at room temperature for 15 min with PBS containing 0.1 µg/ml of propidium

iodide and 20 µg/ml of RNase A, and then washed with PBS to allow the nucleus to become stained. The

stained specimens were examined under a fluorescence microscope equipped with a DXM 1200 digital

camera (Nikon, Tokyo, Japan), with excitation at 450-490 nm (for FITC) and 510-560 nm (for propidium

iodide).

Statistics

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For statistical analysis of time-course experiments, one-way ANOVA followed by Tukey compromise

multiple comparison was applied as a post-hoc test. Mann-Whitney’ U-test was used for other comparisons.

Although the U-test is for non-parametric analysis, SEs were provided for all data.

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Results

Effects of parasympathectomy and sympathectomy on the expression of AQP5 and AQP1

In order to explore the possible involvement of the autonomic nervous system in the regulation of

expression of SMG AQP5 and AQP1, we examined the effects of parasympathectomy and sympathectomy

on the level of these salivary gland water channel proteins. First we examined the effects of CTD on the

AQP levels as well as on the relative SMG weight (gland weight/body weight). When the relative SMG

weights were measured from 1 week to 4 weeks after the operation, the weight of the denervated gland had

decreased to 64.7±1.37% of that of the contralateral gland at 1 week after the operation; and this level was

sustained at least up to 4 weeks. The differences were statistically significant (p<0.01, n=5; Fig. 1a),

confirming that the SMG was strongly dependent on parasympathetic nerves. The ratio of AQP5

expression

Fig. 1a, b

in the denervated gland to that in the contralateral gland decreased slowly and linearly, becoming

significantly different (p<0.05 and p<0.01, n=5) at 2, 3, and 4 weeks after the operation (with no

significant change between values of the contralateral gland and the gland from non-treated control). At 4

weeks after CTD, this ratio for the operated animal was 37±9.45% of that for the non-treated animal (Fig.

1b).

We next examined 1) whether AQP1 was also affected by CTD, and 2) if sympathetic innervation was

also indispensable for persistent expression of AQP5 in the SMG. As shown in Fig. 2a, the AQP1 level

was hardly affected by CTD. This result does not contradict the fact that SMG AQP1 is expressed in the

Fig. 2a, b

capillary endothelial cells (3, 28), which are not innervated by autonomic nerves. Secondly, in order to

confirm whether or not the SMG AQP5 level was specifically regulated by parasympathetic nerves, we

bisected the cervical sympathetic trunk (CSTD) and examined the AQP level. As shown in Fig. 2b,

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sympathetic denervation did not decrease the expression level of AQP5 protein, even at 4 weeks after the

operation. By CSTD, however, the gland weight was decreased to approximately 81.0±3.61% (p<0.01,

n=5), indicating that the sympathetic nerves, although not so strongly as the parasympathetic ones, also

support the SMG. The data presented in Fig. 2b as compared with the result obtained by CTD, also

shown in this figure, obviously indicate that the protein level of SMG AQP5 was positively regulated by

parasympathetic, not sympathetic, nerves.

Effects of administration of cevimeline hydrochloride and pilocarpine on AQP protein expression

Since the expression of AQP5 protein was significantly reduced by CTD, the involvement of muscarinic

receptors in up-regulation of AQP5 was considered likely. Therefore, the effects of 2 muscarinic agonists,

cevimeline hydrochloride and pilocarpine, on AQP5 in the SMG of CTD rats were examined next (Fig. 3a

and b). In this experiment and experiment thereafter except that for Fig. 5, bilateral CTD was performed

(see also Materials and Methods section). The result of Western blotting showed that bilateral CTD

resulted in prominent decrease comparing to unilateral CTD, implying the existence of a “cross-talk”

between the two glands. The oral administration of cevimeline hydrochloride (10 mg/kg BW), an M3

receptor agonist,

Fig. 3a, b

for 7 days significantly (p<0.01, n=5) elevated the AQP5 protein level reduced by CTD without affecting

the gland weight. The level of AQP1, which was not changed by CTD, was significantly (p<0.05, n=5)

increased above the control one by the administration of cevimeline hydrochloride (Fig. 3a). For the

protein levels of AQP1 in the SMG of non-operated rats, the effects of cevimeline were minimal. On the

other hand, the oral administration of pilocarpine had no effect on the SMG AQP5 protein level of the

non-treated or CTD rat (Fig. 3b). We used pilocarpine at a dose (0.3 mg/kg BW) previously reported to

stimulate salivary secretion (18), which dose, in fact, provoked salivary secretion significantly (1236±210

mg/30 min vs. control, 112±65 mg/30 min; p=0.0002, by the Mann-Whitney U-test).

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Since apparent changes in the protein level of AQP5 were observed following CTD and cevimeline

hydrochloride administration after CTD, we thought that the localization of AQP5 after these treatments

should be verified. AQP5 is known to be localized predominantly in the apical (32) and basolateral

membranes (35). The result of immunohistochemistry showed that such localization was not changed

appreciably by CTD or by cevimeline administration after CTD (Fig. 4). However, AQP5 positive

stainings of the plasma membrane and cytoplasm in the acinar cells were decreased by CTD and increased

by cevimeline injection after CTD.

Fig. 4a-g

Effects of CTD and administration of cevimeline hydrochloride on the AQP5 and AQP1 mRNA levels in the

SMG

We next examined the possibility that CTD and/or cevimeline hydrochloride might have affected the

transcription of AQP mRNAs. For this purpose, AQP5 mRNA levels after CTD were first examined by

Northern blotting using the DIG-labeled cDNA probe. In this experiment, a 1.6-kb band corresponding to

AQP5 mRNA was specifically detected in all total RNA samples from the SMG of CTD rats as well as

from those of the control gland (Fig. 5). The AQP5 mRNA level in the denervated SMG was almost the

same as that for the contralateral gland from week 0 to week 4 after the operation.

Furthermore, the administration of cevimeline hydrochloride did not affect the mRNA levels of AQP5

nor AQP1 as determined by RT-PCR (Fig. 6a) and by real-time PCR (Fig. 6b). These data suggest that the

Fig. 5a, b

Fig. 6a, b

changes in the AQP5 protein level caused by CTD and/or cevimeline hydrochloride administration were

regulated not transcriptionally but probably by post-transcriptional events.

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Possible involvement of lysosomal system in AQP5 degradation

The results obtained above suggest that the autonomic regulation of the protein level of SMG AQP5

(and AQP1 in part) may be controlled post-transcriptionally. If this is the case, it is possible that the protein

degradation system is involved in the effects on the protein level of SMG AQP5 by CTD/cevimeline

hydrochloride administration. Among the 2 protein-metabolizing systems (proteasome and lysosomal

systems), possible involvement of the lysosomal system was examined. In order to confirm this possibility,

chloroquine, which is known as a denaturant of lysosomes (44), was administered to normal and CTD rats;

and the protein level of AQP5 was then examined. No significant change in AQP5 level was observed

when chloroquine was administered to non-operated rats. However, administration of chloroquine (50

mg/kg BW, p.o.) elevated the AQP5 protein level decreased by CTD (Fig. 7), supporting the idea that an

enzyme(s) in the lysosomal system is involved in the reduction of the protein level of SMG AQP5 caused

by CTD. We observed that the administration of chloroquine did not affect the gland weight in both

non-operated and CTD rats. Thus, the possibility that a lysosomal extract would be able to degrade AQP5

Fig. 7

was next confirmed by conducting an in vitro experiment (Fig. 8). As shown in Fig. 8, the lysosomal

extract degraded AQP5 protein in the total membrane fraction. Although this is a preliminary experiment

and requires more detailed data, this result implies the possible involvement of some lysosomal enzyme(s)

in

Fig. 8

the autonomic control of the AQP5 protein level in the SMG. Since CTD, an experimental protocol

employed in this study, resulted in a significant decrease in the gland weight, we assumed that this

operation may have elevated the autophagy or induced similar cell physiological changes resulting in

acceleration of protein catabolism in the SMG. Thus we examined the activity of several lysosomal

enzymes as well as the proteasome activity of the SMG for the purpose of knowing if the proteolytic

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enzyme in lysosomes or those in the cytosol were actually changed by CTD and/or cevimeline

administration. Accordingly, the activities of proteasome and some selected lysosomal proteinases, i.e.,

cathepsins B, D, and E, were measured. The results of these assays showed that the activities of these 3

cathepsins, especially cathepsin D, were increased by CTD and that this increase was suppressed by the

administration of cevimeline hydrochloride (Fig. 9a-c). On the other hand, the proteasome activity was not

affected significantly by either CTD or administration of cevimeline hydrochloride (Fig. 9b).

Fig. 9a-d

Since the SMG consists of ducts and acini as major constituents, it was indispensable to verify that the

change observed in the cathepsin activity of the entire tissue also took place in the acinar cells. Thus acini

were isolated from the SMG by the Percoll centrifugation method. As shown in Table I, the purity of the

Table I

acini isolated by the present procedure was verified to be 91.6%-97.2% based on the assay data for

kallikrein, a duct cell marker. Using such cell samples, we measured the cathepsin D activity. As shown in

Table I, the cathepsin D activity in these isolated cell samples changed similarly as in the entire tissue,

suggesting a parallel relation between the changes in the entire tissues and those in the isolated acinar cells.

These results suggest that CTD increased the activities of lysosomal enzyme(s) and thereby elevated

protein metabolism activity in the acinar cells as well.

In addition, the amount of Na+,K+-ATPase α-subunit in the isolated acinar cells was also determined, and

the effect of CTD on the level of this protein was examined. The results showed that CTD effects were not

restricted to the AQP5 protein level but that the denervation also affected at least one other membrane

protein of acinar cells, i.e., the Na+,K+-ATPase α-subunit. Different from AQP5, however, the

Na+,K+-ATPase α-subunit was not up-regulated by cevimeline, thus suggesting a relatively specific effect

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of this agonist on AQP5 induction.

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Discussion

In the present study, we examined the role of the autonomic nervous system in the expression of water

channels AQP5 and AQP1 in the rat SMG by assessing the effects of parasympathectomy, sympathectomy,

and the administration of muscarinic agonists cevimeline hydrochloride and pilocarpine. The level of the

AQP5 protein in the membrane fraction of the SMG was decreased by unilateral denervation of the chorda

tympani nerve (CTD). This decrease was accompanied by a reduction in the gland weight, although the

time course of the AQP5 decrease was a little different from that seen for the decrease in the gland weight.

The AQP5 protein level did not decrease after CSTD, and sham operation in this experiment, suggesting

the selective regulation of AQP5 by parasympathetic nerves in the SMG, and that operation-accompanying

inflammation did not affect the AQP5 expression, which is different from lung tissue (44)

Effects of CTD and/or cevimeline treatment on Na+, K+-ATPase α-subunit and dipeptidyl peptidase

IV (DPPIV), a general marker of plasma membrane, were also examined as control experiments. Similar to

their effects on AQP5, the Na+, K+-ATPase α-subunit level was decreased by CTD and increased by

cevimeline treatment in non-operated control and CTD rats. However, we found that no difference was

observed in the DPPIV level when effects of CTD and/or cevimeline treatment were examined (Data not

shown). Thus the changes in the level of Na+, K+-ATPase α-subunit as well as AQP5 appear to be specific

effects caused by CTD and/or cevimeline treatment.

The secretion of submandibular saliva was measured in rats received CTD and cevimeline

administration. The secretion during initial 5 min after pilocarpine administration was hardly affected by

CTD and/or cevimeline treatment. The secretion between 5 and 10 min after pilocarpine injection was

reduced by CTD and increased by cevimeline. Particularly, 56% increase (p<0.05, n=4) in saliva secretion

was observed by cevimeline when it was given to CTD rats (Data not shown). The result indicates that the

AQP5 level, when altered by the operation/therapy, affects the saliva secretion.

The neural mechanism regulating AQP channels has been reported for the kidney (19) and muscle

(26), although denervation is reported to up-regulate the AQP5 protein level in sweat gland (29). Thus the

neural control of AQP does not appear to be special for the salivary gland.

The rate of decrease in the AQP5 protein level following CTD was very slow. The level decreased

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linearly, and its T1/2 was approximately 25 days. Since the AQP5 mRNA level was not changed by CTD,

it is probable that the activity of the protein degradation system was elevated by CTD, provided that the

rate of protein synthesis was not increased. Such changes would have led to a new equilibrium of synthesis

and degradation of AQP5. This possibility was implied also by immunohistochemical observation, in

which the number of AQP5-bearing vesicles was changed by CTD/cevimeline injection.

Generally, the affinity of an agonist to the receptor determines the physiological response and

selectivity of the agonist among the receptor subtypes. Thus, both cevimeline and pilocarpine provoke the

saliva secretion although its intensity is different (18). The effects of cevimeline on the non-denervated rats

were minimal while the administration of cevimeline hydrochloride after CTD increased the AQP5 levels

reduced by the operation. This difference may be explained by a possibility that CTD may have attenuated

the sensitivity of the muscarinic receptor, since chronic excess agonist can lead to a down-regulation of

receptors, while a deficit of transmitter can lead to increased numbers of receptors and supper sensitivity of

the system (6). In our present study, the second muscarinic agonist, pilocarpine, did not have such an effect

as cevimeline. We speculate that the potency of CTD-induced supper sensitivity may be different for each

receptor subtype. All these elucidations are based on the assumption that effects of cevimeline on the

AQP5 level were due to stimulation of M3 muscarinic receptors (18). This suggestion still remains

tentative and requires more experiments using other M3 muscarinic antagonists or the receptor knock-out

mice, which experiments are now being considered.

Leith et al. (27) showed that the AQP1 protein level increases after application of proteasome

inhibitors in BALB/c fibroblasts spontaneously expressing AQP1, suggesting the involvement of

proteasomes in regulation of the AQP1 protein level. In the present study, on the contrary, the AQP1

protein level in the denervated gland did not change at all as compared with that in the contralateral gland;

and it increased above the control by administration of cevimeline hydrochloride. In the SMG, AQP1 is

known to be localized in capillaries (3, 28) which are not innervated. Despite the lack of apparent

cholinergic innervation, the presence of muscarinic receptors, primarily of the M3 subtype, in the

endothelial cells of most of blood vessel is described (8), which receptor plays a pivotal role for relaxation.

This is probably the reason that denervation did not affect the SMG AQP1 protein level. However, it is not

clear by what mechanism cevimeline hydrochloride induced AQP1 in the SMG. We are now planning an

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experiment using M3/M1 muscarinic receptor knock-out mice to determine if these receptors are involved

in this induction.

Assessment by Northern blotting, RT-PCR, and real-time RT-PCR experiments showed that the AQP5

mRNA level in the SMG was not altered by CTD and/or administration of cevimeline hydrochloride. It is

reported that AQP5 mRNA transcription is suppressed by TNF-α and LPS (47, 52) and induced by

hypoxia and hyper osmolality (57). In a short-term experiment using a murine lung epithelial cell line

(MLE-12), chlorophenylthio-cAMP was shown to induce AQP5 mRNA expression at 6h (53). There is no

report, however, that muscarinic receptors and/or their signaling molecules are responsible for the

regulation of AQP5 mRNA transcription. Our data also show that this is not the case by long term

treatment.

Although the level of AQP5 mRNA did not change after autonomic denervation and/or muscarinic

therapy, the AQP5 protein level was decreased by CTD and increased by cevimeline hydrochloride after

CTD. These data obviously indicate the possibility that either 1) AQP5 protein synthesis is suppressed, 2)

its protein is released into saliva, or 3) AQP5 protein degradation is accelerated by CTD. In the present

study we verified the latter possibility.

As to protein catabolism, the existence of 2 pathways is generally known, i.e., lysosomal proteinases

and the ubiquitin-proteasome system. In the latter system, various protein molecules are metabolized, such

as those involved in cell-cycle regulation, signaling molecules, transcription factors, and oncogene

products. The quality of proteins in the endoplasmic reticulum and cytosol is also known to be controlled

by the ubiquitin-proteasome system. This system removes abnormal proteins such as mutant molecules

or proteins that have failed in normal folding/assembly. The lysosomal enzymes, on the other hand,

degrade extrinsic and intrinsic proteins; and these events are termed heterophagy and autophagy,

respectively (31). The ligands and receptors bound to ligands are decomposed by heterophagy; whereas

various cellular proteins are degraded by autophagy. Autophagy is known to be greatly accelerated under

physiological conditions where animals cannot utilize the food protein, e.g., during fasting/starvation.

In the present study, the SMG weight was decreased by CTD, implying an increase in autophagy or

similar physiological changes took place in acinar cells due to this operation. This idea is supported by

the fact that the activity of some lysosomal proteinases in homogenates (containing lysate of lysosomes)

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was increased by CTD and returned to their normal level by treatment with cevimeline.

Using acinar cells isolated from the SMG, we examined the effects of CTD and administration of

cevimeline on the Na+,K

+-ATPase α-subunit level. The results showed that the level of Na+,K

+-ATPase

α-subunits was decreased by CTD but was not increased by concomitant cevimeline administration. Since

AQP5 and AQP1 protein levels were increased by cevimeline, it is evident that the cevimeline effects were

specific for AQP5 and AQP1. On the other hand, the decrease in AQP5 protein caused by CTD appeared

not to be specific since a similar decrease also was found for the level of Na+,K

+-ATPase α-subunits. This

result again implies an increase in autophagy or similar physiological changes caused by CTD. To

establish this hypothesis, however, the CTD-induced changes of other house keeping proteins originated

form plasma membrane, cytosol, or subcellular structure need to be determined.

Sidhaye et al. (43) reported that the AQP5 protein level reduced by cAMP is increased to its original

level by chloroquine, a denaturant of lysosome, suggesting the possibility that AQP5 is degraded by a

lysosomal enzyme(s). There are also other reports suggesting that AQP2 and AQP8 are degraded by

lysosomal enzymes; e.g., AQP2 expressed in MDCK cells is degraded by lysosomal enzymes after being

ubiquitinated (21), and estrogen-caused reduction in hepatic AQP8 level is provoked by lysosomal

enzymes (9). These reports support our present data suggesting that some lysosomal enzyme(s) is involved

in the regulation of the AQP5 protein level in the SMG. In fact, our preliminary experiment showed that

the extract obtained from the lysosomal fraction of the SMG degraded the AQP5 protein in the total

membrane fraction in vitro. The data obtained in the present study suggest that the salivary gland AQP5

level is regulated by a lysosomal enzyme(s); and therefore the experiment to specify the proteolytic

enzyme(s) responsible for this AQP5 catabolism is now being conducted.

In conclusion, the regulation of the AQP5 protein level in the SMG by CTD and cevimeline is

suggested to be controlled by a lysosomal enzyme(s). In the long-term therapy, transcriptional regulation

of AQP5 mRNA is not likely involved in this neural regulation.

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Acknowledgements

This work was supported in part by grants-in-aid for scientific research (Nos. 13671940 and

13671941) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan, by a grant

from Japan China Medical Association, and by the support of Daiichi Pharmaceutical Co. Ltd.

We are grateful to Daiichi Sankyo Co. Ltd. (Tokyo Japan) for providing us with cevimeline

hydrochloride, and also thank Drs. R. Matsuo of Okayama University and N. Kanamori of our University

for technical suggestions regarding autonomic denervation and statistic analysis, respectively.

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References

1. Akamatsu, T., Parvin, M.N., Murdiastuti, K., Kosugi-Tanaka, C., Yao, C., Miki, O., Kanamori, N.,

and Hosoi, K. Expression and localization of aquaporins, members of the water channel family, during

development of the rat submandibular gland. Pflugers Arch. – Eur. J. Physiol. 446: 641–651, 2003.

2. Agre, P. Molecular physiology of water transport: aquaporin nomenclature workshop. Mammalian

aquaporin. Biol. Cell 89: 225-257, 1997.

3. Agre, P., Bonhiver, M., and Borgnia, M.J. The aquaporin, blueprints for cellular plumbing system. J.

Biol. Chem. 273: 14659-14662, 1998.

4. Bardag-Gorce, F., Li, J., French, B.A., and French, S.W. The effect of ethanol-induced CYP2E1 on

proteasome activity: the role of 4-hydroxynonenal. Experiment. Mol. Pathol. 78: 109-111, 2005.

5. Baskin-Bey, E.S., Canbay, A., Bronk, S.F., Werneburg, N., Guicciardi, M.E., Nyberg, S.L., and

Gores, G.J. Cathepsin B inactivation attenuates hepatocyte apoptosis and liver damage in steatotic

livers after cold ischemia-warm reperfusion injury. Am. J. Physiol. Gastrointest. Liver Physiol. 288:

G396-G402, 2005.

6. Bloom, F.E., Chapter 12, Neurotransmission and the central nervous system, In: Goodman & Gilman’s

The pharmacological basis of therapeutics, 10th edition, edited by. Hardman, J.G.. and Limbird, L.E.

(consulting editor, Gilman, A.G.) New York, MacGraw-Hill, 2001

7. Bradley, R.M. (1995) Salivary secretion. In: Essentials of oral physiology (Bradley, R.M., eds),

p.161-186, Mosby-Year Book, Inc, St. Louis Mo.

8. Brown, J.H. and Taylor P., Chapter 7, Muscarinic agonist and antagonist. In: Goodman & Gilman’s

The pharmacological basis of therapeutics, 10th edition, edited by Hardman, J.G. and Limbird, L.E.

(consulting editor, Gilman, A.G.), New York, MacGraw-Hill, 2001.

9. Carreras, F.I., Lehmann, G.L., Ferri, D., Tioni, M.F., Calamita, G., and Marinelli, R.A. Defective

hepatocyte aquaporin-8 expression and reduced canalicular membrane water permeability in

estrogen-induced cholestasis. Am. J. Physiol. Gastrointes.t Liver Physiol. 292: G905-G912, 2007.

10. Deen, P.M., Verdijk, M.A., Knoers, N.V., Wieringa, B., Monnens, L.A., van Os, C.H., and van

of 46

28

Oost, B.A. Requirement of human renal water channel aquaporin-2 for vasopressin-dependent

concentration of urine. Science 264: 92-95, 1994.

11. Fushimi, K., Uchida, S., Hara, Y., Hirata, Y., Marumo, F., and Sasaki, S. Cloning and expression of

apical membrane water channel of rat kidney collecting tubule. Nature 361: 549-552, 1993.

12. Ishibashi, K., Kuwahara, M., Gu, Y., Kageyama, Y., Tohsaka, A., Suzuki, F., Marumo, F., and

Sasaki, S. Cloning and functional expression of a new water channel abundantly expressed in the

testis permeable to water, glycerol, and urea. J. Biol. Chem. 272: 20782-20786, 1997.

13. Ishibashi, K., Kuwahara, M., Gu, Y., Tanaka, Y., Marumo, F., and Sasaki, S. Cloning and

functional expression of a new aquaporin (AQP9) abundantly expressed in the peripheral leukocytes

permeable to water and urea, but not glycerol. Biochem. Biophys. Res. Commun. 244: 268-274, 1998.

14. Ishibashi, K., Kuwahara, M., Kageyama, Y., Tohsaka, A., Marumo, F., and Sasaki, S. Cloning and

functional expression of a second new aquaporin abundantly expressed in testis. Biochem. Biophys.

Res. Commun. 237: 714-718, 1997.

15. Ishibashi, K., Morinaga, T., Kuwahara, M., Sasaki, S., and Imai, M. Cloning and identification of a

new member of water channel (AQP10) as an aquaporin. Biochim. Biophys. Acta 1576: 335-340,

2002.

16. Ishibashi, K., Sasaki, S., Fushimi, K., Uchida, S., Kuwahara, M., Saito, H., Furukawa, T.,

Nakajima, K., Yamaguchi, Y., Gojobori, T., and Marumo, F. Molecular cloning and expression of

a member of the aquaporin family with permeability to glycerol and urea in addition to water

expressed at the basolateral membrane of kidney collecting duct cells. Proc. Natl. Acad. Sci. U.S.A.

91: 6269-6273, 1994.

17. Itoh, T., Rai, T., Kuwahara, M., Ko, S.B., Uchida, S., Sasaki, S., and Ishibashi, K., Identification of

a novel aquaporin, AQP12, expressed in pancreatic acinar cells Biochem. Biophys. Res. Commun.,

330: 832-838, 2005.

18. Iwabuchi, Y. and Masuhara, T. Sialogogic activities of SNI-2011 compared with those of pilocarpine

and McN-A-343 in rat salivary gland: Identification of a potential therapeutic agent for treatment of

Sjögren’s syndrome. Gen. Pharmacol. 25: 123-129, 1994.

19. Jimi, T., Wakayama, Y., Matsuzaki, Y., Hara, H., Inoue, M., Shibuya, S. Reduced expression of

of 46

29

AQP4 in human muscles with amyotrophic lateral sclerosis and other neurogenic atrophies. Pathol.

Res. Pract., 200: 203-209, 2004.

20. Jung, J.S., Bhat, R.V., Preston, G.M., Guggino, W.B., Baraban, J.M., and Agre, P. Molecular

characterization of an aquaporin cDNA from brain: candidate osmoreceptor and regulator of water

balance. Proc. Natl. Acad. Sci. U.S.A. 91: 13052-13056, 1994.

21. Kamsteeg, E.J., Hendriks, G., Boone, M., Konings, I.B.M., Oorschot, V., Sluijs, P, Klumperman,

J., and Deen, P.M.T. Short-chain ubiquitination mediates the regulated endocytosis of the

aquaporin-2 water channel. Proc Natl Acad Sci USA 103: 18344-18349, 2006.

22. King, L.S., Nielsen, S., and Agre, P. Aquaporins in complex tissues. I. Developmental patterns in

respiratory and glandular tissues of rat. Am. J. Physiol. Cell Physiol. 273: C1541-C1548, 1997.

23. Koyama, Y., Yamamoto, T., Kondo, D., Funaki, H., Yaoita, E., Kawasaki, K., Sato, N.,

Hatakeyama, K., and Kihara, I. Molecular cloning of a new aquaporin from rat pancreas and liver. J.

Biol. Chem. 272: 30329-30333, 1997.

24. Kumagami, H., Loewenheim, H., Beiz, E., Wild, K., Schwarz, H., Yamashita, K., Schultz, J.,

Paysan, J., Zenner, H.-P., and Ruppersberg, J.P. The effect of anti-diuretic hormone on the

endolymphatic sac of the inner ear. Pflugers Arch-Eur. J. Physiol. 436: 970-975, 1998.

25. Kurihara, K., Hosoi, K., Kodama, A., and Ueha, T. A new electrophoretic variant of a subunit of

Na+,K+-ATPase from the submandibular gland of rats, Biochim. Biophys. Acta 1039: 234-240, 1990.

26. Lee, J., Yoo, K., Kim, S.W. Jung K.H., Ma, S.K., Lee, Y.K., Kim, W.Y., Kim, J., Choi K.C.

Decreased expression of aquaporin water channels in denervated rat kidney. Nephron Physiol., 103;

170-178, 2006.

27. Leith, V., Agre, P., and King, L.S. Altered ubiquitination and stability of aquaporin-1 in hypertonic

stress. Proc. Natl. Acad. Sci. USA 98: 2894-2898, 2001.

28. Li, J., Nielsen, S., Dai, Y., Lazowski, K. W., Christensen, E. I., Tabak, L.A., and Baum, B.J.

Examination of rat salivary glands for the presence of the aquaporin CHIP Pflugers Arch. – Eur. J.

Physiol. 428: 455–460, 1994.

of 46

30

29. Ma, L., Huang, Y.-G., He, H., Deng, Y.-C., Zhang, H.-F., Che, H.-L., Tian, J.-Y., Zhao, G. Postnatal

expression and denervation induced up-regulation of aquaporin-5 protein in rat sweat gland. Cell

Tissue Res, 329: 25-33, 2007.

30. Ma, T., Song, Y., Gillespie, A., Carlson, E.J., Epstein, C.J., and Verkman, A.S. Defective secretion

of saliva in transgenic mice lacking aquaporin-5 water channels. J. Biol. Chem. 274: 20071-20074,

1999.

31. Martinez-Vicente, M. and Cuervo, A.M. Autophagy and neurodegeneration: when the cleaning crew

goes on strike. Lancet Neurol. 6: 352-361, 2007.

32. Matsuzaki, T., Suzuki, T., Koyama, H., Tanaka, S., and Takata, K. Aquaporin-5 (AQP5), a water

channel protein, in the rat salivary and lachrymal glands: immunolocalization and effect of secretory

stimulation. Cell Tissue Res. 295: 513-521, 1999.

33. Morello, J. and Bichet, D.G. Nephrogenic diabetes insipidus. Annu. Rev. Physiol. 63: 607-630, 2001.

34. Murdiastuti, K., Miki, O., Yao, C., Purwanti, N., Kosugi-Tanaka, C., Akamatsu, T., Kanamori, N.,

and Hosoi, K. Divergent expression and localization of aquaporin 5, an exocrine-type water channel,

in the submandibular gland of Sprague-Dawley rats. Pflugers Arch-Eur. J. Physiol. 445: 405-412,

2002.

35. Murdiastuti, K., Purwanti, N., Karabasil, M.R., Li, X., Yao, C., Akamatsu, T., Kanamori, N., and

Hosoi, K. A naturally occurring point mutation in the rat aquaporin 5 gene, influencing its protein

production by and secretion of water from salivary glands. Am. J. Physiol. Gastrointest. Liver Physiol.

291: G1081-G1088, 2006.

36. Nielsen, S., Di Giovanni, S.R., Christensen, E.I., Knepper, M.A., and Harris, H.W. Cellular and

subcellular immunolocalization of vasopressin-regulated water channel in rat kidney. Proc. Natl. Acad.

Sci. U.S.A. 90: 11663-11667, 1993.

37. Parvin, M.N., Kurabuchi, S., Murdiastuti, K., Yao, C., Kosugi-Tanaka, C., Akamatsu, T.,

Kanamori, N., and Hosoi, K. Subcellular redistribution of AQP5 by vasoactive intestinal

polypeptide in the Brunner’s gland of the rat duodenum. Am. J. Physiol. Gastrointest. Liver

Physiol.288: G1283-G1291, 2005.

of 46

31

38. Parvin, M.N., Tsumura, K., Akamatsu, T., Kanamori, N., and Hosoi, K. Expression and

localization of AQP5 in the stomach and duodenum of the rat. Biochim. Biophys. Acta 1542: 116-124,

2002.

39. Preston, G.M. and Agre, P. Isolation of the cDNA for erythrocyte integral membrane protein of 28kD:

member of an ancient channel family. Proc. Natl. Acad. Sci. U.S.A. 88: 11110-11114, 1991.

40. Preston, G.M., Carroll, T.P., Guggino, W.B., and Agre, P.. Appearance of water channels in Xenopus

oocytes expressing red cell CHIP28 protein. Science 256: 385-387, 1992.

41. Raina, S., Preston, G.M., Guggino, W.B., and Agre, P. Molecular cloning and characterization of an

aquaporin cDNA from salivary, lacrimal and respiratory tissues. J. Biol. Chem. 270: 1908-1912, 1995.

42. Sasaki, S., Ishibashi, K., and Marumo, F. Aquaporin-2 and -3: representatives of two subgroups of

the aquaporin family colocalized in the kidney collecting duct. Annu. Rev. Physiol. 60: 199-220, 1998.

43. Sidhaye, V., Hoffert, J.D., and King, L.S. cAMP has distinct acute and chronic effects on aquaporin-5

in lung epithelial cells. J. Biol. Chem. 280: 3590-3596, 2005.

44. Sundelin, S.P. and Terman, A. Different effects of chloroquine and hydroxychloroquine on lysosomal

function in cultured retinal pigment epithelial cells. Acta Pathol. Microbiol. Immunol. Scand. 110:

481-489, 2002.

45. Tada, J., Sawa, T., Yamanaka, N., Shono, M., Akamatsu, T., Tsumura, K., Parvin, M.N.,

Kanamori, N., and Hosoi, K. Involvement of vesicle-cytoskeleton interaction in AQP5 trafficking in

AQP5-gene-transfected HSG cells. Biochem. Biophys. Res. Commun. 266: 443-447, 1999.

46. Towbin, H., Stahelin, T., and Gordon, J. Electrophoretic transfer of proteins from polyacrylamide

gels to nitrocellulose sheets: procedure and some applications. Proc. Natl. Acad. Sci. U.S.A. 76:

4350-4354, 1979.

47. Towner, J.E., Krane, C.M., Bachurski, C.J., and Menon, A.G. Tumor necrosis factor-α inhibits

aquaporin 5 expression in mouse lung epithelial cells. J. Biol. Chem. 276: 18657-18664, 2001.

48. Tsuji, A., Akamatsu, T., Nagamune, H., and Matsuda, Y. Identification of targeting proteinase for rat

α1-macroglobulin in vivo. Mast-cell tryptase is a major component of the

α1-macroglobulin-proteinase complex endocytosed into rat liver lysosomes. Biochem. J. 298: 79-85,

of 46

32

1994.

49. Tsukahara, T., Ishiura, S., and Sugita, H. An ATP-dependent protease and ingensin, the

multicatalytic proteinase, in K562 cells. Eur. J. Biochem. 177: 261-266, 1998.

50. Vincent, R. and Nadeau, D. Adjustment of the osmolality of Percoll for the isopycnic separation of

cells and organelles. Anal. Biochem. 141: 322-328, 1984.

51. Walz, T., Hirai, T., Murata, K., Heymann, J.B., Mitsuoka, K., Fujiyoshi, Y., Smith, B.L., Agre, P.,

and Engel, A. The three-dimensional structure of aquaporin-1. Nature 387: 624-627, 1997.

52. Xie, Y.P., Chen, C.P., Wang, J.C., Qian, G.S., Wang, Y.D., and Xiao, Z.L. Experimental study on the

expression and function of aquaporin-1 and aquaporin-5 in rats with acute lung injury induced by

lipopolysaccharide. Zhonghua Jie He He Hu Xi Za Zhi 28: 385-389, 2005.

53. Yang, F., Kawedia, J.D., and Menon, A.G. Cyclic AMP regulates aquaporin 5 expression at both

transcriptional and post-transcriptional levels through a protein kinase A pathway, J. Biol. Chem., 278:

32173-32180, 2003.

54. Yao, C., Karabasil, M.R., Purwanti, N., Li, X., Akamatsu, T., Kanamori, N., and Hosoi, K. Tissue

kallikrein mK13 is a candidate processing enzyme for the precursor of interleukin-1β in the

submandibular gland of mice, J. Biol. Chem. 281: 7968-7976, 2006.

55. Yao, C., Li. X., Murdiastuti, K., Kosugi-Tanaka, C., Akamatsu, T., Kanamori, N., and Hosoi K.

Lipopolysaccharide-induced elevation and secretion of interleukin-1β in the submandibular gland of

male mice. Immunology 116: 213-222, 2005.

56. Yasuda, Y., Kageyama, T., Akamine, A., Shibata, M., Kominami, E., Uchiyama, Y., and

Yamamoto, K. Characterization of new fluorogenic substrates for the rapid and sensitive assay of

cathepsin E and cathepsin D. J. Biochem. 125: 1137-1143, 1999.

57. Zhou, B., Ann, D.K., Li, X., Kim, K.J., Lin, H., Minoo, P., Crandall, E.D. and Borok, Z.

Hypertonic induction of aquaporin-5: novel role of hypoxia inducible factor-1α. Am. J. Physiol.Cell

Physiol. 292: C1280-C1290, 2006.

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Figure legends

Fig. 1 Effects of CTD on the gland weight and AQP5 protein levels of the rat SMG. NT, no

treatment; CTD, chorda tympani denervation; P-Ab, probed with peptide-preabsorbed AQP5

antiserum (sample, from an NT animal). L, left gland; R, right gland; D, denervated gland; C,

contralateral gland. The unilateral chorda tympani nerve was denervated (CTD) as described

in the text, while the contralateral nerve was kept intact. a. Relative gland weight. The relative

gland weights of contralateral (CL) and experimental (CTD) are shown. b. Relative values of

AQPs in the denervated/contralateral gland. The Western blot image was quantified by NIH

Image Software, and means ± SEM for 5 rats are shown. No statistical difference was

observed between the 2 values at 0 week. One-way ANOVA test was then applied to the

non-treated and operated groups. No statistical difference was observed for the non-treated

group but a significant difference was found for the operated group. Tukey’s multiple analysis

was then applied as a post-hoc test. *p<0.05, **p<0.01, significantly different from the

non-treated group before the operation (0 week).

Fig. 2 Effects of sympathectomy and parasympathectomy on the protein levels of AQP5 and

AQP1 in the rat SMG. NT, no treatment; CTD, chorda tympani denervation; CSTD, cervical

sympathetic trunk denervation. P-Ab, probed with peptide-preabsorbed AQP1 antiserum

(sample, from an NT animal). L, left gland; R, right gland; D, denervated gland; C,

contralateral gland, g-AQP1, glycosylated AQP1. a. Effects of CTD on SMG AQP5 and

AQP1. b. Effects of CTD and CSTD on SMG AQP5. Either the left cervical sympathetic

trunk or the left chorda tympani was denervated (CSTD and CTD, respectively). Unilateral

CTD and CSTD were performed as described in the text, and the contralateral nerves were

kept intact in both cases. Rats were killed on 22th day after the operation. The Western blot

images were quantified by using NIH Image Software, and the relative values of AQP5 and

AQP1 in the denervated gland vs. contralateral gland were calculated. The means ± SEM for

5 rats were analyzed by Mann-Whitney U-test. **p<0.01, significantly different from the

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non-treated group. NS, no significant difference from the non-treated group

Fig. 3 Effects of CTD and administration of muscarinic agonists on the AQP5 and AQP1

levels in the SMG of rats. NT, no treatment; CTD, chorda tympani denervation; CM,

cevimeline hydrochloride; PC, pilocarpine; g-AQP1, glycosylated AQP1. Bilateral chorda

tympani nerves were denervated (CTD), and cevimeline hydrochloride or pilocarpine was

administered for 1 week starting from the 15th day after operation. a. Effects of cevimeline

hydrochloride on AQP5 and AQP1 levels in the SMG of CTD rats. b. Effects of pilocarpine

on these levels. The Western blot images were quantified by using NIH Image Software, and

the relative values of AQP5 and AQP1 were calculated. The means ± SEM for 5 rats were

analyzed by Mann-Whitney U-test. **p<0.01, #p<0.05, significantly different from the

non-treated group. §p<0.05, §§p<0.01, significantly different from the CTD group. NS, no

significant difference from the CTD group.

Fig. 4 Immunohistochemical localization of AQP5 in the SMG of CTD rats and CTD rats

administered cevimeline hydrochloride. Bilateral CTD was performed and cevimeline was

administered as described in the text. a. and d., non-treated rats, b. CTD rat, c. CTD rat

administered cevimeline hydrochloride (10 mg/kg body weight). a. – c., anti-AQP5 antiserum.

d. same concentration of anti-AQP5 antiserum preabsorbed with antigen peptide (40 µg/ml). e,

f, and g, are enlarged picture of boxed areas of a, b, and c, respectively. D indicates ducts.

Acinar cells are intensely labeled with anti-AQP5 antibody and shows strong green

fluorescence of FITC. The AQP5 protein as visualized by FITC signals is predominantly

localized at both apical and basolateral membranes in a-c and e-g. Acinar cell size is smaller

in CTD group comparing to non-treated and cevimeline group. Positive stainings for AQP5

are prominent in plasma membrane and cytoplasm of a non-treated rat and a rat administered

with cevimeline, comparing to that of CTD group. Bar indicates 40 µm

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Fig. 5 Effects of CTD on the AQP5 mRNA levels in the rat SMG. L, left gland; R, right

gland; D, denervated gland; C, contralateral gland. Unilateral chorda tympani nerve was

denervated, while the contralateral nerve was left intact. The total RNA from the denervated

SMG (D) and contralateral gland (C) was prepared, and 10 µg of each sample was subjected

to Northern blotting. a. Analysis by Northern blotting. Results of 2 rats in each group are

shown. Upper panel, Northern blotting; lower panel, ethidium bromide staining showing

rRNA. b. Graphical presentation. The Northern blot image was quantified by using NIH

Image Software, and the relative values of AQP5 mRNA were calculated. The means ±

SEM for 5 rats were analyzed by Mann-Whitney U-test. No statistical difference was

observed between control (0 week) and denervated groups.

Fig. 6 Lack of effect of administration of cevimeline hydrochloride on the mRNA levels of

AQP5 and AQP1 in the rat SMG. NT, no treatment; CTD, chorda tympani denervation; CM,

cevimeline hydrochloride. Bilateral chorda tympani nerves were denervated (CTD), and

cevimeline hydrochloride was administered for 1 week starting from the 15th day after the

operation. a. Analysis by RT-PCR. Results of 2 rats in each group are shown. b. Analysis by

real-time PCR. The means ± SEM for 5 rats were analyzed by using the Mann-Whitney U-test.

No statistical difference was observed between control (0 week) and each denervated group.

Fig. 7 Effects of CTD and administration of chloroquine on the AQP5 protein level in the rat

SMG. NT, no treatment; CTD, chorda tympani denervation; CQ, chloroquine administration.

Bilateral chorda tympani nerves were denervated (CTD), and chloroquine was administered

for 1 week starting from the 15th day after the operation. The Western blot images were

quantified by using NIH Image Software, and the relative values of AQP5 were calculated.

The means ± SEM for 5 rats were analyzed by Mann-Whitney U-test. **P<0.01, significantly

different from the non-treated group. §P<0.05, significantly different from the CTD group.

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Fig. 8 Degrading effect of lysosomal extract on protein level of AQP5. Details about the

experimental conditions are described in the text. There was no degradation observed when

the total membrane fraction was incubated under the same condition without added lysosomal

extract.

Fig. 9 Effects of CTD and administration of cevimeline hydrochloride on cathepsin and

proteasome activities of the rat SMG. NT, no treatment; CTD, chorda tympani denervation;

CM, cevimeline hydrochloride. Bilateral chorda tympani nerves were denervated (CTD), and

cevimeline hydrochloride was administered for 1 week starting from 2 weeks after the

operation. a. Cathepsin B activity; b. Cathepsin D activity; c. Cathepsin E activity; d.

Proteasome activity. The means ± SEM for 5 rats were analyzed by Mann-Whitney U-test.

*P<0.05, **P<0.01, significantly different from the non-treated group. §P<0.05, §§P<0.01,

significantly different from the CTD group. NS: no significant difference from the control

group.

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Table 1 Effects of CTD and administration of cevimeline hydrochloride after CTD on the

cathepsin D activity and Na+,K+-ATPase α-subunit levels of isolated SMG acinar cells

Animal treatmentCathepsin D activitya

( mU/mg protein ) (%)Na+,K+-ATPase α-subunit (%)

Relative purityof acinar cell samples(%)c

Non-treated

CTD

CTD+cevimeline hydrochloride

3.44±0.36(100±10.47) b

4.51±0.17(131.10±4.94)*

2.69±0.30(78.20±8.72)**

100±4.48

65.7±1.34§§

57.6±3.28¶.

97.06±1.95

97.20±0.42

91.56±2.5

aThe value after subtraction of the cathepsin E activity from the cathepsin D plus E activities.

bThe values in parentheses indicate the percentage of that of the “non-treated” group. cThe

kallikrein protein in the dissociated cell sample before Percoll separation and in acinar cell

samples after separation was determined by Western blotting. The amount of kallikrein protein

present in the samples before Percoll separation was regarded as 100%. The percentage of

duct cells present in the isolated acinar cell samples was calculated; and the purity of the acinar

cells, determined. ∗p<0.05, §§p<0.01, significantly different from the respective non-treated

group. ∗∗p<0.01, significantly different from the CTD group. ¶, no significant difference from

the CTD group.

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Effects of CTD on the gland weight and AQP5 protein levels of the rat SMG.

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Effects of sympathectomy and parasympathectomy on the protein levels of AQP5 and AQP1 in the rat SMG

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Effects of CTD and administration of muscarinic agonists on the AQP5 and AQP1 levels in the SMG of rats

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Immunohistochemical localization of AQP5 in the SMG of CTD rats and CTD rats administered cevimeline hydrochloride

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Effects of CTD on the AQP5 mRNA levels in the rat SMG

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Lack of effect of administration of cevimeline hydrochloride on the mRNA levels of AQP5 and AQP1 in the rat SMG

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Effects of CTD and administration of chloroquine on the AQP5 protein level in the rat SMG

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Degrading effect of lysosomal extract on protein level of AQP5

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Degradation of submandibular gland AQP5 by parasympathetic denervation of chorda tympani and its...

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