IntroductionAn organ is a mass of diverse cells possessing variousmorphological and functional identities. The diversity of cellsensures that organs develop a highly elaborate multicellularsystem to adapt complex physiological demands. An importantpremise involved in this system is the harmonisation of eachcell activity so as to achieve organ function adequately andefficiently. In most organs, the mechanisms underlying thisharmonisation remain largely unknown.

Cellular activities in virtually all cell types are, despitetremendous diversities in their expression, regulated bycommon intracellular signalling systems, and calcium is oneimportant signalling molecule involved in the regulation ofdiverse cell functions (Berridge et al., 2000). In response toadequate stimuli, [Ca2+]i increases, oscillates and decreases,leading to the activation, modulation and termination of cellfunction. It is likely that the integration of multicellular activityin the organ is achieved through the regulation of Ca2+

signalling. Although extensive studies have elucidatedsubcellular mechanisms responsible for Ca2+ signalling, littleis known of its regulation at the supracellular level.

The salivary gland is an exocrine organ capable of secretingfluid and macromolecules under the autonomic nerveregulation (Garrett, 1987). It is composed of tubular epithelia

that can be divided into two major domains. The distal end isthe secretory unit, the acini, where the primary saliva isproduced. At the proximal part are the ducts, which modify theprimary saliva by absorbing or secreting certain ions, such asNa+, Cl– and HCO3– (Young et al., 1987). By using high-speedconfocal microscopy, we previously studied Ca2+ signalling inthe rat salivary ducts (Yamamoto-Hino et al., 1998) and acini(Takemura et al., 1999). Millisecond analyses demonstratedthat [Ca2+]i responses in the ducts started in some ‘pioneercells’ and then spread to the neighboring cells, showingasynchrony and heterogeneity (Yamamoto-Hino et al., 1998),whereas those in the acini occurred synchronously (Takemuraet al., 1999). These data prompted us to study whether the Ca2+

signalling system in salivary glands is constructed accordingto the tissue architecture.

In the present study, we analysed details of calciumsignalling in the rat parotid gland. We dissociated the glandswith collagenase to obtain morphologically and functionallyintact preparation of acini and ducts (Segawa et al., 1985).These specimens were loaded with fluo-3AM and stimulatedwith calcium mobilising agents carbachol (CCh) and ATP inthe presence or absence of extracellular Ca2+. Changes in[Ca2+]i of acini and ducts were analysed by high-speedconfocal microscopy both at the regional level and at the single

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Organ function requires coordinated multicellularactivities, which may require proper control of cellsignalling dynamics at the supracellular level. By usinghigh-speed confocal microscopy, we studied how calciumsignalling is organised in the dissociated rat parotid gland.Salivary gland function is accomplished primarily by thecompartmentalized epithelial domains, acini and ducts, theformer involved in the production of primary saliva and thelatter involved in its modification. Upon muscarinicstimulation with carbachol, both domains showed anincrease in intracellular free calcium concentration([Ca2+]i) with distinctive spatiotemporal kinetics, asindicated by the fluo-3 fluorescence. Acini respondedinitially, and the ducts followed with a time lag of more than0.3 second. Cells comprising an acinus respondedsynchronously, whereas those in the ducts respondedheterogeneously with respect to the latency period,magnitude of response and the requirement of extracellular

calcium to raise [Ca2+]i. ATP also elicited a non-synchronous [Ca2+]i response in the duct domain, under apattern different from that of carbachol. The synchronousoscillations seen in the acinar domain were madeasynchronous by octanol, an agent known to inhibit gap-junction function. Accordingly, a gap junction component,connexin 32, was immunolocalised predominantly betweenthe acinar cells. Moreover, expression of the type 2 inositol(1,4,5)-trisphosphate receptor [Ins(1,4,5)P3R] washomogeneous in the acinar domain but heterogeneous inthe duct domain. Together, these data suggest that thecalcium signalling system in salivary glands is constructedspecifically according to the tissue architecture.

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Key words: Calcium signalling, Salivary gland, Gap junction,Ins(1,4,5)P3 receptor, Epithelia

Summary

Calcium signalling in tissue: diversity and domain-specific integration of individual cell response insalivary glandsAkihisa Segawa 1,*, Haruo Takemura 2 and Shohei Yamashina 1

1Department of Anatomy, School of Medicine, Kitasato University, 1-15-1 Kitasato, Sagamihara, Kanagawa 228-8555, Japan2Department of Pharmacology, School of Medicine, Sapporo Medical University, South 1, West 17, Sappro 060-8556*Author for correspondence (e-mail: segawa@kitasato-u.ac.jp)

Accepted 20 February 2002Journal of Cell Science 115, 1869-1876 (2002) © The Company of Biologists Ltd

Research Article

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cell level. The possible role of intercellular communicationthrough the gap junction was studied functionally by octanolpretreatment and morphologically by connexin 32immunohistochemistry. The distribution of inositol (1,4,5)-trisphosphate receptor type 2 [Ins(1,4,5)P3R2], a receptormolecule involved in the release of Ca2+ from intracellularstores, was also examined.

Materials and MethodsCell dissociationMale Wistar rats (6-10 weeks) were killed by ether anaesthesia andexsanguination. Parotid glands were cut into small pieces and digestedwith collagenase (CLS4, Worthington Biochemical Corporation,Lakewood, NJ) at 3 mg/ml for 20 minutes at 37°C in Dulbecco’smodified Eagle medium (DMEM) containing 25 mM Hepes withconstant shaking and gentle pipetting. Following digestion, thedissociated glands were washed twice with DMEM and thenincubated with 10 µM fluo-3AM (Molecular Probes, Eugene, OR) for20 minutes at room temperature. After washing, the dissociatedtissues were resuspended in Krebs-Ringer Hepes (KRH) medium (120mM NaCl, 5.4 mM KCl, 1.0 mM CaCl2, 0.8 mM MgCl2, 11.1 mMglucose, 20 mM Hepes) containing 0.2% bovine serum albumin Foromission of extracellular Ca2+, Ca2+-free KRH was supplementedwith 0.2 mM EGTA. The dissociated tissues were able to respond toCCh and ATP for up to 1 hour in KRH at room temperature.

High-speed confocal microscopyThe fluo-3-loaded dissociated tissues were placed on the coverslipcoated with Cell Tak (Collaborative Biomedical Products, Bedford,MA) (Segawa, 1999). They were viewed with a inverted lightmicroscope (Nikon Diaphot 300, Nikon, Tokyo, Japan) equipped witha high-speed confocal microscope (Oz with InterVision software,Noran Instruments Inc., Middleton, WI). For stimulation of thedissociated tissues, 10 µM carbachol (Wako Pure Chemical Industries,Osaka, Japan) or 100 µM ATP (adenosine 5′-triphosphate: SigmaChemical, St Louis, MO) were added directly onto the tissues. Whennecessary, pretreatment with octanol (Wako Pure Chemical Industries)was performed at 3 mM for 3 minutes before addition of CCh. Thespecimens were observed with the plan apo objective lenses ×20, ×40,×60 with NA 0.75, 1.00 and 1.20, respectively. The plane ofapproximately 20-50 µm depth from the coverslip was observed.Confocal images were taken every 8-33 milliseconds. Excitation wasperformed with an Ar/Kr laser at 488 nm and the emission signalswere collected through a 515 nm barrier filter. Pseudo-ratio imagingwas performed by dividing raw fluorescence images by an imageimmediately before addition of CCh or ATP, representing the restingdistribution of fluo-3 fluorescence in the specimens. The pseudo-ratioimages were used for the quantitative measurement.

Scanning electron microscopyDissociated tissues were fixed with 2% glutaraldehyde in 0.1 Mcacodylate buffer, pH 7.2, for 1 hour at 4°C, and post-fixed with 1%OsO4 for 1 hour at 4°C. They were dehydrated through a graded seriesof ethanol, dried in a critical point dryer and coated with Au/Pd at 4nm by a sputter coater. The specimens were observed by a Hitachi S-4500 field emission scanning electron microscope (Hitachi Co. Ltd.,Hitachi, Japan) operated at 5 kV.

ImmunohistochemistryPieces of parotid glands were fixed with 10% formalin for 1 hour at4°C, and cut in a cyrostat at 20 µm. The sections were pretreated with1% bovine serum albumin (BSA) in phosphate buffered saline (PBS)

at 4°C for 1 hour. They were then treated with mouse monoclonalantibodies against Ins(1,4,5)P3R2 (IgGIIa, KM1083) (Sugiyama etal., 1994) and connexin 32 (IgG1, Zymed Laboratories, Inc., CA) at4°C for overnight. Control specimens were incubated solely withBSA-PBS. After washing with PBS, the sections were treated withRITC-labelled anti-mouse IgGIIa and FITC-labelled anti-mouseIgG1, mounted with anti-fader reagent Fluoro-Guard (Bio-Rad Lab.,CA) and observed under the conventional confocal microscope (Bio-Rad MRC 1024, Nippon Bio-Rad Lab., Tokyo, Japan). Serial confocalimages were taken along the Z-axis at a distance of 1.5 µm and thenreconstructed into the ‘through focus’ image by summating eachimage.

ResultsMorphology of the dissociated glandsDigestion of rat parotid glands with collagenase yieldedrelatively large cell clumps. Scanning electron microscopy ofthese specimens (Fig. 1) showed that the ducts exhibit theelongated tubular structure, whereas acini reveal the roundstructure located at the end of the duct segment. A lot of aciniwere grouped together, so that the overall morphology of acini-duct complex revealed the ‘bunch of grape’-like appearance.Comparable images were obtained by using confocalmicroscopy with transmitted light imaging (Fig. 2B).

Changes in [Ca2+]i in acini and ducts in response toCChFollowing application of CCh to the tissues, both acinar andductal segments exhibited a rise in fluo-3 fluorescenceintensity, indicating a rise in [Ca2+]i. Fluo-3 loading wasefficiently applied in the outer layers of cell clumps so that theresponse of acini was defined in those located in the peripheryof the specimen. Fig. 2 shows the typical response. Acini were

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Fig. 1.Dissociated rat parotid glands shown by scanning electronmicroscopy. A lot of acini (a) are grouped together and located at theend of the duct (d). Bar, 30 µm.

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always the first to respond (0.32-2.5 seconds after CCh;mean=0.76 seconds, n=28), and the ducts followed (0.87-3.1seconds after CCh, mean=1.70 seconds, n=12) with a time lagof 0.38-2.0 seconds (mean=1.0 seconds, n=12). We did notobserve any propagation of Ca2+ waves from acini to the ducts.Instead, within the ducts, the rise in [Ca2+]i started in some‘pioneer cells’ and then spread to the neighboring cells (Fig.2C), as described previously in the ducts of submandibularglands (Yamamoto-Hino et al., 1998).

[Ca2+]i response to CCh and ATP in the acini and ducts:analyses at the regional levelThe response of acini and ducts to CCh and ATP was analysedby the consecutive application of each reagent in the presence

or absence of extracellular Ca2+. Acini and ducts exhibited adistinct response pattern. A typical example is shown in Fig. 3(n=14).

When CCh was applied to the specimens in the absence ofextracellular Ca2+ (Fig. 3C), [Ca2+]i increased in the acinararea but not in the ductal area. However, addition of Ca2+ tothe perfusion medium resulted in a rise in [Ca2+]i in the ductalarea (Fig. 3D). After washing the specimen thoroughly withCa2+-free KRH, and adding ATP in the absence of extracellularCa2+, the acinar response was weak or negligible (Fig. 3E). Bycontrast, the ductal area exhibited a rise in [Ca2+]i, althoughthe response was transient and [Ca2+]i soon declined. WhenCa2+ was added to the perfusion medium under theseconditions, the ductal area exhibited a rise in [Ca2+]i andshowed a sustained plateau (Fig. 3F).

Fig. 2.High-speed confocal microscopy ofdissociated rat parotid gland showing calciumsignalling of acini and ducts in response toCCh. (A) Fluorescence image. (B) Mergedimage of transmitted light and fluorescenceimage shown in A. (C) Quantitativemeasurement of the fluorescent changescalculated from the pseudo-ratio imaging.Data were obtained from the correspondinglynumbered squares shown in A.(D,E) Sequential changes of fluo-3fluorescence following addition of CCh.Acinar response precedes the ductal response.An arrow denotes ‘pioneer’ cells from whichthe ductal response is initiated. CCh wasapplied at time zero. Confocal images weretaken every 8 milliseconds.

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When the application protocol was reversed and ATP wasapplied prior to CCh, the acinar and ductal responses weresimilar to those described above (not shown). Thus thesequence of application of ATP and CCh did not seem to alterthe pattern of acinar and ductal response.

[Ca2+]i response to CCh and ATP in the acini and ducts:analyses at the individual cell levelAnalyses of the individual cell response (Fig. 4) were carriedout using the same preparation as that shown in Fig. 3. Thepattern of acinar cell response was similar to that measured atthe regional level. As described below, each acinar cellexhibited an almost homogenous pattern of response. Bycontrast, ductal cells exhibited an heterogeneous pattern ofresponse. Although the gross measurement of the specimenfailed to detect the ductal response to CCh in the absence of

extracellular Ca2+ (Fig. 3D), a few duct cells were found torespond under this condition (Fig. 4B). Addition of Ca2+ to theperfusion medium caused [Ca2+]i levels to rise in most ductcells (Fig. 4C) but several cells showed a weak or negligibleresponse; ATP stimulation caused [Ca2+]i levels to rise in theseduct cells (Fig. 4D). In the absence of extracellular Ca2+, theresponse of each cell was transient, but in the presence ofextracellular Ca2+ it exhibited a sustained plateau (Fig. 4E).The magnitude of the response and the latency period differedfrom one cell to another.

Effect of octanol on the synchronised [Ca2+]i response inthe acini10 µM CCh treatments often caused an oscillation in the[Ca2+]i response in acinar cells (Fig. 5A,C). The oscillatory

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Fig. 3. [Ca2+]i dynamics in ducts and acini in response to the consecutive application of CCh and ATP in the presence or absence ofextracellular Ca2+. (A) Transmitted light image. (B) Fluorescence image. Ductal area (red) and acinar area (blue) were selected and measured.The application protocol was as follows: (C) CCh in the absence of extracellular Ca2+. (D) CCh in the presence of extracellular Ca2+.(E) ATP in the absence of extracellular Ca2+. (F) ATP in the presence of extracellular Ca2+. Arrows denote the application time of each reagent.

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response occurred synchronously among the cells within anacinus. Gap junctions have been shown to be involved in thesynchronised [Ca2+]i response in several cell systems (Staufferet al., 1993; Guerineau et al., 1998). We thus analysed theeffects of octanol, a gap junction inhibitor, on the acinar cellresponse. Following pretreatment of the specimen with octanolfor 3 minutes, the response became asynchronous (Fig. 5B,D).

Distribution of connexin 32 and Ins(1,4,5)P3R2 in theacini and ductsThe involvement of gap junctions in the observed [Ca2+]iresponse was further analysed by immunohistochemistry usingconnexin antibody. As shown in Fig. 6A, positiveimmunofluorescence was found predominantly in the acinararea, whereas the ductal area was mostly devoid of positivesignals. An exceptional area was the distal end of theintercalated ducts, in which a positive reaction was sometimesdetected. In an attempt to discover the intracellular moleculeresponsible for the supracellular regulation of Ca2+ signalling,we also observed the distribution of Ins(1,4,5)P3R2.Ins(1,4,5)P3R2 immunofluorescence was detected along theapico-lateral area of acinar cells (Fig. 6B). This area

corresponds to the intercellularcanaliculi, a specialised formof the lumen in the ratparotid acini (Takemura et al.,1999). In the ducts, strongimmunofluorescence wasobserved along the apical area;many cells showed positivesignals, but some cells werefound to exhibit weak ornegative fluorescence. Thus, incontrast to the acini, thedistribution of Ins(1,4,5)P3R2in the ducts exhibits hetero-geneity. The merged image(Fig. 6C) clearly showed thedifferent immunolocalisation ofconnexin 32 and Ins(1,4,5)P3R2in the acini and ducts. Controlspecimens treated withoutprimary antibodies did notreveal any specific staining.

DiscussionExocrine organs function isprimarily achieved by thecompartmentalized epithelialdomains, acini and ducts, whichalso exhibit characteristicmorphological specialisation(Young and van Lennep, 1978;Pinkstaff, 1980). In the presentstudy, we found that the patternof Ca2+ signalling is alsodistinct between acini and ductsin the rat parotid gland inresponse to CCh and ATP

stimulation. Acinar cells within an acinus respondedsynchronously and behaved as a functional syncitium, whereasductal cells responded heterogeneously, showing diversesignalling kinetics. In addition, the molecular distribution ofconnexin 32 and Ins(1,4,5)P3R2 was distinct between the twodomains. Collectively, these data indicate that the Ca2+

signalling system in the rat parotid gland is organised inparallel with the tissue architecture.

Many investigators have observed the elevation of [Ca2+]i inthe acini and ducts of salivary glands in response to CCh andATP stimulation (Valdez and Turner, 1991; Gromada et al.,1993; Dinudom et al., 1993; Hurley et al., 1993; Soltof et al.,1993; Hurley et al., 1994; Jorgensen et al., 1995; Xu et al.,1996; Lee et al., 1997a; Lee et al., 1997b; Tojyo et al., 1997a;Tojyo et al., 1997b; Bird et al., 1998). As discussed previously,the time course of [Ca2+]i elevation proceeds rapidly, makingit difficult to clarify the spatiotemporal details of Ca2+

signalling dynamics in the tissue (Takemura et al., 1999). High-speed confocal microscopy of dissociated glands allowedanalysis of the changes in [Ca2+]i of the order of a millisecond,with precise measurement of acinar and ductal responses fromthe supracellular to the subcellular level.

CCh stimulation caused a rise in [Ca2+]i in the acini before

Fig. 4. [Ca2+]i dynamics in ducts andacini measured at the individual celllevel. (A) Fluorescence image. Cellularareas were selected from the acinus(yellow) and duct (green, orange, blue,red) and used for the quantitativemeasurement. (B) CCh in the absenceof extracellular Ca2+. (C) CCh in thepresence of extracellular Ca2+. (D) ATPin the absence of Ca2+. (E) ATP in thepresence of extracellular Ca2+. Arrowsdenote the application time of eachreagent.

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the ducts (Fig. 2). We did not observe any Ca2+ waves fromacini to ducts; thus direct propagation of Ca2+ signals fromacini to ducts seems unlikely. 3D propagation of Ca2+ signalsout of and into the plane of focus might be another explanation.Since CCh triggers fluid secretion from the acini, it is alsopossible that acinar fluid elicited the ductal response from theluminal side (Xu et al., 1996; Lee et al., 1997a; Lee et al.,1997b). However, it was noted that the presence of acini wasnot obligatory to initiate the ductal response (Fig. 3). Thisindicates that the ductal system has its own intrinsic controlsystem for [Ca2+]i mobilisation. Based on the fact that the Ca2+

response started in certain cells from which the Ca2+ wave thenpropagated to neighboring cells (Fig. 2), it is likely that thesecells act as ‘pioneer’ or ‘pacemaker’ cells to control theregional ductal response (Yamamoto-Hino et al., 1998).

The pattern of Ca2+ response to CCh and ATP in thepresence or absence of extracellular Ca2+ was considerablydifferent in acini and ducts (Fig. 3). The acinar domainresponded well to CCh but not to ATP, and the responsivenessto CCh did not require the presence of extracellular Ca2+. Thisindicates that the acinar domain raises [Ca2+]i by the releaseof Ca2+ from the intracellular pool upon muscarinic receptorstimulation. Purinergic receptor stimulation seems to have littleeffect on both the intra- and extracellular mechanisms of Ca2+

mobilisation in the acinar domain. By contrast, the ductaldomain responded both to CCh and ATP, but in the absence ofextracellular Ca2+ the responsiveness to CCh was abolished.This suggests that the observed ductal response to CCh occursmostly through the entry of Ca2+ from the extracellular fluid.

Alternatively, the release of Ca2+ from the intracellular storemight require the presence of extracellular Ca2+ in the case ofCCh stimulation. In the same duct, ATP triggered both therelease of Ca2+ from the intracellular pool (Fig. 3E) and theCa2+ entry from the extracellular fluid (Fig. 3F). However, itshould be pointed out that the response of individual duct cellsvaried from one to the other (Fig. 4). This indicates that theintracellular signalling cascade downstream of receptoractivation is diverse among the duct cells.

Extensive studies have elucidated that Ca2+ signallinginvolves a molecular cascade of reactions, includingphosphoinositide turnover and the activation ofreceptors/channels present on the plasma membrane and theintracellular stores, leading to the elevation of [Ca2+]i either byentry of Ca2+ from the extracellular fluid or release of Ca2+

from the intracellular stores (Berridge et al., 2000; Meldolesiand Pozzan, 1998; Michikawa et al., 1996). Our findingssuggest that such subcellular mechanisms are not equallyactivated in the cells of duct domain. Since salivary ducts areknown to be composed of heterogeneous cell populations (Satoand Miyoshi, 1988; Sato and Miyoshi, 1998), it is possible thatthe heterogeneity of Ca2+ signalling reflects the heterogeneityof cell types. Whether this involves the heterogeneity of ERCa2+ stores (Meldolesi and Pozzan, 1998) remains to beelucidated. Further studies are needed to clarify the biologicalsignificance of diverse Ca2+ signalling occurring in the ductdomain.

In contrast to the ducts, acini exhibited a synchronous[Ca2+]i response (Fig. 5). Gap junctional communication has

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Fig. 5.Effects of octanol on the synchronised [Ca2+]i response evoked by CCh in the acinus. (A,C) Untreated control. (B,D) Octanolpretreatment for 3 minutes. Data for the quantitative measurements were obtained from the correspondingly colored squares shown in A and B.Arrows denote the application time of CCh.

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been implicated in synchronised intercellular Ca2+ signalling(Stauffer et al., 1993; Guerineau et al., 1998). In mammaliansalivary glands, the presence of gap junctions between theacinar cells has been shown by electron microscopy (Deweryand Barr, 1964; Hand, 1972; Nagato and Tandler, 1986) andconnexin immunohistochemistry (Hirono et al., 1995; Lee etal., 1998; Shimono et al., 2000). Although there are debates asto whether gap junctions are present between the duct cells(Dewery and Barr, 1964; Hirono et al., 1995; Lee et al., 1998),our findings indicate that the ductal domain is mostly devoidof gap junctions and, if present, they are located at the distalend of the intercalated ducts close to the acini (Fig. 6). Theimmunofluorescence spots observed in the intercalated ductarea presumably correspond to the gap junctions betweenmyoepithelial cells (Lee et al., 1998) or intercalated duct cellsthemselves (Dewery and Barr, 1964; Hirono et al., 1995).Whatever the explanations are, previous data and our studiesindicate that the acinar domain is the principal site where gapjunctions are present in salivary glands. The abolishment by

octanol of the synchronised Ca2+

response in the acini (Fig. 5)strongly suggests that the acinardomain acts as the functionalsyncitium through the gapjunction.

Immunohistochemical distri-bution of Ins(1,4,5)P3R2 washomogeneous in the acini butheterogeneous in the ducts (Fig.6). The distribution ofIns(1,4,5)P3R2 in the intercellularcanaliculi area of salivary acinihas been observed by us(Yamamoto-Hino et al., 1998;Takemura et al., 1999) and byothers (Lee et al., 1997a).Heterogeneous expression of

Ins(1,4,5)P3Rs in salivary ducts has also been noted by ourprevious study (Yamamoto-Hino et al., 1998). Heterogeneousexpression of Ins(1,4,5)P3Rs has also been observed in airwayepithelia (Sugiyama et al., 1996), kidney collecting ducts(Monkawa et al., 1998) and gastrointestinal epithelia(Matovick et al., 1996), and thus is considered as the commonfeature in tubular epithelia. Since Ins(1,4,5)P3Rs play key rolesin the release of Ca2+ from the intracellular stores (Berridge etal., 2000; Meldolesi and Pozzan, 1998; Michikawa et al.,1996), the particular tissue distribution of Ins(1,4,5)P3R2 islikely to determine the type of Ca2+ signalling: either acinar(synchronised) or ductal (non-synchronised).

Our observations indicate that acini and ducts create distinctlocal Ca2+ signalling communities. Since acini and ductsconstitute the elementary units of exocrine organs, theorganised Ca2+ signalling might have some impact on themechanisms underlying the harmonised organ function.Coordination of Ca2+ signalling according to the tissue unit hasbeen observed in the liver (Nathanson et al., 1995; Robb-

Fig. 6. Immunofluorescencemicrographs showing the expressionof connexin 32 and Ins(1,4,5)P3R2 inthe rat parotid gland. (A) Connexin32. (B) Ins(1,4,5)P3R2. (C) Mergedimage. (D) Schematic representation.The ductal (d) area is devoid ofconnexin 32 immunofluorescence.The distal end of the intercalatedducts (i) exhibits a fewimmunofluorescence spots. Theexpression of Ins(1,4,5)P3R2 in theducts is strong in the apicalcytoplasm of some cells (arrows), butweak or negligible in other cells(arrowheads). In the schematicrepresentation, Ins(1,4,5)P3R2 in theduct cells is localised in the apicalvesicles based on the immunoelectronmicroscopic observation (Yamamoto-Hino et al., 1998). ‘Through-focus’images reconstructed from nine serialconfocal sections taken at a distanceof 1.5 µm. Bar, 50 µm.

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Gaspers and Thomas, 1995), and suggested to facilitate diversecells to perform the integrative, organ-level response. DistinctCa2+ signalling among different domains (i.e. the acini andducts), may help to develop the versatility needed for the organto perform its complex functions.

We thank T. Sugiyama (Kyowa Hakko Kogyo Co Ltd, Tokyo,Japan) and K. Mikoshiba (The Institute of Medical Science,University of Tokyo, Tokyo, Japan) for the gift of monoclonalantibody against Ins(1,4,5)P3R2. We are also grateful for M.Miyakawa (Hitachi High-Technologies Co Ltd., Atsugi, Japan) for theassistance in high-speed confocal microscopy, and for the staff in theLight and Electron Microscopy Laboratory Center (KitasatoUniversity, Sagamihara, Japan) for assistance in scanning electronmicroscopy. This work was supported in part by the Grant-in-Aid forScientific Research (C) from Japan Society for the Promotion ofScience (No.12671781).

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