Histamine stimulates ion transport by dog pancreatic duct epithelial cells through H1 receptors

Toan D. Nguyen, Charles N. Okolo, and Mark W. Moody

Department of Medicine, University of Washington, and Veterans Affairs Puget Sound Health Care System, Seattle, Washington 98108

    ABSTRACT
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Histamine affects pancreatic secretion, but its direct action on ion transport by pancreatic duct epithelial cells (PDEC) has not been defined. We now characterize the secretory effects of histamine on cultured, well-differentiated, and nontransformed dog PDEC. Histamine stimulated, in a concentration-dependent manner (1-100 µM), a cellular 125I- efflux that was inhibited by 500 µM 5-nitro-2-(3-phenylpropylamino)benzoic acid, 2.5 mM diphenylamine-2-carboxylate, and 500 µM DIDS and thus mediated through Ca2+-activated Cl- channels. Histamine-stimulated 125I- efflux was 1) inhibited by 100 µM diphenhydramine, an H1 receptor antagonist, 2) resistant to 1 mM cimetidine, an H2 receptor antagonist, 3) not reproduced by 1 mM dimaprit, an H2 agonist, and 4) inhibited by 50 µM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid-AM, a Ca2+ chelator, suggesting that it was mediated through H1 receptors acting via increased cytosolic Ca2+. Histamine also stimulated a 86Rb+ efflux that was sensitive to 100 nM charybdotoxin and thus mediated through Ca2+-activated K+ channels. When PDEC monolayers were studied in Ussing chambers, a short-circuit current of 21.7 ± 3.1 µA/cm2 was stimulated by 100 µM histamine. This effect was inhibited by diphenhydramine but not cimetidine, was not reproduced with dimaprit, and was observed only after serosal addition of histamine, suggesting that it was mediated by basolateral H1 receptors on PDEC. In conclusion, histamine, acting through basolateral H1 receptors, activates both Ca2+-activated Cl- and K+ channels; in this manner, it may regulate PDEC secretion in normal or inflamed pancreas.

chloride channels; potassium channels; Ussing chamber

    INTRODUCTION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

SINCE ITS DISCOVERY at the beginning of the century, the physiological and pathological role of histamine has been expanding. It is released from nerve endings to act as a neurotransmitter, from endocrine cells to act as a hormone, and from mast cells to act as a mediator of inflammation. These functions are mediated through the following three specific receptors: H1 receptors acting through phospholipase C to increase cytosolic free Ca2+ concentration ([Ca2+]i), H2 receptors activating the cAMP cascade, and presynaptic H3 receptors inhibiting histamine synthesis and release (3, 9). In the digestive system, histamine stimulates secretion in certain cells [e.g., parietal cells and colonocytes (23)], whereas it inhibits secretion in others [e.g., duodenal cells and D cells (4, 22)]. The effect of histamine on pancreatic secretion has been examined using different experimental systems involving anesthetized animals, isolated pancreas, pancreatic lobules, or pancreatic segments. Depending on the model studied, histamine interacts with either H1 receptors to stimulate pancreatic exocrine secretion, with H2 receptors to stimulate or inhibit secretion, or with H3 receptors to inhibit secretion (6, 8, 16-19). This varied response may reflect the different species studied; it also illustrates the complex components and determinants of pancreatic exocrine secretion.

Pancreatic exocrine function consists of the secretion of digestive enzymes, mediated by pancreatic acinar cells, and the secretion of fluid and electrolytes, mainly bicarbonate, mediated by pancreatic duct epithelial cells (PDEC). Most of the reports mentioned above characterized the effect of histamine on the final product of these two secretory processes, the pancreatic juice collected by cannulation. Although acinar and ductal function are generally reflected by the enzyme content and volume of pancreatic juice, pancreatic duct cannulation does not allow clear distinction between histamine effects on acinar cells and PDEC. The investigations using pancreatic lobules, on the other hand, focused mainly on the secretion of enzymes by acinar cells and did not address the effect of histamine on PDEC.

Of further note, because the pancreatic tissue studied in these reports contains many cell types, it is unclear whether the observed effects result from the direct interaction between histamine and the secretory cell or whether they are indirectly mediated through other cells. Indeed, histamine has been shown to interact with presynaptic H3 receptors on intrinsic pancreatic nerves to inhibit pancreatic secretion (6).

Thus the direct effect of histamine on PDEC is yet undefined. Compared with acinar cells, studies of the secretory function of PDEC have been hampered by the lack of a practical model for these cells (11). Oda et al. (15) recently developed a method to culture well-differentiated dog PDEC without transformation; we recently demonstrated that these cells express functional cAMP- and Ca2+-activated Cl- channels (14) and Ca2+-activated K+ channels (14a). In this report, these cells were used to examine the direct effects of histamine on ion transport by PDEC. Our aims were to 1) determine the effects of histamine on ion transport pathways, such as K+ and Cl- channels, 2) characterize the subtype and localization of the responsible histamine receptors, and 3) define the signal-transduction pathway mediating this effect.

    MATERIALS AND METHODS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Chemicals and reagents. Histamine, diphenhydramine, cimetidine, dimaprit, DIDS, charybdotoxin, and tissue culture medium and supplements were from Sigma (St. Louis, MO), and 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM was from Calbiochem (San Diego, CA). Diphenylamine-2-carboxylate (DPC) was from Fluka (Ronkonkoma, NY), and 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) was from Research Biochemicals (Natick, MA). Na125I (16 mCi/mg iodide) was purchased from Amersham (Arlington Heights, IL), and 86RbCl (4.66 mCi/mg rubidium) was from NEN (Boston, MA).

Cell culture. Dog PDEC, isolated from the accessory pancreatic duct of a dog, were cultured in Eagle's MEM, containing 10% fetal bovine serum, 2 mM L-glutamine, 20 mM HEPES, 100 IU/ml penicillin, 100 µg/ml streptomycin, 5 µg/ml bovine insulin, 5 µg/ml human transferrin, and 5 ng/ml sodium selenite, and placed on Transwell inserts (Costar, Cambridge, MA) coated with 0.5 ml of a 1:1 solution of Eagle's MEM-Vitrogen (Collagen, Palo Alto, CA). The Transwell inserts allow the PDEC to share a common medium with a feeder layer of human gallbladder myofibroblasts cultured on the bottom of the well in which the insert is suspended. These myofibroblasts are believed to secrete the growth factors necessary for maintaining and propagating the well-differentiated PDEC. These PDEC secrete mucin (15) and express functional Ca2+-activated Cl- channels, cAMP-activated Cl- channels corresponding to the cystic fibrosis transmembrane conductance regulator (CFTR; see Ref. 14), and Ca2+-activated K+ channels (14a). The cells used in this report were between passages 9 and 30.

Efflux studies. In many investigations of Cl- transport, 125I- is the preferred substitute marker for 36Cl- because of its high specific activity, favorable selectivity with most Cl- channels, relative low cost, and shorter half-life. The use of cellular 125I- efflux to study Cl- channel activation has been validated (21) and was effective for characterizing Cl- channels on PDEC (14). Similarly, 86Rb+ has been used as a marker for K+ for studies of K+ channels (21).

PDEC were grown to confluence on Transwell inserts as described above. The membranes and overlying cells were excised from the insert and washed two times with 1 ml of efflux buffer consisting of (in mM) 140 NaCl, 4.7 KCl, 1.2 CaCl2, 1 MgCl2, 10 glucose, and 10 HEPES, pH 7.4. Radioactive tracer loading was achieved by incubating the cells for 45 min at 37°C with 1.5 ml of efflux buffer containing either ~2 µCi/ml Na125I or ~1 µCi/ml 86RbCl. The cells were next washed four times with 2 ml of isotope-free buffer. Isotope efflux was measured by sequential addition and removal of 1 ml of isotope-free buffer at 15-s intervals for a 5-min period. To establish baseline efflux, no secretagogue was added for the first minute; in the remaining 4 min, the secretagogue tested was included in the buffer. When inhibitors were tested, they were added at the beginning of the monitoring period. The radioactivity of these sequential samples and the radioactivity associated with the cells at the end of the experiment were measured with a gamma counter (Isodata 120; ICN, Huntsville, AL) for 125I- and with a liquid scintillation counter (Tri-Carb model 1600TR; Packard, Meriden, CT) for 86Rb+.

The radioactivity contained in the cells at a particular time point was calculated as the sum of the radioactivities released in subsequent efflux samples and remaining in the cells at the end of the experiment. The efflux rate coefficient (r) for a certain time interval was also calculated using the formula
<IT>r</IT> = [ln(R<SUB>1</SUB>) − ln(R<SUB>2</SUB>)]/(<IT>t</IT><SUB>1</SUB> − <IT>t</IT><SUB>2</SUB>)
where R1 and R2 are the percentage of counts initially loaded remaining in the cells at times t1 and t2.

In certain experiments, peak stimulated efflux rate coefficients were compared. These peak rate coefficients were calculated by subtracting the baseline rate, the lowest efflux rate before the addition of histamine, from the peak stimulated efflux rate, the highest efflux rate after the addition of histamine.

Ussing chamber studies. Confluent monolayers of PDEC and membrane support were excised from the Transwell insert and mounted in modified Ussing chambers with an aperture area of 0.95 cm2 (13). Both sides of the monolayer were bathed in a Ringer solution (in mM: 115 NaCl, 1.2 CaCl2, 1 MgCl2, 0.4 KH2PO4, 2.4 K2HPO4, 25 NaHCO3, and 10 glucose), warmed to 37°C with a circulating water jacket, and gently mixed and aerated with a constant inflow of 95% air-5% CO2. During secretory studies, spontaneous tissue potential differences were short-circuited by an automatic voltage clamp (model DVC-1000; WPI, Sarasota, FL) with Ag-AgCl2 electrodes, and the corresponding short-circuit current (Isc) was recorded continuously using a model MP100 analog-to-digital converter and the Acknowledge 2.0 software program (BioPak Systems, Goleta, CA). Instrument calibration was performed using membranes devoid of cells. In most cases, the magnitude of the Isc response was estimated using peak stimulated Isc increases, corrected for baseline activity; in some instances, the area under the curve was also calculated using the Acknowledge software.

Because the amplitude of the Isc response can vary with time (probably reflecting growth conditions and cell passage), comparative experiments used cell monolayers cultured under the same conditions and studied concurrently.

Statistics. All experiments were repeated at least three times, and results are expressed as means and SE. Comparisons were performed using unpaired two-tailed Student's t-test, and the Stat View 512+ software (Abacus Concepts, Calabasas, CA) was used to determine the corresponding P values.

    RESULTS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Iodide efflux studies. The effect of different concentrations of histamine on 125I- efflux from preloaded PDEC was first evaluated. Marked increases in efflux were observed with concentrations of histamine >= 10 µM; these increases were rapid and transient, reaching a peak 30 s after the addition of histamine (Fig. 1A). The peak efflux rate coefficients were, respectively, 0.25 ± 0.02, 0.54 ± 0.05, 0.63 ± 0.04, and 0.69 ± 0.09/min for concentrations of histamine of 1 µM, 10 µM, 100 µM, and 1 mM (n = 6 experiments). The corresponding efflux rate coefficient in the absence of histamine was 0.23 ± 0.02/min. Thus histamine stimulated an increased 125I- efflux in a concentration-dependent manner, with a maximal effect at ~100 µM.


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Fig. 1.   Stimulation of 125I- efflux by histamine. 125I- efflux from pancreatic duct epithelial cells (PDEC) was determined as detailed in MATERIALS AND METHODS, and the efflux rate coefficient was calculated. Means and SE from 6 (A) or 3 (B-D) experiments are shown. A: after 1 min of baseline determination, histamine was added to the final concentrations shown. B: after 1 min of baseline determination, 100 µM histamine was added in the presence (bullet ) or absence (open circle ) of 500 µM 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB). C: after 1 min of baseline determination, 100 µM histamine was added in the presence (bullet ) or absence (open circle ) of 2.5 mM diphenylamine-2-carboxylate (DPC). D: after 1 min of baseline determination, 100 µM histamine was added in the presence (bullet ) or absence (open circle ) of 500 µM DIDS.

To determine that the 125I- efflux was mediated through activated Cl- channels, the effects of previously established inhibitors of Cl- channels on these PDEC were evaluated. The 125I- efflux stimulated by 100 µM histamine was completely abolished by 500 µM NPPB (Fig. 1B) and by 2.5 mM DPC (Fig. 1C). In addition, it was also markedly inhibited by 500 µM DIDS, from a control peak efflux rate coefficient of 0.82 ± 0.04/min to a peak rate of 0.30 ± 0.03/min in the presence of DIDS (mean peak stimulated efflux rate coefficient: 0.588 ± 0.049 vs. 0.205 ± 0.026/min, 65% inhibition, P = 0.002 by unpaired 2-tailed t-test with 4 degrees of freedom; Fig. 1D). This inhibitory profile suggests that the I- efflux is mediated through Ca2+-activated Cl- channels (14).

The receptor subtype mediating histamine action was next defined using different histamine receptor agonists and antagonists. As shown in Fig. 2A, the 125I- efflux stimulated by 100 µM histamine was abolished by 100 µM diphenhydramine, a specific H1 antagonist. On the other hand, the histamine-stimulated 125I- efflux was not affected by a high concentration (1 mM) of cimetidine, an H2 antagonist (Fig. 2B). In addition, the effect of histamine was not reproduced by a high concentration (1 mM) of the specific H2 agonist, dimaprit (Fig. 2C). In the aggregate, this profile suggests that the stimulatory effect of histamine was mediated by the H1, and not H2, histamine receptor subtype.


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Fig. 2.   Mediation of histamine effect by H1 receptors and increased cytosolic Ca2+ concentration ([Ca2+]i). 125I- efflux from PDEC was determined as detailed in MATERIALS AND METHODS, and the efflux rate coefficient was calculated. Means and SE from 3 experiments are shown. A: after 1 min of baseline determination, 100 µM histamine was added in the presence (bullet ) or absence (open circle ) of 100 µM diphenhydramine. B: after 1 min of baseline determination, 100 µM histamine was added in the presence (bullet ) or absence (open circle ) of 1 mM cimetidine. C: after 1 min of baseline determination, 1 mM dimaprit (bullet ) or vehicle (H2O) was added. D: after 1 min of baseline determination, 100 µM histamine was added in the presence (bullet ) or absence (open circle ) of 50 µM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM.

Because histamine effects mediated through H1 receptors are coupled to increased [Ca2+]i, the signal-transduction pathway responsible for histamine-stimulated I- efflux was studied using BAPTA-AM, a membrane-permeant Ca2+ chelator. Once BAPTA-AM is loaded into PDEC, the ester bond is cleaved by cytosolic esterase, trapping the active chelator BAPTA intracellularly and depleting [Ca2+]i. When PDEC were pretreated with BAPTA-AM for 45 min, the increased I- efflux produced by histamine was abolished (Fig. 2D). This inhibition suggests that the effect of histamine is dependent on increased [Ca2+]i.

86Rb+ efflux studies. Using 86Rb+ as a marker for K+, we demonstrated the presence on PDEC of Ca2+-activated K+ channels (14a). These channels are likely to be activated by histamine, which stimulates an increased [Ca2+]i. The effects of histamine on the cellular efflux of 86Rb+ were therefore evaluated. Histamine stimulated an increased 86Rb+ efflux from PDEC, which reached a peak rate coefficient of 0.158 ± 0.003/min (peak stimulated efflux rate coefficient: 0.120/min after baseline correction, n = 3) 30 s after the addition of 100 µM histamine (Fig. 3A).


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Fig. 3.   Stimulation of 86Rb+ efflux by histamine. 86Rb+ efflux from PDEC was determined as detailed in MATERIALS AND METHODS, and the efflux rate coefficient was calculated. A: after 1 min of baseline determination, 100 µM histamine (bullet ) or vehicle (H2O) (open circle ) was added. B: after 1 min of baseline determination, 100 µM histamine was added in the presence (bullet ) or absence (open circle ) of 100 nM charybdotoxin.

To verify that this efflux is mediated through activated K+ channels, the effect of charybdotoxin, an inhibitor of Ca2+-activated K+ channels in these cells, was evaluated. As shown in Fig. 3B, 100 nM charybdotoxin inhibited the histamine-stimulated 86Rb+ efflux by 86%. Indeed, the mean peak stimulated efflux rate coefficient was 0.016 ± 0.001/min for charybdotoxin-treated cells vs. 0.111 ± 0.004/min for control untreated cells (P = 0.0001, 2-tailed unpaired t-test with 4 degrees of freedom).

Ussing chamber studies. An advantage associated with these PDEC is their ability to grow as monolayers of polarized cells with tight junctions, generating a transepithelial electrical resistance adequate for studies in Ussing chambers. Using this experimental system, we can localize the H1 receptors to either the apical or basolateral side of PDEC and characterize the effects produced by the sequential addition of antagonists and agonists.

Confluent PDEC monolayers were mounted in Ussing chambers; in this system, the serosal compartment contains the buffer in contact with the filter and basolateral membrane of the cell, whereas the luminal compartment contains the buffer in contact with the apical membrane. The addition of 100 µM histamine to both compartments produced a quick increase in the Isc that reverted to normal over 5 min; occasionally, a small shoulder was also observed (Fig. 4A). Because the amplitude of the Isc can vary with time (probably reflecting growth conditions and cell passage), the average peak Isc increase (corrected for baseline Isc) from 14 different experiments was calculated; it was 21.7 ± 3.1 µA/cm2.


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Fig. 4.   Effect of histamine on net electrogenic ion transport across PDEC monolayers mounted in Ussing chambers. Monolayers of confluent PDEC were mounted in Ussing chambers as described in MATERIALS AND METHODS, and short-circuit current (Isc) was measured (µA/filter of 0.95 cm2). In A and B, both traces were derived from monolayers cultured and studied at the same time. Experiments were repeated 3 times. A: after 5 min of baseline determination, histamine (100 µM, tracing A) or the vehicle (H2O, tracing B) was added. Because of the variability of the amplitude of the Isc response at different times, the Isc increase in this tracing, 40 µA/filter, is larger than the average response of 21 µA/filter obtained from 14 determinations; this response is otherwise typical of the stimulatory effect of histamine. B: after 5 min of baseline determination, 100 µM histamine was added to either the serosal compartment, contiguous with the basolateral membrane, or the luminal compartment, contiguous with the apical membrane of PDEC. Exclusive response to the serosal addition of histamine was verified in all 3 experiments.

The localization of the histamine receptors on PDEC is clarified in Fig. 4B, where 100 µM histamine was added to either the serosal or luminal compartment of the Ussing chamber. Histamine stimulated an Isc increase only when added to the serosal compartment, but not when added to the luminal compartment. This finding suggests that expression of the histamine receptors is limited to the basolateral side of PDEC.

The receptor subtype was also verified using this system. As shown in Fig. 5A, addition of 100 µM diphenhydramine, the H1 antagonist, by itself did not affect the Isc. However, it fully inhibited the subsequent Isc response to 100 µM histamine. In additional experiments, we observed that this inhibitory effect was reversible: histamine stimulated an Isc increase from cells that were washed after treatment with diphenhydramine (data not shown).


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Fig. 5.   Mediation of histamine effect in Ussing chambers by H1 receptors. Monolayers of confluent PDEC were mounted in Ussing chambers as described in MATERIALS AND METHODS, and Isc was measured (µA/filter of 0.95 cm2). In A-D, both traces were derived from monolayers cultured and studied at the same time. A: after 5 min of baseline determination, histamine was added to a final concentration of 100 µM in the absence (tracing A) or presence (tracing B) of 100 µM diphenhydramine. Complete inhibition of histamine-stimulated Isc increase by diphenhydramine was verified in 4 experiments. B: after 5 min of baseline determination, histamine was added to a final concentration of 100 µM in the presence (tracing A) or absence (tracing B) of 1 mM cimetidine. Absence of significant inhibition was shown in 3 experiments (see text for details). C: after 5 min of baseline determination, 100 µM histamine (tracing A) or 1 mM dimaprit (tracing B) was added; after an additional 15 min, 100 µM histamine was added to cells previously exposed to dimaprit (tracing B) and vice versa (tracing A). The lack of a stimulatory effect of dimaprit and its inhibition of histamine was verified in 4 experiments (see text for details). D: effect of the sequential addition of 1 mM dimaprit followed by 100 µM histamine was assessed in the presence (tracing A) and absence (tracing B) of 1 mM cimetidine. The lack of an effect of cimetidine on the subsequent inhibition of histamine-stimulated Isc increase by dimaprit was verified in 3 experiments. A smaller scale is shown on the ordinate, corresponding to the smaller Isc observed in this particular series of experiment.

The action of cimetidine, the H2 antagonist, was also evaluated. Cimetidine had no effect on the Isc when added by itself; it also did not inhibit the subsequent Isc increase stimulated by histamine (Fig. 5B). In some experiments, after the addition of cimetidine, the Isc response was slightly sharper (higher amplitude and shorter duration), but the overall areas under the curve were comparable [area with cimetidine: 96 ± 15% (n = 3) of control response without cimetidine]. The inhibition of histamine effect in an Ussing chamber by an H1, but not H2, antagonist is consistent with the findings observed in previous efflux studies; it suggests again that the stimulatory effect of histamine is mediated through H1 receptors.

The effect of dimaprit, the H2 agonist, and its interaction with histamine are shown in Fig. 5C. Dimaprit did not stimulate an Isc increase when added in isolation or when added after histamine. On the other hand, dimaprit appears to partially inhibit the subsequent response to histamine. Indeed, the peak Isc stimulated by 100 µM histamine was inhibited from a control value of 26.9 ± 3.1 µA/cm2 to a value of 10.4 ± 3.1 µA/cm2 after addition of 1 mM dimaprit (n = 4, P = 0.009 by 2-tailed unpaired t-test, 6 degrees of freedom).

To examine whether the inhibitory effect of dimaprit was mediated through H2 receptors, the effect of 1 mM cimetidine on the ability of dimaprit to subsequently inhibit histamine was evaluated. When 1 mM cimetidine, 1 mM dimaprit, and 100 µM histamine were added in sequence, the final response to histamine was the same in the presence or absence of cimetidine (Fig. 5D). Because the inhibitory effect of dimaprit was not blocked by prior addition of the H2 antagonist cimetidine, it probably was not mediated through an H2 receptor.

    DISCUSSION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Exocrine pancreatic secretion is composed of the secretion of pancreatic enzymes, mediated by acinar cells, and the secretion of fluid and electrolytes, mediated by PDEC. Histamine has been observed to affect this secretion. However, depending on the system and the species used, histamine may have different effects, mediated through different mechanisms. Injected arterially into anesthetized dogs, histamine stimulated pancreatic secretion as evidenced by an increased volume, bicarbonate concentration, and protein content of pancreatic juice obtained from main pancreatic duct cannulation. The agonist and antagonist profile of this effect suggested mediation through H2 receptors (1, 5). When rabbits were studied instead of dogs, a more complex effect was produced by histamine: through H1 receptors, histamine stimulated pancreatic secretion, but through H2 receptors, it inhibited secretion (16, 17). When whole pancreata or pancreatic lobules from rabbits were studied, histamine stimulated an increased production in both the volume and enzymatic content of pancreatic secretion through H1 receptors (10). In guinea pigs, histamine also stimulated enzyme production, K+ release, and [Ca2+]i increase in whole pancreata and pancreatic segments (18, 19). More recently, histamine has been shown to inhibit pancreatic secretion in guinea pigs, acting through H3 receptors on intrinsic pancreatic nerves to decrease acetylcholine release (6).

In these studies of relatively intact tissue, it is difficult to determine which cell type is directly affected by histamine and which receptor subtype mediates that specific action. The increased pancreatic secretion from cannulated ducts can reflect stimulation of both acinar and ductular components. In addition, regulatory interactions between different cell types can also occur; as discussed above, histamine may decrease pancreatic secretion indirectly through the inhibition of acetylcholine release (6). This report is, to our knowledge, the first study examining the direct action of histamine on PDEC.

Through studies of 125I- and 86Rb+ effluxes, we demonstrated that histamine activated both Cl- and K+ channels on dog PDEC. The uses of 125I- and 86Rb+ effluxes to study Cl- and K+ channels have been validated (21) and led to the successful characterization of Cl- and K+ conductances in PDEC (Refs. 14 and 14a).

We previously demonstrated that dog PDEC express two distinct Cl- channels: a cAMP-activated channel corresponding to CFTR, inhibited by NPPB and DPC but resistant to DIDS, and a Ca2+-activated Cl- channel inhibited by NPPB, DPC, and DIDS (14). Because the histamine-stimulated 125I- efflux is inhibited by NPPB, DPC, and DIDS, it most likely is mediated through Ca2+-activated Cl- channels.

The partial inhibition by DIDS merits further discussion. This partial inhibition was previously observed with the Ca2+-activated Cl- channel in PDEC (14) and is consistent with a role for this channel in mediating histamine-stimulated 125I- efflux; however, it does not exclude a partial role for the DIDS-resistant CFTR Cl- channel in this efflux. On the other hand, the abolition of this efflux by the Ca2+ chelator BAPTA suggests dependence on increased [Ca2+]i and thus a principal role for the Ca2+-activated Cl- channel. In control experiments, BAPTA has no effect on cAMP-activated I- efflux mediated by CFTR and stimulated with forskolin (data not shown).

In PDEC, the function of Cl- channels may be coupled to the Cl--bicarbonate exchanger; a potential effect of DIDS on this exchanger may indirectly affect 125I- flow through Cl- channels. However, such an indirect effect should affect both the CFTR and the Ca2+-activated Cl- channels; in our system, only the Ca2+-activated, but not the cAMP-activated, 125I- efflux is inhibited by DIDS (14). An indirect effect of DIDS is therefore unlikely, even though it cannot be excluded; either way, DIDS only inhibited the 125I- efflux mediated through Ca2+-activated Cl- channels.

We also recently observed the presence, on PDEC, of Ca2+-activated K+ channels sensitive to charybdotoxin (14a). The increased [Ca2+]i stimulated by histamine would be expected to activate these K+ channels. Indeed, histamine stimulated a 86Rb+ efflux from PDEC that was sensitive to the K+ channel inhibitor charybdotoxin. Activation of K+ channels is also consistent with the report that histamine evokes release of K+ from mouse and guinea pig pancreatic segments (18).

The secretory effect of histamine occurs at concentrations as low as 10 µM and appears to be mediated through specific receptors. Of the three histamine receptor subtypes described, H1 receptors are coupled to phospholipase C and increased [Ca2+]i, H2 are coupled to adenylate cyclase and increased cAMP, and H3 receptors mediate neuroendocrine regulation. The secretory effects of histamine on PDEC, dependent on increased [Ca2+]i, are most likely mediated through H1 receptors. H1 receptor involvement is further supported by the inhibitory effect of the specific H1 antagonist, diphenhydramine [no specific H1 agonist has been identified (9)]. The stimulatory role of H2 receptors was excluded by the ineffectiveness of dimaprit, an H2 agonist, and the resistance to cimetidine, an H2 antagonist. On the other hand, it remains possible that PDEC express H2 receptors that do not participate in these effects of histamine and therefore were silent when assessed for these functions. H3 receptors are only expressed in the nervous system and on endocrine cells (7, 9, 22); they would not mediate secretion in epithelial cells.

In Ussing chambers, dimaprit inhibits the subsequent secretory response to histamine, raising the possibility that H2 receptors may mediate an inhibitory action. This action would be consistent with the in vivo studies in which H2 agonists and antagonists, respectively, inhibited and stimulated pancreatic flow and enzyme output in rabbits (17). However, the inhibitory action of dimaprit was resistant to cimetidine, suggesting that it was not mediated through H2 receptors. Considering the high concentration of dimaprit used relative to histamine (1 vs. 0.1 mM), it is possible that dimaprit may cross-react with the H1 receptor as an antagonist and inhibit histamine action in this manner.

Histamine acts mainly in a paracrine or neuroendocrine mode, and the secretory effects described in this report are only relevant if histamine is present in the pancreas. The presence and distribution of histamine have been determined fluorometrically and biologically in tissues of different species. The concentrations of histamine in human and canine pancreas are, respectively, 4.1 and 11.4 µg/g fresh weight. These concentrations are higher than the concentration of 1 µg/g found in fat, mesentery, or muscle; the highest concentration, 19 µg/g, is found in the body of the stomach (12). In addition to neurons and endocrine cells, mast cells are also a major source of histamine; these cells are present in guinea pig pancreas (20) and in both normal and inflamed human pancreas (T. D. Nguyen and M. P. Bronner, unpublished data). Histamine may also play a role in the inflammatory response of pancreatitis; elevated levels of blood and pancreatic histamine were observed in a rat model of pancreatic inflammation from chronic dietary magnesium deficiency (2). Histamine is therefore available for interaction with pancreatic duct cells.

In summary, we have shown that histamine interacts with basolaterally located H1 receptors on PDEC to increase [Ca2+]i and activate Cl- and K+ channels. This effect may be relevant to the regulation of pancreatic ductal secretion and the pathological manifestations of pancreatitis.

    ACKNOWLEDGEMENTS

We thank Dr. Sum Lee (Division of Gastroenterology, University of Washington) for advice on the culture and characterization of pancreatic duct epithelial cells.

    FOOTNOTES

This research was supported in part by funds from the Cystic Fibrosis Foundation and the Department of Veterans Affairs.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Address for reprint requests: T. D. Nguyen, GI Section (111 GI), VA Medical Center, 1660 S. Columbian Way, Seattle, WA 98108.

Received 5 January 1998; accepted in final form 19 March 1998.

    REFERENCES
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

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Am J Physiol Gastroint Liver Physiol 275(1):G76-G84
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