Department of Medicine, University of Washington, and Veterans Affairs Puget Sound Health Care System, Seattle, Washington 98108
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ABSTRACT |
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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
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INTRODUCTION |
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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.
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MATERIALS AND METHODS |
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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
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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.
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RESULTS |
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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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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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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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DISCUSSION |
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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.
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ACKNOWLEDGEMENTS |
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We thank Dr. Sum Lee (Division of Gastroenterology, University of Washington) for advice on the culture and characterization of pancreatic duct epithelial cells.
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FOOTNOTES |
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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.
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