Department of Medicine, University of Colorado Health Sciences Center, Denver, Colorado 80262
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ABSTRACT |
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In
cholangiocytes, adenine nucleotides function as autocrine/paracrine
signals that modulate ductular ion transport by activation of
purinergic receptors. The purpose of these studies was to identify cellular signals that modulate ATP release and nucleotide processing in
polarized normal rat cholangiocytes. In Ussing chamber studies, selective exposure of the apical and basolateral membranes to ATP or
adenosine 5'-O-(3-thiotriphosphate) (ATPS)
stimulated increases in short-circuit current. Apical purinergic
receptor agonist preference was consistent with the P2Y2
subtype. In contrast, basolateral ADP was more potent in stimulating
transepithelial currents, consistent with the expression of different
basolateral P2 receptor(s). Luminometric analysis revealed that both
membranes exhibited constitutive ATP efflux. Hypotonic exposure
enhanced ATP release in both compartments, whereas decreases in ATP
efflux during hypertonicity were more prominent at the apical membrane. Increases in intracellular cAMP, cGMP, and Ca2+ also
increased ATP permeability, but selective effects on apical and
basolateral ATP release differed. Finally, the kinetics of ATP
degradation in apical and basolateral compartments were distinct. These
findings suggest that there are domain-specific signaling pathways that
contribute to purinergic responses in polarized cholangiocytes.
P2 receptors; extracellular nucleotides; adenosine 5'-triphosphate release and processing; transepithelial ion transport
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INTRODUCTION |
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FLUID ABSORPTION AND SECRETION across intrahepatic bile
ducts play a key role in modifying the volume and composition of bile. Bile formation by the liver results from the combined complementary interactions and functions of two distinct liver cell types. Secretion is initiated by hepatic parenchymal cells (~80% of liver mass) that
actively transport bile salts and other organic solutes into the
canalicular space between cells (8, 18). Subsequently, canalicular bile
flows into the lumen of an extensive network of intrahepatic ducts,
where it undergoes dilution and alkalization as a result of
cholangiocyte Cl and
HCO
3 secretion (17). Despite the
comparatively small number (3-5% of liver mass) of biliary
epithelial cells lining the intrahepatic bile ducts (28), cholangiocyte
secretion is thought to account for up to 40% of human bile flow (18). In humans, for example, the intrahepatic ductular network is estimated to be 1-2 km in length (15) and hormonal stimulation of ductular bile formation by secretin increases
HCO
3 output and bile flow from 0.67 to
1.54 ml/min with no effect on bile salt transport (14). At the cellular
level, the response to secretin is thought to involve binding to
basolateral receptors (6), stimulation of exocytosis via a
cAMP-dependent mechanism (13), and efflux of apical
Cl
and HCO
3 through
opening of cystic fibrosis transmembrane regulator-associated
Cl
channels and enhanced
Cl
/HCO
3 exchange
(1, 3, 16, 17). Little is known regarding alternative cAMP-independent
pathways that modulate ductular secretion. However, definition of such
pathways might provide attractive strategies for pharmacological
treatment of liver disease in cystic fibrosis, for enhancing bile flow
in cholestasis, and for increasing the solubility of cholesterol in
patients with gallstone formation.
ATP and its metabolites are found in bile and interstitial fluid in
liver (2). Recent studies suggest that these nucleotides might serve as
potent autocrine/paracrine signaling molecules affecting cholangiocyte
transport by binding to purinergic receptors in the plasma membrane
(15, 21, 22, 25, 30, 34). The cellular origin of ATP and the factors
that modulate its local concentrations have not been defined. In
isolated bile duct units, stimulation of purinergic receptors in the
basolateral membrane increases cytosolic Ca2+ concentration
([Ca2+]) (19). Stimulation of apical purinergic
receptors in an in vitro model of cholangiocytes in culture causes
rapid and substantial increases in the Cl
permeability of the apical membrane, favoring efflux of
Cl
from the cell into the lumen (25). Because
cholangiocytes are a polarized epithelium with high-resistance tight
junctions (>1,000
· cm2) between
cells, there is strict localization of transport proteins to specific
domains. Thus the apical and basolateral membranes, through effects on
in vivo ATP release and degradation, may distinctively modulate
cholangiocyte function.
Previous studies of cholangiocyte ATP permeability and metabolism have been limited in part by the lack of polarized model systems and by the resolution of ATP assay systems that might not permit detection of ATP in the nanomolar concentrations sufficient to initiate purinergic signaling. ATP and/ or UTP release has been detected in other epithelial and nonepithelial cell types (4). A broad range of stimuli, including mechanical stress (9, 10, 26), cytosolic [Ca2+] (22), cytosolic cAMP concentration (27), phosphatidylinositol 3-kinase (7), and changes in cell volume (22, 24, 32), appear to modulate release according to the cell under investigation. Consequently, the purpose of these studies was to utilize a polarized model of cholangiocytes in culture that permits selective access to the apical and basolateral domains to evaluate the cellular signals responsible for cholangiocyte ATP release and degradation. The observation that there are significant differences between the apical and basolateral compartments in regard to purinergic receptor expression, constitutive ATP efflux, non-receptor-mediated nucleotide release, and pathways of exogenous ATP degradation suggests that local nucleotide concentrations are tightly regulated in a domain-specific manner.
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METHODS |
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Cells and method of culture. Polarized normal rat cholangiocytes (NRC) in long-term culture (passages 9-25) were propagated in vented Falcon tissue culture flasks (75 cm2) on a thick layer (1-2 mm) of rat tail collagen as substratum as previously described (31) with the following modifications. Cells were maintained in antibiotic-free DMEM nutrient mixture F-12 (DMEM-F-12) supplemented with the following reagents: MEM nonessential amino acid solution (0.01 ml/ml), chemically defined lipid concentrate (0.01 ml/ml), MEM vitamin solution (0.01 ml/ml), L-glutamine (2 mM), soybean trypsin inhibitor (0.05 mg/ml), insulin-transferrin-selenium-S supplement (0.01 ml/ml), and fetal bovine serum (5%). Immediately before media replacement, an aliquot of base medium was additionally supplemented with bovine pituitary extract (30 µg/ml), dexamethasone (393 ng/ml), epidermal growth factor (25 ng/ml), forskolin (4.11 µg/ml), and 3,3',5-triiodo-L-thyronine (3.4 µg/ml). Cells were passaged at ~95% confluence, as assessed by microscopic inspection. The collagen slab was dislodged and then digested using a filter-sterilized solution of dispase (2 U/ml) and collagenase (10 mg/ml) in nonsupplemented DMEM-F-12. Cells were washed twice with PBS solution, allowed to pellet by natural gravitation between washes, and resuspended in supplemented DMEM-F-12. Cells were seeded into either T75 flasks containing a sterile, prewashed (with PBS) collagen slab (~1-2 mm thickness) and supplemented base medium (12 ml/T75 flask) or collagen-coated Transwells, as described in Electrophysiology.
For Ussing chamber analysis, NRC were seeded onto 4.52-cm2 collagen-coated semipermeable (0.4-µm pore size) supports (Costar, Cambridge, MA) at a plating density of 0.5 × 106 cells/well. Cells were grown to confluence in serum-free, antibiotic-free DMEM-F-12 supplemented with the following reagents: MEM nonessential amino acid solution (0.01 ml/ml), chemically defined lipid concentrate (0.01 ml/ml), MEM vitamin solution (0.01 ml/ml), L-glutamine (2 mM), and soybean trypsin inhibitor (0.05 mg/ml). Selective supplementation of the apical and basolateral media with insulin-transferrin-selenium-S supplement (0.01 ml/ml), bovine pituitary extract (30 µg/ml), dexamethasone (393 ng/ml) and/or 3,3',5-triiodo-L-thyronine (3.4 µg/ml) was used to improve electrophysiological properties. Transepithelial resistance was assessed using an epithelial voltohmmeter (EVOM; World Precision Instruments, Sarasota, FL). For bioluminescence ATP detection assay, NRC were plated onto 0.79-cm2 semipermeable (0.2-µm pore size) supports (Nalge Nunc International, Naperville, IL) coated with rat tail collagen type 1 (Collaborative Biomedical Products, Bedford, MA) at 300 µg/ml. Cells were seeded at a density of 0.1 × 106 cells/well and maintained as described above for Ussing chamber analysis.Electrophysiology.
Electrophysiological studies were performed using both an EVOM to
monitor development of transepithelial resistance
(Rt) and an Ussing chamber (Jim's Instrument
Manufacturing, Iowa City, IA). Monolayers reached confluence, as
assessed by microscopic examination, 5-7 days after plating.
Transmembrane resistance increased from <600 (resistance of solution + membrane support) to >2,000 · cm2
as confluence was achieved. Rt reached plateau
values after 10-14 days in culture and was stable for ~3 wk
thereafter. Only NRC monolayers exhibiting Rt
2,000
· cm2 were used for
experimentation (range 2,084-3,702
· cm2; mean 2,817 ± 45
· cm2). Transferring monolayers to
the Ussing chamber resulted in a ~30% decrease in
Rt as measured by EVOM immediately before transfer to the chamber. Short-circuit current (Isc) either
slightly declined or was stable for 30-60 min after mounting.
Bioluminescence ATP detection assay. Cellular ATP release was detected via bioluminescence using the firefly luciferin-luciferase assay as recently described (22, 29). Luminescence was detected with a TD-20/20 luminometer (Turner Designs, Sunnyvale, CA) that houses a chamber that accommodates a platform to hold 35-mm petri dishes. The amount of light produced is proportional to the amount of ATP in the extracellular solution.
Before analysis, apical and basolateral compartments were washed twice with PBS (500 µl · washReagents.
The following reagents were purchased from GIBCO-BRL (Grand Island,
NY): DMEM-F-12, MEM nonessential amino acid solution, chemically
defined lipid concentrate, MEM vitamin solution,
L-glutamine, soybean trypsin inhibitor,
insulin-transferrin-selenium-S supplement solution, dispase, PBS, and
OPTI-MEM I reduced serum medium. Dexamethasone, 3,3',5-triiodo-L-thyronine, forskolin, collagenase,
CPT-cAMP (sodium salt), ATP (disodium salt), adenosine
5'-O-(3-thiotriphosphate) (ATPS; tetralithium salt),
ADP (sodium salt), AMP (sodium salt), adenosine (free base),
2-methylthioadenosine 5'-triphosphate (2-MeS-ATP; tetrasodium
salt),
,
-methyleneadenosine 5'-triphosphate (AMP-PCP; sodium salt),
,
-methyleneadenosine 5'-triphosphate
(AMP-CPP; lithium salt), and UTP (sodium salt) were purchased from
Sigma Chemical (St. Louis, MO). Thapsigargin, ionomycin (free acid), 8-Br-cGMP (sodium salt), BAPTA-AM, Rp-cAMPS, calphostin C, and luciferin-luciferase reagent (ATP assay kit) were purchased from Calbiochem-Novabiochem (La Jolla, CA). Defined fetal bovine serum was
purchased from Hyclone Laboratories (Logan, UT). The remaining reagents, bovine pituitary extract and epidermal growth factor, were
purchased from Upstate Biotechnology (Lake Placid, NY).
Statistics.
For Ussing chamber analyses, relative dose-response relationships for
ATP and ATPS were determined by calculating the difference between
Isc immediately before addition of the reagent and
the maximal current response induced after addition of the reagent. Data were plotted using SigmaPlot for Windows. For bioluminescence ATP
detection assays, data were compiled into Microsoft Excel spreadsheets
in which the mean ± SE was calculated for each time point in each set
of experimental time courses. Rate of change in ATP efflux was
determined by calculating the change in the slope of the line from one
time point (baseline) to another for each condition (control vs.
reagent). Values were normalized by designating the baseline value as
zero and then dividing the remaining time points by the absolute
baseline value for the change in the slope of the line.
Statistical analyses were performed using paired student's
t-test (SigmaStat) and were based on the results of relative
change in ALU within the 2-min time frame. A P value of <0.05
was considered significant.
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RESULTS |
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Domain-specific nucleotide-induced changes in Isc.
To determine whether cells in culture retained biological
responsiveness to exogenous nucleotides, we mounted confluent NRC monolayers in an Ussing chamber and measured Isc
under basal conditions and after exposure to different nucleotides.
Purinergic receptor responses were identified on both the apical and
basolateral membranes of NRC monolayers. Exposure to ATP (0.01 µM-1 µM) stimulated dose-dependent increases in
transepithelial ion transport, as indicated by an increase in
Isc (Fig. 1). Changes
in Isc occurred rapidly (within ~10 s) and
reached maximal values within 60-90 s after addition of reagent.
Despite the continued presence of ATP, Isc tended to return toward basal values within 5-60 min. Similar results were observed with the nonhydrolyzable analog ATPS (0.01 µM-1 µM), indicating that receptor activation does not require ATP hydrolysis.
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Pharmacological properties of apical vs. basolateral electrogenic
responses.
The pharmacological profile of the apical response to nucleotides was
consistent with expression of purinergic receptors of the P2U
(P2Y2) receptor subtype (4, 22, 25). ATP, UTP, and ATPS
were equally efficient at stimulating transepithelial currents, with
half-maximal increases (
) in Isc, at ~300 nM. ADP, AMP, and adenosine were without effect. These results are consistent with previous results in NRC monolayers (25), with the
exception that the concentration of ATP
S (~300 nM) required to
produce a half-maximal increase in Isc was lower
than that previously reported (~2-3 µM).
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Polarized cholangiocytes exhibit vectorial constitutive ATP release. Monolayers were continuously monitored in the luminometer with the luciferin-luciferase reagent added selectively to the apical or basolateral compartment to determine whether NRC are capable of constitutive ATP efflux. Basal ATP release was detected in all monolayers studied. Values of 77.00 ± 5.21 ALU (n = 45) measured in the apical compartment were consistently approximately fivefold greater than the values of 15.16 ± 2.89 ALU (n = 45) measured in the basolateral compartment.
To determine whether these differences reflect a property of polarized cells or differences in apical/basolateral photon detection, additional studies were performed under cell-free conditions with exogenous ATP. The efficiency of bioluminescence detection, as determined through the development of an ATP standard curve, was decreased in the presence of a blank (unseeded) Transwell insert (Fig. 3). The apparent decrease in efficiency was greater in the apical (48% of control) compared with the basolateral (67% of control) compartment. This likely reflects light scattering induced by the distance between the lid of the petri dish and the Transwell insert. Thus the ALU values reported are likely to underestimate the actual concentration of ATP present. Technical differences in measurement do not account for the exaggerated apical ATP release. On the basis of standard curve analysis, a mean value of 77.00 ALU in the apical compartment would correspond to ~250 nM ATP, whereas a mean value of 15.16 ALU in the basolateral compartment would correspond to ~50 nM ATP.
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Regulation of vectorial ATP release by cellular signaling pathways.
Multiple stimuli have been reported to modulate cellular ATP release in
different model systems (4, 22). Consequently, studies were performed
to assess potential regulatory effects of cAMP, cGMP, and
Ca2+-dependent signaling in NRC. The results are summarized
in Table 1, with a representative tracing
illustrated in Fig. 4. In all studies,
there was a consistent gradual decrease in ALUs with time.
Consequently, same-day controls were performed in parallel and results
of test reagents were compared with control values. In control studies,
addition of OPTI-MEM I reduced serum medium (vehicle control for
water-soluble reagents) or OPTI-MEM I reduced serum medium with 0.001%
(final concentration) DMSO was without effect, thereby eliminating
significant effects on ATP release caused by mechanical forces or
chemical solvents.
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Effects of cell volume changes on vectorial ATP release.
Epithelial cells undergo regulated changes in cell volume in response
to multiple physiological stimuli. Because cell volume has been
reported to modulate ATP release, the effects of cell volume increases
(hypotonic exposure) and decreases (hypertonic exposure) on
bioluminescence were assessed. Results are shown in Table
2, with representative tracings illustrated
in Fig. 5. Hypo- and hypertonic exposure
had contrasting effects on ATP release. In the apical compartment,
increases in cell volume (hypotonic exposure; ~240 mosmol/kg)
increased bioluminescence approximately twofold, whereas decreases in
cell volume (hypertonic exposure; ~360 mosmol/kg) decreased
bioluminescence approximately twofold. In the basolateral compartment,
increases in cell volume also increased bioluminescence to a degree
approximately fourfold greater than same-day controls. However,
exposure of the basolateral membrane to hypertonic solution caused a
slight but significant decrease in ALU. Thus cell volume represents an
important determinant of vectorial ATP release.
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Domain-specific ATP degradation.
Extracellular ATP concentrations are regulated not only through the
rate of ATP efflux but also through multiple degradation pathways (4,
22). To further elucidate the dynamic relationship between vectorial
release and degradation of ATP, the medium bathing NRC monolayers was
selectively (apical vs. basolateral) loaded with exogenous ATP (1 µM). Changes in bioluminescence were monitored continuously until
relative ALU reached zero. Composite tracings (n = 3 per
compartment) of membrane-specific changes in extracellular ATP
concentrations after addition of 1 µM ATP are illustrated in Fig.
6. Addition of 1 µM ATP to the apical
compartment increased bioluminescence ~40-fold higher than basal
values before administration (42.31 ± 2.65 vs. 1,752.00 ± 29.05 ALU). The time course of degradation (Fig. 6C) was described by
a single exponential (y = ae0.058
min; r = 0.99). Approximately 13% of
extracellular ATP degraded within the first minute. By comparison, in
the basolateral compartment, the same concentration of
exogenous ATP increased bioluminescence ~70-fold higher than basal
values (16.88 ± 0.18 vs. 1,224.67 ± 10.02 ALU). The time
course of degradation was more complex, with a rapid initial clearance
of 30% of exogenous ATP within 1 min, followed by a more gradual
decline. The rate of decrease (Fig. 6D) was well described by a
double exponential (y = ae
0.037 min + ce
0.001 min; r = 0.99).
Therefore, it appears that a single degradation pathway exists in the
apical compartment. In contrast, ATP degradation in the basolateral
compartment shows biphasic or multiple degradation pathways, with rapid
initial clearance followed by a more gradual decline.
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DISCUSSION |
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Purinergic signaling is postulated to regulate cholangiocyte transport by binding of extracellular nucleotides to purinergic receptors in the plasma membrane. Although the cellular origin of purinergic agonists has not been clearly defined, the recent observations that hepatocytes, cholangiocytes, and other epithelia are capable of regulated ATP release under different conditions suggest that modulation of nucleotide release is tissue specific. The principal findings of these studies are 1) purinergic receptor expression in cholangiocytes is membrane specific; 2) cholangiocytes exhibit high levels of constitutive vectorial ATP release; 3) multiple intra- and extracellular signals modulate the rate of vectorial ATP release; and 4) regulation of local ATP concentrations is domain specific. Substantial differences in these parameters were noted between the apical and basolateral compartments of polarized rat cholangiocytes. Together, these findings provide evidence for an intricate network of cellular mechanisms controlling local nucleotide concentrations and purinergic signaling in biliary epithelial cells.
Previous studies indicate that cholangiocyte transport is sensitive to
small changes in local nucleotide concentrations (22). Exposure of
different hepatobiliary cell models to ATP or UTP in nanomolar
concentrations results in opening of K+ and
Cl channels (11, 25) and stimulation of
Na+/H+ exchange (5, 34). In polarized cells in
monolayer culture, the response is mediated by stimulation of
P2Y2 receptors in the apical membrane, with subsequent
increases in the transepithelial transport of Cl
(25). These findings suggest that extracellular nucleotides might
represent a mechanism for local autocrine/paracrine regulation of
cholangiocyte transport and bile formation.
The small size and intrahepatic location of cholangiocytes in vivo limit the ability to perform direct physiological studies. However, culture of NRC under strict and defined conditions leads to the formation of polarized monolayers that exhibit well-defined apical and basolateral domains (31), thus allowing for adequate in vitro assessment of transepithelial processes. These cells and other nonpolarized biliary epithelial cell models were shown previously to express P2Y2 receptors (25, 33) and to respond to exogenous ATP with an increase in Isc (25). In addition, nonpolarized heptobiliary cell models were shown to release ATP in response to increases in cell volume (7, 24, 32). Moreover, cholangiocytes express the cystic fibrosis transmembrane conductance regulator (cftr) and P-glycoprotein products of multidrug resistance (mdr) genes, which are ATP-binding cassette proteins and have been implicated in the regulation of ATP release in other cell models (22). Thus nucleotide processing by cultured NRC is likely to be representative of the in vivo parent cell functions. However, it is acknowledged that these results must be confirmed and further delineated if improved in vitro cell models become available.
Under basal conditions, NRC showed constitutive release of ATP into the apical and basolateral chambers. ALU values measured in the apical chamber were consistently approximately fivefold greater than those in the basolateral chamber. This difference could not be attributed to technical considerations because the presence of the culture insert in the luminometer decreased the apparent ALU response to a greater degree in the apical chamber. It is important to note that determination of "bulk" ATP concentrations in extracellular media using luminometry may not accurately represent the local availability of ATP at the membrane surface, where there are rapid changes in nucleotide release and degradation and nucleoside salvage. These technical considerations suggest that the measurements of ALU reported here are likely to underestimate total ATP concentrations and to minimize differences in the apical vs. basolateral chambers.
The large differences in apical vs. basolateral nucleotide constitutive
release were matched by other significant differences in responsiveness
to exogenous nucleotides, intracellular signals evoking ATP release,
and processes of exogenous degradation. In general, apical nucleotide
processing appears to be more straightforward as follows. 1)
The high constitutive rate of release was largely unaffected by
maneuvers designed to increase intracellular cAMP, Ca2+, or
cGMP. 2) ATP release was volume sensitive,
increasing with maneuvers that increase cell volume and decreasing with
maneuvers that decrease cell volume. 3) The agonist profile of
the secretory response to nucleotides (ATP UTP
ATP
S; no
response to ADP or adenosine) was consistent with expression of a
single class of P2Y2 receptors as previously described.
4) Nucleotide degradation followed a time course readily
described by a single exponential, consistent with functional
expression of a single, dominant ATP degradation pathway.
In contrast, basolateral nucleotide processing was more complex and differentiated substantially from apical processing as follows. 1) Basal ATP release could be modulated significantly by multiple signaling pathways, with a relative potency of Ca2+ > cGMP > cAMP. 2) Maneuvers that increase cell volume substantially increased basolateral ATP release to a degree greater than that observed in the apical domain. 3) The agonist profile of the secretory response to nucleotides cannot be explained by a known class of P2Y2 receptors. Because ADP and 2-MeS-ATP each stimulate secretion, expression of other P2 receptor subtypes is postulated. 4) Nucleotide degradation shows an initial rapid time course followed by a slower time course, consistent with more than one mechanism for ATP degradation. Together, these findings indicate that there are substantial differences in the cellular mechanisms controlling local nucleotide availability in the apical vs. basolateral compartments.
Assuming that these findings are relevant to cholangiocytes in vivo, several additional points merit emphasis. First, because the apical pole of cholangiocytes lines the lumen of intrahepatic ducts, it seems likely that cholangiocytes themselves account for a portion of the ATP found in bile and that this constitutive apical release contributes to local regulation of cholangiocyte secretion. Indeed, recent studies indicate that exposure of the apical membrane to the ATPase apyrase inhibits basal and volume-sensitive secretory currents (21). Second, because ADP is a potent basolateral agonist, it is attractive to postulate that the rapid conversion of ATP to ADP seen in the basolateral domain might serve as an additional control point for regulation of secretion. These findings differ from observations in isolated duct units, in which basolateral ATP and UTP stimulate a rise in cytosolic Ca2+ but have no apparent effect on secretion (19). The reasons for these differences are not known, but they may reflect differences in the models under investigation. Alternatively, the Ussing chamber analysis used here allows for measurement of electrogenic transport under voltage-clamp conditions but does not provide an actual measurement of fluid movement.
These observations also highlight several uncertainties. First, the molecular basis of ATP release has not been established. Consequently, it is not clear whether the domain-specific responses reflect different transport proteins or different modes of regulation of the same transport protein. Interestingly, although cftr has been implicated in cAMP-dependent regulation of ATP permeability, increases in cAMP had minute effects on apical ATP release, whereas inhibition of cAMP-dependent kinase slightly but significantly increased volume-sensitive ATP release. In contrast, inhibition of cytosolic Ca2+ mobilization or PKC suppressed both constitutive and volume-sensitive ATP release. Second, because cellular ATP release has also been shown to be sensitive to changes in extracellular halide concentrations (12), alterations in media tonicity may have other effects on the ATP transporter(s) in addition to changes in cell volume. Third, the full complement of basolateral nucleotide receptors has not been defined, and the mechanisms that regulate their expression are unknown. The secretory response to ADP is particularly interesting because it is not observed in the apical domain. Finally, cholangiocytes appear to be capable of rapid nucleotide degradation, but the specific enzymes involved have not been identified.
In conclusion, multiple levels for the regulation of local ATP concentrations appear to exist in cultured polarized cholangiocytes, with notable differences between the apical and basolateral domains. These findings provide a rationale for future studies aimed at pharmacological modulation of biliary ATP concentrations in diseases affecting hepatobiliary transport. In addition, molecular characterization and localization of ATPases might provide alternative strategies for selective regulation of apical and basolateral nucleotide availability through effects on enzyme activity.
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FOOTNOTES |
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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 and other correspondence: K. D. Salter, Dept. of Medicine, Campus Box B158, Rm. 6412, Univ. of Colorado Health Sci. Ctr., Denver, CO 80262 (E-mail: kelli.salter{at}uchsc.edu).
Received 6 June 1999; accepted in final form 25 October 1999.
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