Domain-specific purinergic signaling in polarized rat cholangiocytes

Kelli D. Salter, J. Gregory Fitz, and Richard M. Roman

Department of Medicine, University of Colorado Health Sciences Center, Denver, Colorado 80262


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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) (ATPgamma S) 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


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 Omega  · 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.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 Omega  · 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 Omega  · cm2 were used for experimentation (range 2,084-3,702 Omega  · cm2; mean 2,817 ± 45 Omega  · 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.

An Ussing chamber specifically designed to house 4.52-cm2 Costar Transwells with solution reservoirs of ~20 ml was used for electrophysiological analyses. Each half-chamber was filled with standard NaCl-rich extracellular solution (pH 7.4; osmolality ~295 mosmol/kg) containing (in mM) 140 NaCl, 4 KCl, 1 KH2PO4, 2 MgCl2, 1 CaCl2, 10 glucose, and 10 HEPES-free acid. Solutions were preequilibrated with 100% O2 and maintained at 37°C before use in the chamber. On mounting of the cells, the apical and basolateral compartments were bathed in preequilibrated standard extracellular solution that was independently and continuously bubbled with 100% O2 and maintained at 37°C using a recirculating heated water bath and a standard mercury thermometer to intermittently determine the temperature.

Two agarose-KCl electrodes (3% agar-1 M KCl) connected each half of the chamber to a current clamp amplifier (model EC-825; Warner Instruments, Hamden, CT). Open-circuit transepithelial voltage (Vt), transepithelial conductance (Gt, mS/cm2) and Isc (µA/cm2) were determined immediately after mounting. Isc was measured when Vt was clamped to 0 mV. The current deflection (I) response to a 10-mV offset lasting 3 s was obtained every 90 s. Gt and its inverse (Rt) were then calculated using Ohm's law (V = IR) and the relationship Gt = I/Vt. The clamp was zeroed before each experiment by inserting a blank Transwell as described above and placing preequilibrated standard extracellular buffer into each half-chamber. A 10-min equilibration period was allowed for stabilization of Isc and Rt after cells were mounted into the chamber. Individual experiments lasted ~30 min after addition of test substrates. Reagents were added to either the apical or basolateral solution of each half-chamber as indicated.

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 · wash-1 · compartment-1). Inserts were placed in the middle of a 35-mm petri dish containing 200 µl of OPTI-MEM I reduced serum medium; pegs elevated the Transwell ~1 mm above the surface of the petri dish and allowed for adequate bathing of the basolateral membrane. Subsequently, 200 µl of OPTI-MEM I reduced serum medium were added to the apical compartment. Luciferase-luciferin reagent (50 µl/ml) was added to one compartment only so that apical or basolateral ATP could be specifically detected. The dish containing the insert was lowered directly into the luminometer chamber in complete darkness and allowed to stabilize for 5 min to allow for dissipation of ATP release resulting from mechanical perturbation caused by washing. Subsequently, bioluminescence, in arbitrary light units (ALU), was recorded continuously with 15-s photon collection intervals for an additional 5 min.

The effects of test reagents on ATP release were assessed by addition of 20 µl of stock solutions to achieve final concentrations of 500 µM, 8-(4-chlorophenylthio)-adenosine 3',5'-cyclic monophosphate (CPT-cAMP), 1 mM 8-bromoguanosine 3',5'-cyclic monophosphate (8-BrcGMP), 2 µM thapsigargin, and 2 µM ionomycin. In other studies, cell volume increases or decreases were induced by adding water to decrease solution osmolality 20% or saline solution to increase final NaCl concentration by 20%. In all cases, the test solution contained supplemented luciferin-luciferase to ensure a constant level of enzyme. Inhibition of calcium-, cAMP-, and protein kinase C (PKC)-dependent signaling was performed by preincubating (37°C, 95%-O2-5% CO2) cells in medium supplemented with 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM (100 µM), Rp diastereomer of adenosine 3',5'-cyclic monophosphothioate (Rp-cAMPS; 1 mM) or calphostin C (500 nM), respectively. Identical volumes of isotonic media (OPTI-MEM I reduced serum medium for cAMP, cGMP, cell volume, and Rp-cAMPS studies) or vehicle (OPTI-MEM I reduced serum medium + 0.001% DMSO for thapsigargin, ionomycin, BAPTA-AM, and calphostin C studies) were added in control studies performed in parallel to dissociate reagent-induced and volume-induced ATP release from mechanosensitive ATP release. All solutions added contained luciferase-luciferin (50 µl/ml of stock concentrate) to avoid dilution of the enzyme. Membrane-specific ATP degradative pathways were assessed by the addition of a known quantity of ATP (1 µM) to either the apical or the basolateral compartment. Bioluminescence was recorded continuously every 15 s for 60 min and then every 5 min thereafter until a value of zero was achieved.

Reagents. 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) (ATPgamma S; tetralithium salt), ADP (sodium salt), AMP (sodium salt), adenosine (free base), 2-methylthioadenosine 5'-triphosphate (2-MeS-ATP; tetrasodium salt), beta ,gamma -methyleneadenosine 5'-triphosphate (AMP-PCP; sodium salt), alpha ,beta -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 ATPgamma S 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.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 ATPgamma S (0.01 µM-1 µM), indicating that receptor activation does not require ATP hydrolysis.



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Fig. 1.   Dose-response relationship for apical and basolateral P2 receptors. Transepithelial currents (change in short-circuit current; Delta Isc) in response to purinergic stimulation were measured in normal rat cholangiocyte (NRC) monolayers mounted in an Ussing chamber (see METHODS). Compared with apical membranes (A), basolateral membranes (B) were more sensitive to selective addition of ATP and adenosine 5'-O-thiotriphosphate (ATPgamma S). Values are means ± SE; n = 3-5 monolayers/time point.

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 ATPgamma S were equally efficient at stimulating transepithelial currents, with half-maximal increases (Delta ) 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 ATPgamma S (~300 nM) required to produce a half-maximal increase in Isc was lower than that previously reported (~2-3 µM).

The pharmacological properties of the basolateral response were different, with an agonist selectivity of ADP >=  2-MeS-ATP >=  ATP > ATPgamma >=  UTP >> AMP. Adenosine, AMP-PCP, and AMP-CPP were without notable effect (Fig. 2). The maximal current response of the basolateral membrane was obtained at ~500 nM for both ATP and ATPgamma S, with half-maximal increases in Delta Isc at 50 and 100 nM, respectively (Fig. 1). These differences in agonist selectivity are not consistent with any cloned P2 receptor subtype. Moreover, the substantial response to both ADP and 2-MeS-ATP cannot be attributed to activation of P2Y2 receptors (4). Together, however, they indicate that the basolateral membrane of NRC is distinctly different from the apical membrane in regard to both purinergic receptor expression and sensitivity to exogenous purines.


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Fig. 2.   Pharmacological properties of basolateral membrane sensitivity to nucleotides. Peak Delta Isc during selective addition of reagents to basolateral membranes are shown. Peak currents occurred at a nucleotide concentration of ~300 nM. Values are means ± SE; n = 3-5 monolayers/category. AMP-PCP, beta ,gamma -methyleneadenosine 5'-triphosphate; AMP-CPP, alpha ,beta -methyleneadenosine 5'-triphosphate; 2-MeS-ATP, 2-methylthioadenosine 5'-triphosphate.

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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Fig. 3.   Bioluminescence ATP detection assay. Extracellular medium ATP was detected using the luciferase-luciferin assay (see METHODS). A: standard curve for ATP concentration ([ATP]) in absence of tissue culture insert (20 µl ATP stock in 200 µl OPTI-MEM I reduced serum medium containing 50 µl/ml luciferase-luciferin reagent). Values are means ± SE; n = 3 separate analyses per [ATP]. ALU, arbitrary light units. B: efficiency of detection of ATP in presence and absence of tissue culture insert. Compared with addition of ATP to medium on culture dishes (filled bars), luminescence was significantly decreased when similar ATP concentrations were added to medium in apical and basolateral chambers of culture inserts (open bars). Concentrations of 200 and 50 nM ATP were used for apical and basolateral studies, respectively (note fewer ALU for latter). Values are means ± SE; n = 3 inserts/compartment.

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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Table 1.   Regulation of ATP efflux



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Fig. 4.   Basolateral ATP permeability is sensitive to increase in intracellular Ca2+. A representative tracing of non-receptor-mediated basolateral ATP efflux is shown. Changes in ATP release are represented by ALU (y-axis). Compared with control, exposure to thapsigargin (2 µM, arrow) to mobilize intracellular Ca2+ induced a rapid and sustained increase in basolateral membrane ATP release.

Addition of CPT-cAMP (500 µM), 8-BrcGMP (1 mM), ionomycin (2 µM), or thapsigargin (2 µM) to the apical compartment did not induce absolute increases in bioluminescence, as recorded by the luminometer. However, a slight but consistent increase above control values was reflected when the rate of change in extracellular ATP concentrations was determined. In contrast, ionomycin, cGMP, and thapsigargin each increased bioluminescence (1.75-, 1.14-, and 1.13-fold, respectively) when added to the basolateral compartment. Basolateral addition of CPT-cAMP also caused a slight but significant increase in bioluminescence. Moreover, as shown in Table 1, the relative ability of each stimulus to enhance apical and basolateral ATP permeability differed: ionomycin = 8-BrcGMP > thapsigargin > CPT-cAMP for apical release and ionomycin > thapsigargin > 8-BrcGMP > CPT-cAMP for basolateral release. Thus the signals responsible for regulation of ATP release from apical and basolateral cholangiocyte membranes are distinct.

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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Table 2.   Inhibition of cell swelling-induced ATP efflux




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Fig. 5.   Membrane specificity of volume-sensitive ATP efflux. Representative tracings that demonstrate characteristic changes in bioluminescence in apical and basolateral compartments during changes in cell volume are shown. A: addition of water (hypotonicity) or NaCl (hypertonicity) to change osmolarity ~20% (arrow) rapidly increased and decreased apical ATP release, respectively, compared with control. B: in a similar fashion, medium dilution (hypotonicity; arrow) led to a large increase in basolateral luminescence compared with control. Effect of NaCl addition (hypertonicity) on ATP permeability was less pronounced.

Both [Ca2+]i and PKC have been shown to modulate liver cell response to cell volume increases (20, 23). Because [Ca2+]i also appears to modulate ATP release, the role of multiple intracellular signaling pathways in ATP release was assessed through the use of specific inhibitors. Results are shown in Table 2. Cell swelling-induced ATP efflux was significantly inhibited (~80% apical; ~68% basolateral) by chelation of intracellular Ca2+ (BAPTA-AM; 100 µM). Similarly, partial decreases in bioluminescence (~25% for both apical and basolateral) occurred when PKC activity was inhibited (calphostin C; 500 nM). In contrast, volume-sensitive ATP efflux in the apical compartment was slightly but significantly increased through inhibition of cAMP signal transduction (1 mM Rp-cAMPS). Neither increase nor inhibition of cell swelling-induced ATP efflux occurred in the basolateral compartment on exposure to Rp-cAMPS. Thus Ca2+-dependent signaling mechanisms appear to be the primary pathways contributing to regulation of both constitutive and volume-sensitive ATP release.

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 = ae-0.058 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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Fig. 6.   Kinetics of ATP degradation differ between apical (A and C) and basolateral (B and D) cholangiocyte membranes. ATP degradation was assessed after addition of ATP (1 µM) as described in METHODS. A and B: rate of decline in bioluminescence differed when ATP was added (arrow) to apical (continuous) compared with basolateral (biphasic) compartments of NRC monolayers. C and D: regression analysis revealed distinct kinetics of exogenous ATP degradation in apical and basolateral compartments. Values are means ± SE; n = 3 monolayers/time point.


    DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 approx  UTP approx  ATPgamma 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.


    FOOTNOTES

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.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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