Cyclic ADP-ribose, a putative Ca2+-mobilizing second messenger, operates in submucosal gland acinar cells

Kan Sasamori,1 Tsukasa Sasaki,1 Shin Takasawa,2 Tsutomu Tamada,1 Masayuki Nara,1 Toshiya Irokawa,1 Sanae Shimura,1 Kunio Shirato,3 and Toshio Hattori1

1Divisions of 1Respiratory and Infectious Diseases and 33Cardiovascular Medicine, Tohoku University Graduate School of Medicine, Sendai 980-8574; and 22Department of Biochemistry, Tohoku University Graduate School of Medicine, Sendai 980-8575, Japan

Submitted 24 December 2003 ; accepted in final form 18 February 2004


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cyclic ADP-ribose (cADPR), a putative Ca2+-mobilizing second messenger, has been reported to operate in several mammalian cells. To investigate whether cADPR is involved in electrolyte secretion from airway glands, we used a patch-clamp technique, the measurement of microsomal Ca2+ release, quantification of cellular cADPR, and RT-PCR for CD38 mRNA in human and feline tracheal glands. cADPR (>6 µM), infused into the cell via the patch pipette, caused ionic currents dependent on cellular Ca2+. Infusions of lower concentrations (2–4 µM) of cADPR or inositol 1,4,5-trisphosphate (IP3) alone were without effect on the baseline current, but a combined application of cADPR and IP3 mimicked the cellular response to low concentrations of acetylcholine (ACh). Microsomes derived from the isolated glands released Ca2+ in response to both IP3 and cADPR. cADPR released Ca2+ from microsomes desensitized to IP3 or those treated with heparin. The mRNA for CD38, an enzyme protein involved in cADPR metabolism, was detected in human tissues, including tracheal glands, and the cellular content of cADPR was increased with physiologically relevant concentrations of ACh. We conclude that cADPR, in concert with IP3, operates in airway gland acinar cells to mobilize Ca2+, resulting in Cl secretion.

patch clamp; CD38; calcium store; inositol 1,4,5-trisphosphate; calcium; acetylcholine


TRANSEPITHELIAL Cl secretion is critical for the production of properly hydrated secretion in airways, thus contributing to the airway defense mechanisms. Recurrent intractable airway infection in cystic fibrosis (CF), a hereditary disease with compromised airway Cl secretion, attests to the importance of this function since lung disease is the primary cause of mortality in CF patients (8, 64). Because human airway epithelium is likely to be primarily absorptive (5, 22, 26, 62, 63), a major fraction of the airway fluid seems to be derived from the submucosal gland (2, 48) and follows an active Cl secretion from the glandular acini (11, 14, 48). The airway secretion appears to be under tonic regulation by cholinergic nerves because oral administration of atropine reduces the secretion (35) and vagal cooling reduces it by ~40% (59). An in vivo observation indicated that cholinergic agents were much more potent stimulators of gland secretion than were adrenergic agonists when estimated using hillock formations of a powdered tantalum layer coating the airway surface (43). Also, primary cultures of human submucosal gland secreted Cl in response to both cholinergic and adrenergic stimulation, with cholinergic stimulation being more potent (65). The acinar cells of the airway submucosal gland are the predominant site for the expression of cystic fibrosis transmembrane conductance regulator (CFTR), a protein that is flawed in CF disease (2, 12, 64). Recently, CFTR has been shown to be involved in cholinergically stimulated, Ca2+-activated Cl secretion in cultured human airway cells (42) and in porcine bronchial submucosal gland (2). These findings demonstrate the major role of Ca2+-mediated Cl secretion stimulated by cholinergic agents in airway submucosal glands. However, the cellular mechanism of Ca2+ mobilization that ultimately leads to activation of Cl channels is only partly understood in airway submucosal glands.

The muscarinic receptors have been identified on swine (67), ferret (49), and feline (21) submucosal glands. Muscarinic agonists activate, via a guanine nucleotide-binding protein (G protein) coupled with the receptor, the plasma membrane phospholipase C, generating inositol 1,4,5-trisphosphate (IP3) and diacylglycerol. IP3 binds to its specific receptor on the cytosolic Ca2+-storage compartment to release Ca2+ into the cytosol, thus initiating the Ca2+ signal (4). Inositol phosphate formation in submucosal glands has been shown in response to agents, including the muscarinic agonist carbachol, in bovine airway (18). Moreover, IP3-mediated Ca2+ release has been found to play a critical role in starting acetylcholine (ACh)-evoked Cl secretion in human and feline tracheal glands in patch-clamp experiments and intracellular Ca2+ measurements (51). Interestingly, however, prevention of the IP3-mediated signaling pathway by a MAb against IP3 receptors (IP3R) did not totally abolish the ACh-induced response in tracheal glands (51). This observation raised the possibility that there may be an additional Ca2+-mobilizing mechanism independent of the IP3-mediated one in airway gland acinar cells.

A likely candidate for such a second messenger is cyclic ADP-ribose (cADPR), a metabolite of the ubiquitous pyridine nucleotide {beta}-NAD+ (7, 46, 53). Since the identification of cADPR as a potent mediator of Ca2+ release in sea urchin egg microsomes (7), its Ca2+-mobilizing activity has been shown in rodent pituitary cell (27), dorsal root ganglion cell (10), pancreatic {beta}-cell (46, 53), and successively in rodent exocrine glands, including pancreas (56), salivary (32), and lacrimal (17) glands. Moreover, the CD38 protein, originally identified on leukocytes as one of the surface antigens, was revealed to retain enzymatic activities in both the synthesis and degradation of cADPR (19, 54). CD38 appears to work in rat parotid acinar cells in raising the cellular level of cADPR (34). These findings suggest that cADPR may be involved in the Ca2+ mobilization in exocrine acinar cells. However, none of these reports (17, 32, 56) addressed whether cADPR can occur naturally in response to physiological stimuli in exocrine glands. It is also unclear whether cADPR can operate not only in rodent glands but also in human acinar cells. To address these issues, electrophysiological, biochemical, and molecular biological experiments were carried out using freshly isolated human and feline tracheal submucosal glands. This is the first report that describes cADPR involvement in Ca2+ homeostasis in human acinar cells, which may give some insights into the mechanism underlying deteriorated Ca2+-activated Cl secretion in CF submucosal glands.


    METHODS
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 ABSTRACT
 METHODS
 RESULTS
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Cell preparations. Submucosal glands were isolated either from the tracheae of human surgical specimens from patients with laryngeal cancer or those of cats (2–5 kg body wt) anesthetized with intramuscular ketamine hydrochloride (30 mg/kg) and intravenous thiopental sodium (30 mg/kg). The external surface of the cat trachea was cleaned of fat and connective tissues, cut into rings 3–4 cm long, and pinned in the extracellular solution with the posterior (membranous) wall side up. Light through a flexible fiber bronchoscope (FBS-1; Machida, Tokyo, Japan) placed inside the tracheal ring was used to transilluminate the membranous portion. In the human tracheal piece, the mucosal membrane together with the underlying connective tissues were peeled off from the cartilage layer and pinned on a silicon-rubber-coated chamber with the submucosal aspect directed upward. Light was shed from the lateral side of the tissue. The submucosal gland could then be easily distinguished from the surrounding connective tissue under a stereoscopic microscope (x60–80 magnification). Fresh, unstained submucosal glands were isolated using two pairs of tweezers and microscissors (20, 21, 51, 55). The isolated glands were further dispersed enzymatically into single or clustered acinar cells by incubating them with enzyme solution containing collagenase (200 U/ml), DL-dithiothreitol (0.31 mg/ml), and trypsin inhibitor (1 mg/ml) for 30 min at 37°C. After dispersion and being washed three times with centrifugation at 180 g, the cells were resuspended in a standard extracellular solution (see below) until use.

Electrical recordings. Ionic currents were measured with a patch-clamp amplifier (EPC9; HEKA Electronic), low-pass filtered at 2.9 kHz, and monitored on both a built-in software oscilloscope and a thermal pen recorder (RECTI-HORIZ-8K; Nippondenki San-ei, Tokyo, Japan). Patch pipettes were made of glass capillary with an outer diameter of 1.5 mm using a vertical puller (PP-83; Narishige Scientific Instruments, Tokyo, Japan) and had a tip resistance of 2–6 M{Omega}. The junction potential between the patch pipette and bath solution was nulled by the amplifier circuitry. After establishing a high resistance (>1 G{Omega}) tight seal, the whole cell configuration was obtained by rupturing the patch membrane with negative pressure applied to the pipette tip. Membrane currents were monitored at two different holding potentials (Hp), i.e., 0 and/or –80 mV, each of which roughly corresponded to the Cl and K+ equilibrium potential, respectively, under the present electrolyte conditions. The double current monitoring, i.e., alternate recording of the ionic currents corresponding to Hp of 0 and –80 mV, was accomplished by applying 200-ms voltage pulses of –80 mV at a frequency of 2 Hz to the pipette voltage of 0 mV (20, 50, 51, 55). The upward or downward deflection of the current tracing represents outward (Io) or inward current (Ii), respectively. The solutions employed were of the following compositions (in mM): extracellular (bath) solution, 120 NaCl, 4.7 KCl, 1.13 MgCl2, 1.2 CaCl2, 10 glucose, and 10 HEPES; and intracellular (pipette) solution, 120 KCl, 1.13 MgCl2, 0.5 EGTA, 1 Na2ATP, 10 glucose, and 10 HEPES. For the extracellular solution used in the experiments with external free Ca2+, Ca2+ was removed from the extracellular solution and replaced with 1 mM EGTA. The fluids were superfused over the cell(s) by hydrostatic pressure-driven application (20–30 cmH2O) through polyethylene tubes. All solutions were at pH 7.2, and all experiments were carried out at room temperature (22–25°C).

Preparation of cADPR. cADPR was prepared enzymatically from NAD+ with ADP-ribosyl cyclase purified from ovotestis of Aplysia kurodai, a species common around the Japan coast (53). cADPR was then purified by HPLC and characterized by 1H nuclear magnetic resonance and fast atom bombardment mass spectrometry.

Ca2+ release assay. Microsomes were prepared as previously described (52, 53). In brief, cells were homogenized with a pellet mixer (Treff, Degersheim, Switzerland) in 0.2 ml of acetate intracellular medium composed of 250 mM potassium acetate, 250 mM N-methylglucamine, 1 mM MgCl2, and 20 mM HEPES (pH 7.2) supplemented with 0.5 mM ATP, 4 mM phosphocreatine, creatine phosphokinase (2 U/ml), 2.5 mM benzamidine, and 0.5 mM phenylmethylsulfonyl fluoride. After the homogenates had been centrifuged for 45 s at 13,000 g, the microsomes were prepared by Percoll density gradient centrifugation at 20,000 g for 40 min at 10°C. Ca2+ release was monitored in 3 ml of intracellular medium composed of 250 mM potassium gluconate, 250 mM N-methylglucamine, 1 mM MgCl2, and 20 mM HEPES (pH 7.2) supplemented with 1 mM ATP, 4 mM phosphocreatine, creatine phosphokinase (2 U/ml), 2.5 mM benzamidine, 0.5 mM phenylmethylsulfonyl fluoride, and 3 µM fluo 3, a fluorescent dye Ca2+ indicator. Fluo 3 fluorescence was measured at 490 nm excitation and 535 nm emission with a Jasco CAF-110 intracellular ion analyzer (Tokyo, Japan) at 37°C with a circulating water bath. The amounts of Ca2+ released were measured as the peak responses to either agent. We added a 3-µl aliquot of 1 µM standard Ca2+ solution into the cuvette, giving rise to a final amount of 3 nmol of Ca2+ in the bath. The Ca2+ released in response to IP3 and/or cADPR was estimated by comparing with the standard 3 nmol of Ca2+ added.

Detection of CD38 mRNA. Total RNAs were isolated as described previously (54). RT-PCR was carried out as described (54). The sequences of sense and antisense primers used for CD38 were 5'-GCTCTAGAGCCCTATGGCCAACTGCGAGTT-3' and 5'-GCTCTAGAGCTCAGATCTCAGATGTGCA-3', respectively (54), and those for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were 5'-CATTGACCTCAACTACATGGT-3' and 5'-TTGTCATACCAGG-AAATGAGC-5', corresponding to nucleotides 112–132 and 928–948 of human GAPDH cDNA (1), respectively. The PCR conditions were as follows: denaturation at 94°C for 30 s, annealing at 68°C (CD38) or at 60°C (GAPDH) for 1 min, and extension at 72°C for 2 min. The cycle was repeated 40 times for CD38 and 30 times for GAPDH as an internal control. The reaction mixture was analyzed on a 1.5% agarose gel in HCl/acetate/EDTA buffer.

Quantification of cellular IP3. The cellular content of IP3 was estimated with a D-myo-inositol 1,4,5-trisphosphate [3H] assay system (TRK 1000; Amersham Pharmacia Biotech UK, Buckinghamshire, UK) either in the presence or absence of ACh (10–9, 10–8, or 10–7 M). The suspension of enzymatically dispersed acinar cells in 1 ml of external solution (with or without ACh) in 2-ml test tubes was incubated for 1 min, and the supernatant was discarded after centrifugation at 2,000 g for 5 min at 4°C and stocked at –80°C. The cells were homogenized in 200 µl of ice-cold 10% (vol/vol) perchloric acid with an ultrasonic processor (Astrason; Heat Systems, Farmingdale, NY) for 10 s. The samples were then neutralized by the addition of 0.15 M KOH (1 ml), and a 2-µl aliquot was kept for protein quantification. After sedimentation of KClO4 by centrifugation at 2,000 g for 15 min at 4°C, the supernatant was used for IP3 measurement. The assay was performed according to the manufacturer's instructions.

Quantification of cellular cADPR. The cells were prepared with the same protocol as described above. The frozen pellet samples were homogenized in 370 µl of perchloric acid solution (2.5%, vol/vol) and centrifuged for 10 min at 13,000 g. cADPR was extracted and concentrated as follows (52). The supernatant (300–350 µl) of the homogenates was then mixed with 150 µl of a suspension of Norit A (27 mg/ml in H2O; Nacalai tesque, Kyoto, Japan). After a 30-min incubation at 37°C, the samples were again centrifuged, and the supernatant was discarded. The pellet was washed three times with 1.0 ml of H2O, resuspended in a pyridine/ethanol/H2O mixture (10:50:40, vol/vol/vol), and incubated for 120 min at 37°C. After a further centrifugation, the supernatant was collected and evaporated (Speedvac; Savant Instrument, Farmingdale, NY). The recovery of cADPR, monitored by the recovery of [3H]cADPR added in each homogenate, was 55.8 ± 2.61% (n = 20). Correction was introduced for the recovery of cADPR. The cADPR content of the cell extracts was measured by a radioimmunoassay. Briefly, the evaporated materials eluted from Norit A charcoal were resuspended with 50 µl of H2O and then incubated at 25°C for 2 h with bovine alkaline phosphatase and venom phosphoesterase (Worthington) at final concentrations of 50 and 2 U/ml, respectively, in 100 mM imidazole HCl (pH 7.5), 2 mM MgCl2, 100 mM NaCl, and 400 mM KCl. The reaction was terminated by adding a solution of trichloroacetic acid (at a final concentration of 4%, wt/vol) and was kept on ice for 20 min. A clear supernatant was obtained after centrifugation at 13,000 g for 10 min. An aliquot (10–20 µl) of the supernatant was immediately neutralized with a solution of 2 M Tris base and subjected to the procedure for cADPR measurement (52).

Animal care and human surgical specimen. This study was approved by the Animal Care and Use Committee and the Ethics Committee on Human Investigations of the Tohoku University School of Medicine. The care and handling of the animals were performed in accordance with National Institutes of Health guidelines for the care and handling of animals (60).

Reagents. Fluo 3 was purchased from Molecular Probes (Eugene, OR). Tetraethylammonium (TEA)-Cl and collagenase were from Wako Pure Chemicals, Osaka, Japan. HEPES was from Dojindo Laboratories, Kumamoto, Japan. Charybdotoxin was from Peptide Institute, Osaka, Japan. FK-506 was a generous gift from Fujisawa Pharmaceuticals, Osaka, Japan. All other reagents were from Sigma Chemical, St. Louis, MO.


    RESULTS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
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ACh-induced ionic currents in submucosal glands. The fresh submucosal gland acinar cells generated ionic currents in the presence of a wide range of ACh concentrations (4 x 10–9-10–6 M). Lower concentrations of ACh (up to several tens of nM) induced repetitive transient activations of the ionic currents recorded at Hp of 0 and –80 mV. The ionic current activated at 0 mV Hp (the upward deflection of the current-trace shown in Fig. 1A; Io) was abolished by charybdotoxin, a specific inhibitor of the Ca2+-activated large conductance K+ channel (BK channel) (39) (100–200 nM, n = 5; Fig. 1A). The downward deflection of the current recorded at –80 mV (Ii) was inhibited in the presence of diphenylamine-2-carboxylate (DPC, 1 mM, n = 6), a Cl channel inhibitor (2) (Fig. 1B). TEA (2–5 mM), a known inhibitor of BK channels (28), abolished both Io and Ii (data not shown, n = 8). As reported previously (20, 51), extremely high concentrations of ACh induced a sustained (not oscillatory) activation (see Fig. 2, A and B), and the sustained Io and Ii were carried by K+ and Cl, respectively, as evidenced by experiments using channel blockers and electrolyte-replacement studies (51). Thus the regenerative current spikes (or the oscillation) activated by low concentrations of ACh shared the same characteristics with the sustained currents stimulated by high doses of ACh. These ionic currents were abolished totally either with the extracellular membrane-permeable Ca2+ chelator BAPTA-AM (50 µM, n = 3) or with intracellular high EGTA (10 mM, n = 3), indicating Ca2+ dependence of the ACh-stimulated ionic currents (data not shown). As shown in Fig. 1, C–E, the oscillatory Ii increased in both frequency and amplitude as the concentration of ACh was raised, although the sensitivity to ACh was largely variable from cell to cell. A sustained current component became visible and was accompanied by oscillatory activation with stronger stimuli (Fig. 1, B and E).



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Fig. 1. Representative original recordings showing acetylcholine (ACh)-induced ionic currents in tracheal submucosal gland acinar cells. A conventional whole cell patch-clamp technique was used. A: membrane currents were monitored at 2 different holding potentials (Hp), i.e., 0 and –80 mV, each of which roughly corresponded to the Cl and K+ equilibrium potential, respectively. This dual current monitoring was accomplished by applying 200-ms voltage pulses of –80 mV at a frequency of 2 Hz to the pipette Hp of 0 mV. The upward or downward deflection of the current tracing represents outward (Io) or inward current (Ii), respectively. The periodic spikes imposed on the current traces (looking like a band of shadow) are parts of the capacitance currents of the plasma membrane. These artificial currents arise at the respective moments, just after the switching of the Hp from 0 to –80 mV, or the opposite. In B–G, ionic currents were monitored with Hp fixed at –80 mV so that only Ii was recorded. In the presence of low concentrations of ACh, the submucosal gland generated repetitive spiky ionic currents. A: Io was abolished by charybdotoxin (ChTx), a K+ channel blocker. B: Ii was abolished by diphenylamine-2-carboxylate (DPC), one of the Cl channel blockers. C–E: original current recordings showing the effect of increasing doses of ACh. These traces were obtained from the same cell with increasing concentrations of ACh. The oscillatory response was increased in frequency and amplitude with incremental ACh, giving rise to a sustained component mixed with the spiky currents (E). The sustained component is also shown in B, in which the activated Ii returned to baseline with the Cl channel blocker DPC. As discernible from these recordings, the sensitivity to ACh was variable from cell to cell, which may reflect differences in the cell population, i.e., serous and mucous cells. F and G: representative recordings showing the effects of extracellular FK-506 and ryanodine on the ACh-induced oscillatory Ii. Both the agents inhibited Ii in a reversible manner. FK-506 required a longer interval for the cell to regain activity.

 


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Fig. 2. Representative dual current recordings generated by intracellular infusion of inositol 1,4,5-trisphosphate (IP3) and/or cyclic ADP-ribose (cADPR) in human tracheal glands. Typical responses to a high concentration of ACh in human (A) and feline (B) submucosal glands are shown (top). Note the abrupt and large surge in Ii followed by a delayed activation of sustained Io, commonly observed in both species. In infusion experiments (C–I), the whole cell configuration was established at the point indicated ({bullet}) in each recording. C: 20 µM IP3 included in the pipette solution induced a current activation quite similar to the beginning of the ACh response but was followed by a rapidly decayed small current. D: 23 µM cADPR in pipette activated both Io and Ii but was distinct from those activated by extracellular high ACh or intracellular high IP3. E: IP3 combined with cADPR (20 µM each) in pipette induced a long-lasting large current with rapid onset. F and G: low IP3 or cADPR alone in pipette were without effects on the cells that could respond to extracellular ACh. H: a combined infusion of low IP3 and cADPR mimicked the oscillatory activation of the currents with low ACh. I: 6 µM cADPR in pipette induced a slow activation of Io alone.

 
Effects of FK-506 and ryanodine on the oscillatory Cl current. FK-506 (tacrolimus, 10 µg/ml), an immunosuppressant drug, abolished the ACh-induced oscillatory Cl current (n = 8, Fig. 1F). Ryanodine (10 µM), an agent that acts via the endoplasmic reticulum Ca2+ release channel ryanodine receptor (RyR), also inhibited Ii in a way similar to FK-506 (n = 12, Fig. 1G), although the duration required for the recovery was far longer with FK-506.

Actions of IP3 and/or cADPR introduced into cytoplasm. FK-506 has been shown to bind to one of the immunophilin FK-506-binding proteins (FKBP) associated with the RyR (29, 45) or IP3R (6), thus regulating the intracellular release of Ca2+. We, therefore, performed experiments with intracellular infusion of IP3 and/or cADPR, a putative RyR agonist, into human tracheal gland acinar cells. As shown in Fig. 2, we chose relatively high (>=20 µM, Fig. 2, C–E) and low (2–4 µM, Fig. 2, F–H) doses of IP3 (or cADPR) for infusing into the cytoplasm via the patch pipette. This was intended to reconstitute the bioelectric responses of the gland to ACh in high and low concentrations. The infusion of highly concentrated IP3 alone activated abrupt but transient responses in both directions (n = 7, Fig. 2C), which closely mimicked the initiation of ACh responses in both human and feline tracheal acinar cells (Fig. 2, A and B) but lacked the successive activation. A high dose of cADPR (23 µM) included in the patch pipette activated both Io and Ii (n = 7) with a gradual onset without the initial surge in Ii (Fig. 2D). In contrast, the combined infusion of IP3 and cADPR (both 20 µM) induced a rapid and long-lasting current in the presence of extracellular Ca2+ (n = 4, Fig. 2E), whereas the current decayed rapidly in the absence of external Ca2+ (n = 4, data not shown).

As shown in Fig. 2, F and G, infusions of 2–4 µM of IP3 alone or 2 µM of cADPR alone did not induce any responses up to 12 min after the establishment of the whole cell configuration (n = 11 for IP3 and n = 16 for cADPR). However, the combined infusion of IP3 and cADPR (2 µM each) evoked oscillatory current responses with quiet intervals of 1–2 min after the establishment of the whole cell configuration (4 positive responses out of 9 experiments, Fig. 2H). The pipette cADPR concentration required for the activation of the ionic current was >5 µM. As shown in Fig. 2I, 6 µM cADPR included in the pipette, however, did not induce an oscillation but stimulated gradually the Io alone without Ii (n = 3).

In a separate series of experiments, we applied cADPR on the cytosolic aspect of excised inside-out patches (n = 18). However, we did not find any channels activated in the presence of cADPR (4–20 µM). cADPR did not affect the activity of the Ca2+-dependent large conductance K+ channels contained in the isolated patches (n = 7).

Measurement of microsomal Ca2+ release. We then investigated the presence or absence of the intracellular Ca2+ pool that was sensitive to these agents. The microsome fraction obtained from the feline tracheal glands responded to both IP3 and cADPR to release Ca2+. IP3 and cADPR (1 µM each) released 0.65 ± 0.13 (n = 4, means ± SE) and 0.86 ± 0.10 nmol (n = 4) of Ca2+, respectively. The protein content in the cuvette was 105 µg/3 ml. The Ca2+ stores sensitive to either agent were independent of each other because the microsomes desensitized with IP3 by repetitive stimulations still responded to cADPR, or vice versa (n = 5, Fig. 3, A and B). The IP3-induced Ca2+ release was abolished by adding heparin (250 µg/ml) to the bath, but cADPR still released Ca2+ from the heparin-treated microsomes (n = 2, Fig. 3C).



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Fig. 3. Representative records of microsomal Ca2+ release assay. Microsome fraction was prepared from feline submucosal gland and used fluo 3 as a Ca2+ indicator (see METHODS). A: cADPR released Ca2+ in microsome desensitized to IP3. B: IP3 liberated Ca2+ in microsome desensitized to cADPR. C: heparin abolished the response to IP3 without affecting the potency of cADPR. Vertical scale bar at right of each panel represents 1 nM Ca2+.

 
Detection of CD38 mRNA. CD38 has been shown to catalyze the formation and degradation of cADPR (19, 25, 34, 54). To examine whether the airway glands have the ability to synthesize cADPR, we carried out experiments to detect the mRNA of CD38 using human materials, including submucosal glands (Fig. 4). CD38 mRNA was expressed strongly in human tracheal submucosal glands by RT-PCR (Fig. 4) as well as in exocrine pancreas but not in tracheal smooth muscle.



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Fig. 4. Expression of CD38 mRNA (A) in human tissues and cell lines. Lane 1, tracheal gland; lane 2, tracheal smooth muscle; lane 3, pancreas; lane 4, kidney; lane 5, NB-1 cells; lane 6, HeLa cells. RNAs from pancreas, kidney, NB-1 cells, and HeLa cells were the same RNAs that had been used for Northern blot hybridization to detect CD38 mRNA (33). PCR reactions using glyceraldehyde-3-phosphate dehydrogenase (GADPH; B)-specific primers demonstrated that similar amounts of cDNA had been used and similarly amplified.

 
Changes in cellular content of IP3 and cADPR by ACh. We next examined whether cADPR can occur naturally in the presence of physiological concentrations of ACh. We measured the cellular content of IP3 and cADPR stimulated for 1 min with 10–9 to 10–7 M ACh. The IP3 contents were not changed within the concentration range of ACh used, i.e., 116.6 ± 24.9, 126.8 ± 41.1, 85.7 ± 32.4, and 222.7 ± 81.6 nmol/µg protein for control, 10–9, 10–8, and 10–7 M of ACh, respectively (n = 4 each; P > 0.05 by ANOVA). The mean protein content in each sample was 124.0 ± 21.1 µg/ml (n = 16). In contrast, the cADPR content was markedly raised by ACh at a concentration as low as 10–9 M, i.e., the nontreatment control gland yielded 4.4 ± 1.2 fmol/µg of protein (n = 10) and was increased to 152.0 ± 40.5, 233.7 ± 54.9, and 295.0 ± 154.7 in the presence of 10–9, 10–8, and 10–7 M ACh, respectively (n = 3–4). These results are summarized in Fig. 5.



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Fig. 5. Changes in cellular content of IP3 and cADPR in the presence of ACh. The vertical axis on the right represents the IP3 concentration, and the left axis denotes the concentration of cADPR. SE is smaller than the symbol size in the first point in cADPR. {circ}, cADPR; {bullet}, IP3.

 

    DISCUSSION
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The airway submucosal gland is a mixed serous and mucous gland, and the serous phenotype consists of ~60% of the cell population (3). We could not discriminate the cell types under the present experimental settings. However, both cell types appeared to be activated by ACh to generate ionic currents because most of the cells (>=95%) responded to high concentrations of ACh (10–7 to 10–6 M) with a strikingly similar pattern, although ~20% of the cells failed to respond to a low concentration of ACh (10–8 M) (51, 55). Both the mucous and serous cells have been shown to respond equally to cholinergic stimuli to cause secretions in a rheological (33) and a morphological (58) experiment. In addition, an earlier bioelectric study performed on submaxillary salivary gland, which is a mixed gland similar to the submucosal gland, revealed that all the acinar cells responded to chorda tympani stimulation through randomized intracellular microelectrode recordings (36). These observations support our notion that the ionic currents could be induced by ACh regardless of the cell type.

The rise in intracellular Ca2+ concentration ([Ca2+]i) can activate Ca2+-sensitive ion channels on the plasma membrane, and, in exocrine acinar cells, the opening of Ca2+-gated Cl channels leads to a release of Cl out of the cell, which has been correlated with water secretion physiologically (47, 66). In the present study, ACh stimulated Io and Ii in tracheal submucosal glands, corresponding to the efflux of K+ and Cl from the cell, respectively, since Io was inhibited by charybdotoxin and Ii was blocked by DPC. In addition, the total replacement of K+ with Cs+ abolished Io, and the substitution of Cl with gluconate (or SO42–) inhibited the major fraction of Ii (51). Both the currents were dependent on [Ca2+]i because the membrane-permeable Ca2+ chelator BAPTA-AM abolished the currents and because the cells perfused with a high EGTA could no longer respond to ACh. In the presence of physiologically relevant concentrations of ACh, Io and Ii showed oscillatory activations (Fig. 1), the magnitudes of which increased with the augmentation of the stimulus, and finally gave rise to sustained activations with extreme concentrations of ACh (Fig. 2, A and B). These steps may reflect the regulation of fluid secretion in physiological situations. Interestingly, both the oscillatory Io and Ii in the presence of low-ACh concentrations were inhibited concurrently by TEA, a known BK channel blocker (28). A similar observation has been reported in sheep parotid gland (9) and was attributable to the atropine-like action of TEA.

Mobilization of intracellular Ca2+ in mammalian exocrine glands is mediated by two major mechanisms: IP3-induced Ca2+ release and Ca2+-induced Ca2+ release (CICR) (61), although the latter mechanism seemed absent in avian nasal gland (37). Accumulating evidence suggests that cADPR may be an endogenous regulator of CICR through RyR (15, 29, 30, 31, 41, 53, 56). In addition, FK-506 has been shown to disturb the Ca2+ release due to a competition with cADPR in binding to FKBP that is allied with the RyR (29, 45) or IP3R (6). In the present study, FK-506 and ryanodine reversibly abolished the oscillatory current response in submucosal gland (Fig. 1F). Moreover, the infusion of cADPR into the cell induced an activation of ionic currents (Fig. 2, D and I), and cADPR released Ca2+ from a Ca2+ store distinct from that of IP3 (Fig. 3). These results indicated the involvement of RyR in the ACh-induced response in submucosal gland. It is of note, however, that the pattern of activation in ionic currents by cADPR infusion was different from that induced by ACh. A relatively small amount of cADPR showed an outward K+ current (Io) alone (Fig. 2I), which closely resembled the response evoked by intracellular ryanodine or extracellular caffeine, as reported previously (51). The high dose of cADPR, on the other hand, caused Ii as well as Io (Fig. 2D). This may indicate that high doses of cADPR mobilize enough Ca2+ to activate Cl channels because the Ca2+ sensitivity of the exocrine Cl channel is lower by one factor than that of the K+ channel (13). In contrast, the infusion of a high concentration of IP3 evoked a current response quite similar to that initiated by a high concentration of ACh (Fig. 2C) but was followed by a rapid decay.

Low concentrations of IP3 (2 or 4 µM in the pipette solution) could not induce any response in tracheal gland acinar cells (Fig. 2F). This is a common observation in infusion studies in exocrine glands using patch-clamp methodology (24, 51, 56, 57, 61) in which 10–100 µM IP3 were included in the patch pipette and at least 5 µM were required to evoke current responses (24). It is very unlikely, however, that the agonists act by generating such relatively high IP3 levels in physiological conditions, since the microsomes liberated Ca2+ in the presence of a lower dose (1 µM) of IP3. We cannot reconcile the apparent discrepancy of dosage between the two experimental conditions. One possible explanation may be the difficulty of the agent in diffusing out of the pipette into the cytosol. In our previous study, heparin (mol wt 3,000) included in the pipette solution failed to affect the ACh response (51). Nevertheless, the microsome fraction treated with heparin showed an attenuation of Ca2+ release as also shown in the present experiments (Fig. 3). Presumably, heparin could not reach its effector sites due, perhaps, to the large-molecular-weight and/or to the electric charge. Another likely explanation for the necessity of an extremely high amount of IP3 in evoking the response is that an additional factor other than IP3 is involved in the signaling pathway employed by ACh.

Importantly, the combined application of IP3 and cADPR mimicked the responses induced by ACh both at higher concentrations (Fig. 2E) and lower concentrations (oscillatory response; Fig. 2H). These currents decayed rapidly in the absence of external Ca2+, indicating the requirement of a replenishment of the internal Ca2+ stores to maintain the continuous activation of the currents. Thus the ionic currents stimulated by ACh could be reconstituted by IP3 with cADPR but not by either agent alone.

In the present Ca2+-release assay, we demonstrated the presence of a cADPR-sensitive Ca2+ pool in submucosal gland microsomes (Fig. 3). The microsome fraction obtained from the whole gland is derived from morphologically distinct cell populations, i.e., serous and mucous cells. There is a possibility that IP3 and cADPR released Ca2+ from different cell types. However, if IP3 and cADPR were synthesized separately in distinct cell types by ACh, we would have found at least two patterns of current activations, because intracellular applications of IP3 or cADPR alone elicited different and distinct patterns of responses by the respective agents. Yet, as described above, almost all the cells responded to ACh with a pattern strikingly similar to each other that was mimicked by the combined infusion of IP3 and cADPR. This strongly suggested the coexistence of IP3- and cADPR-sensitive Ca2+ stores in a single cellular cytoplasmic compartment.

CD38 has been found to be involved in cADPR metabolism (19, 25, 54), acting as ADP-ribosyl cyclase for the synthesis of cADPR and as cADPR hydrolase for its degradation. We identified CD38 mRNA in submucosal gland, which suggests the possibility that the cells synthesize cADPR. In pancreatic {beta}-cells, the relative balance of the two catalytic activities is regulated by cellular ATP to control the cellular level of cADPR (54). We do not have any evidence, so far, concerning a regulatory mechanism of the cADPR concentration through muscarinic receptor activation. It has long been known that the cGMP level is raised after muscarinic receptor activation in exocrine gland (23), but the physiological relevance of the increased cGMP in these cells has not yet been established. Recently, cGMP was reported to increase [Ca2+]i by stimulating cADPR synthesis in sea urchin eggs (16). We also found that nitric oxide, a cGMP enhancer, may act as a potent secretagogue in human airway submucosal glands (44). It would be of interest to study the possibility that cGMP may act directly or via a cGMP-dependent phosphorylation on CD38 protein, resulting in the increase in cADPR. Whatever the possible mechanism, it was evident that the cellular content of cADPR was markedly increased in response to physiological ACh stimuli (Fig. 5). In contrast, the IP3 content was unchanged within the concentration range of ACh used in the present experiment (10–9 to 10–7 M). This was compatible with the report that measured 3H-labeled inositol phosphate formation in bovine submucosal gland (18). A significant increase in the cellular inositol phosphate was observed at 10–6 M or higher concentrations of carbachol. Similarly, in a study using physiologically relevant concentrations of cholecystokinin on pancreatic acini, the investigators could not detect a significant rise in IP3 with a radioreceptor assay, although an increase in [Ca2+]i was evident (38).

We have shown that cADPR operates in submucosal gland acinar cells by acting on a Ca2+ store distinct from that of IP3. The resultant increase in [Ca2+]i may activate K+ channels in the immediate vicinity of the cADPR-sensitive Ca2+ store. The activation of the K+ channels induces membrane hyperpolarization (51), forming an electrochemical environment favorable to the efflux of Cl ions (47) and to the influx of Ca2+ into the cytosol (40), thus assuring continuous Cl secretion. We propose that IP3 and cADPR are both involved in a cooperative manner in developing the ACh-induced response.


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This work was supported by Grant-in-Aid 09670593 for Scientific Research from the Ministry of Education, Science, Sports, and Culture, Japan.


    ACKNOWLEDGMENTS
 
We thank Dr. Hiroshi Okamoto for the cADPR measurements and for many helpful discussions. We also thank Brent K. Bell for reading the manuscript.


    FOOTNOTES
 

Address for reprint requests and other correspondence: K. Sasamori, Division of Respiratory and Infectious Diseases, Tohoku Univ. Graduate School of Medicine, 1-1 Seiryo-machi, Aoba-ku, Sendai 980-8574, Japan (E-mail: sasamori{at}int1.med.tohoku.ac.jp).

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. Section 1734 solely to indicate this fact.


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