Nicotinic acid adenine dinucleotide phosphate (NAADP) has recently been shown to act as a second messenger controlling intracellular Ca2+ responses in mammalian cells. Many questions remain regarding this signaling pathway, including the role of the ryanodine receptor (RyR) in NAADP-induced Ca2+ transients. Furthermore, the exact metabolic pathway responsible for the synthesis of NAADP in vivo has not been determined. Here, we demonstrate that the NAADP mediated Ca2+ release system is present in human myometrial cells. We also demonstrate that human myometrial cells use the NAADP second messenger system to generate intracellular Ca2+ transients in response to histamine. It has been proposed in the past that the NAADP system in mammalian cells is dependent on the presence of functional RyRs. Here, we observed that the histamine-induced Ca2+ transients are dependent on both the NAADP and inositol 1,4,5-trisphosphate signaling pathways but are independent of RyRs. The enzyme CD38 has been shown to catalyze the synthesis of NAADP in vitro by the base-exchange reaction. Furthermore, it has been proposed that this enzyme is responsible for the intracellular generation of NAADP in vivo. Using CD38 knockout mice, we observed that both the basal and histamine stimulated levels of NAADP are independent of CD38 and the base-exchange reaction. Our group is the first to demonstrate that NAADP is a second messenger for histamine-elicited Ca2+ transients in human myometrial cells. Furthermore, the NAADP mediated mechanism in mammalian cells can be independent of RyRs and CD38. Our data provides novel insights into the understanding of the mechanism of action and metabolism of this new second messenger system.
- cADP ribose
- inositol 1,4,5-trisphosphate
- endoplasmic reticulum
- ryanodine channel
- nicotinic acid adenine dinucleotide phosphate
- base-exchange reaction
the release of Ca2+ from intracellular stores is a widespread component of several signaling pathways (5, 13). Nicotinic acid adenine dinucleotide phosphate (NAADP) is a recently discovered nucleotide with intracellular Ca2+-releasing properties in several mammalian cells and tissues (10, 13, 22, 23). In fact, NAADP has been implicated as an intracellular second messenger (7, 9, 19, 29, 33, 40, 41).
In view of the potential role of NAADP as a second messenger, it is pivotal to determine the mechanisms of NAADP-induced Ca2+ release and its metabolism in mammalian tissues. In fact, many aspects of the NAADP signaling system in mammalian cells have yet to be defined. With regard to the mechanism of NAADP-induced Ca2+ transients, it has been proposed that NAADP induces Ca2+ release from lysosomal like Ca2+ stores (18, 29, 40). In addition, it has been proposed that the ryanodine receptor (RyR) is necessary for NAADP-induced Ca2+ release in mammalian cells (20, 25, 28). In fact, some authors (20, 24, 25, 28) have proposed that the RyR is the site of action of NAADP in mammalian cells. Here, we demonstrate that the agonist histamine induces Ca2+ transients in myometrial cells that are dependent on NAADP but independent of the RyR. This new finding indicates that, similar to what is observed in invertebrates (10, 16, 30), the RyR is not necessary for the effect of NAADP in mammalian cells.
Synthesis of this second messenger by the so-called base-exchange reaction has been described in several mammalian tissues, including the brain, heart, liver, spleen, and kidney (8, 11, 15). In the base-exchange reaction, the nicotinamide residue from NADP is replaced by a nicotinic acid resulting in the formation of NAADP (14). This reaction is catalyzed by NADases in the presence of the substrates NADP and nicotinic acid (14). Furthermore, it has also been reported (1) that ADP-ribosyl cyclase is capable of catalyzing the base-exchange reaction leading to synthesis of NAADP. This enzyme was first described as being responsible for the synthesis of another second messenger, cADPR, and a link between these two signaling pathways has been proposed (1, 11). In mammalian cells, ADP-ribosyl cyclase (CD38) is the major enzyme involved in the synthesis of cADPR in many mammalian tissues (38). Furthermore, CD38 has also been reported to catalyze the synthesis of NAADP (1, 11). In addition, we have recently described that CD38 is the major enzyme responsible for the in vitro synthesis of NAADP catalyzed by the base-exchange reaction in mammalian tissues (11). In fact, it has been proposed by many authors that the CD38 catalyzed base-exchange reaction is the physiological pathway for the synthesis of NAADP (1, 11). However, whether CD38 can indeed generate NAADP via the base-exchange reaction in vivo has not been described to date. Under the present experimental conditions, the concentrations of substrate needed for the base-exchange reaction, namely nicotinic acid, are several times higher than would be expected to be present in intact cells (14). Furthermore, the optimal pH for this reaction is out of the physiological range (1, 8). However, compartmentalization of nicotinic acid and NADP into an acidic environment could theoretically provide a possible milieu for the synthesis of NAADP in vivo. With the recent development of a sensitive and specific bioassay for the measurement of intracellular levels of NAADP (19), the question of whether CD38 and the base-exchange reaction are responsible for the in vivo synthesis of NAADP can be now approached.
In the present study, we provide experimental evidence to suggest that CD38 and the base-exchange reaction are not involved in the in vivo generation of NAADP. These data provide the framework for the discovery of the true physiological metabolic pathway for the in vivo synthesis of NAADP.
MATERIALS AND METHODS
CD38 wild-type and knockout mice.
CD38 knockout mice (C57BL/6J.129 CD38−/−, N12 back-cross) were produced as described previously (3) and were maintained in the Mayo Clinic and Trudeau Institute Animal Breeding facility in accordance with all Institutional Animal Care and Use Committee guidelines.
In accordance with procedures reviewed and approved by the Mayo Foundation Institutional Review Board, human myometrium was obtained from patients undergoing elective hysterectomy. Human myometrial cells were isolated using techniques previously described (4, 39). Briefly, the tissue was minced in Hanks’ balanced salt solution (HBSS) containing 1 g/l glucose and 10 mM HEPES (pH 7.4). The tissue was then suspended in fresh HBSS, aerated with 95% O2-5% CO2, and incubated in a 37°C water bath with gentle shaking for 2 h in the presence of 20 U/ml papain and 2,000 U/ml DNase. Subsequently, the tissue was incubated for an additional 1 h at 37°C, with the addition of 1 mg/ml type IV collagenase. Human myometrial cells were released by trituration, were centrifuged, and then were resuspended in Dulbecco’s modified Eagle’s medium (DMEM) containing 10% FBS, 100 U/l penicillin, 100 mg/l streptomycin, and 2.5 mg/ml amphotericin B. Cultures were grown and maintained in 75 cm2 plastic flasks in a humidified incubator supplied with 5% CO2-95% air at 37°C. Subcultures were obtained as needed by detaching the cells with a Ca2+/Mg2+-free HBSS solution containing 0.25% trypsin and 5 mM EDTA. Only cultures between passages 2 and 10 were used. Cells isolated by this procedure stained positive for α-smooth muscle actin and negative for keratin. For experiments, cells were made quiescent by replacement of the growth medium with DMEM without serum. Cell medium was again replaced with DMEM containing test reagents solubilized in 0.1% DMSO or water added to the final concentrations. HL60 cells were prepared as described previously (34).
“Global” intracellular [Ca2+] imaging in human myometrial cells.
Cultured human myometrial cells were plated onto 9 × 22 mm coverslips for spectrofluorometer microscopy at a density of 25,000 cells/coverslip and grown until ∼80% confluent in DMEM with 10% FBS. Cells were made quiescent by changing the medium to serum-free DMEM for 24 h and then incubated with 5 μM fura 2-AM for 2 h at 37°C. Cells were then washed in HBSS medium with or without Ca2+, depending on the experiment and imaged alternately with 340/380-nm light (emission 510 nm) using a spectrofluorometer (model F-2000; Hitachi, Tokyo, Japan).
Isolation of microsomes.
Microsomal fractions were isolated (at 0–4°C) from human myometrial tissue. The myometrium, minced with a razor blade, was suspended in a buffer containing (in mM) 300 sucrose, 10 HEPES, 0.1 EDTA, and 0.5 phenylmethylsulfonyl fluoride (pH 7.4), and homogenized with the use of a Polytron homogenizer. The homogenate was centrifuged for 10 min at 2,000 g, and the pellet was discarded. The supernatant was further centrifuged at 20,000 g for 20 min, and the supernatant thus obtained was centrifuged at 100,000 g for 1 h. The pellet was resuspended in a small volume of homogenizing buffer with a Dounce homogenizer. The microsomes were either used fresh for measurement of Ca2+ release or were divided into aliquots, quickly frozen, and stored at −70°C.
Ca2+ release in human myometrium microsomes.
Ca2+ uptake and release were measured in a medium containing 250 mM N-methyl glucamine, 250 mM potassium gluconate, 20 mM HEPES buffer (pH 7.2), 1 mM MgCl2, 2 U/ml creatine kinase, 4 mM phosphocreatine, 1 mM ATP, 25 μg/ml leupeptin, 20 μg/ml aprotinin, 100 μg/ml soybean trypsin inhibitor, and 3 μM fluo-3. With the use of a Hitachi F-2000 spectrofluorometer, fluo-3 fluorescence was monitored at 490-nm excitation and 535-nm emission in a 250-μl cuvette contained in a 37°C temperature-regulated cuvette holder and mixed continuously with a magnetic stirring bar. The addition of stock solutions of various reagents did not exceed 2% of the volume in the cuvette.
Confocal intracellular [Ca2+] imaging and microinjection of NAADP.
Dissociated human myometrial cells attached to coverslips were incubated in 5 μM fluo-3/AM (Molecular Probes, Eugene, OR) at 37°C for 30 min, and then placed on an open slide chamber (Warner Instruments, Hamden, CT) mounted on a Nikon Diaphot inverted microscope. The chamber was perfused with HBSS containing 1 mM Ca2+ at 2–3 ml/min at room temperature.
Fluo-3 loaded cells were visualized using an Odyssey XL real-time confocal system (Noran Instruments, Middleton, WI) attached to the Nikon microscope, and equipped with an Ar-Kr laser. In previous studies (37), we have used real-time confocal imaging to examine both spatial and temporal aspects of intracellular [Ca2+] ([Ca2+]i) dynamics. However, in the present study, the system was used predominantly to examine temporal aspects, within small regions of uterine smooth muscle cells. Our previous experiments have determined that 30 frames/s was sufficient to determine the dynamic [Ca2+]i response of myometrial cells without introducing frequency aliasing. An Olympus ×40/1.3 oil-immersion objective lens was used for imaging with image size set to 640 × 480 pixels (0.06 μm2/pixel). Optical section thickness was set to 1 μm, with regions of interest of 5 × 5 pixels (1.5 μm2).
Myometrial cells displaying stable basal [Ca2+]i levels (i.e., no spontaneous fluctuations in resting [Ca2+]i levels) were located, brought into the imaging field, and were impaled with single-lumen glass micropipettes (1.5 mm outer diameter; World Precision Instruments) that were pulled to a fine tip with the use of a Brown-Flaming electrode puller. The electrode tip resistance was found to be <100 KΩ when filled with NAADP concentrations in 150 mM KCl. The average tip diameter was estimated to be <5 μm. The NAADP solution was injected into the myometrial cells using pressure injection (PicoSpritzer; General Valve) during simultaneous monitoring of [Ca2+]i responses. Because the final NAADP concentration in the cell following injection was dependent on cell volume, the volumes of ∼10 myometrial cells was estimated a priori from online length and breadth measurements. On the basis of this cell volume, the total time and amplitude of the pressure pulse was then set such that the volume injected resulted in a final intracellular concentration of one of two values (1 μM and 1 mM).
Lysotracker red DND-99 and BODIPY-FL thapsigargin labeling.
Simultaneous fluorescent labeling of sarcoplasmic reticulum and lysosomes were performed using BODIPY-FL-thapsigargin (TG; B-7487, Molecular Probes) and Lysotracker red (L-7528; Molecular Probes). Cells were grown to confluence in 8-well chamber slides and the medium was changed to fresh growth medium before the fluorescent probes were added. The concentration of BODIPY-FL-TG and Lysotracker red were, respectively, 1 μM and 75 nM. The cells were incubated simultaneously with both labels for a period of 30 min to 1 h under growth conditions and imaging was done with an Olympus Fluoview confocal imaging station (BODIPY-FL-thapsigargin excitation was 488 nm and emission 540 nm and Lysotracker red excitation was 568 nm, and emission 590 nm). Cells were also labeled with each probe separately and the effect of unlabeled excess TG (10 μM) and 50 μM glycylphenylalanine-2-naphthylamide (GPN) were determined.
ADP-ribosyl cyclase activity was measured using the NGD technique as previously described (21). Microsomal preparations were incubated in a medium containing 0.2 mM NGD, 0.25 M sucrose, and 40 mM Tris·HCl (pH 7.2) at 37°C. Activity was determined using a fluorescence assay at 300 nm excitation and 410 nm emission. In key experiments, results were also confirmed with the use of NAD, which is the natural substrate of the enzyme.
Tissues were removed from mice that were under anesthesia, and immediately frozen in liquid nitrogen. The tissues were then resuspended in 10% TCA and homogenized on ice using a Potter-Elvehjelm tissue grinder and the resultant homogenate was centrifuged for 2 min, 12,000 g at 4°C. The supernatant was removed and TCA was extracted using water-saturated ether. The pH of the supernatant was then adjusted to 8.0, injected onto an HPLC and the NAADP fraction collected as stated previously. The sample was then dried under a vacuum, washed three times with 1 ml methanol, and resuspended in the same volume as injected onto the column using GluIM, composed of (in mM) 250 N-methyl-glucamine, 250 potassium gluconate, 1 MgCl2, and 20 HEPES, pH 7.2. NAADP content was then determined using a modification of the binding protocol stated previously using GluIM. Briefly, 25 μl of 0.5% (final concentration) sea urchin egg homogenate were aliquotted into 12 × 75 mm tubes, followed by 25 μl of cell extract. The reaction was allowed to incubate on ice for 30 min. [32P]NAADP (50,000 cpm/200 μl) was then added and the binding terminated after 20 additional minutes, as described previously, using ice-cold GluIM. Concentration of NAADP in the cellular extract is then calculated by comparing to a standard curve with known quantities of NAADP.
NAADP synthesis by the base-exchange reaction.
Membrane fractions (1 mg/ml) were incubated with 1 mM NADP and 40 mM nicotinic acid at 37°C in a buffer containing 40 mM triethanolamine-acetic acid buffer (pH 7.2). Aliquots (3–7 μl) were removed after different incubation times and NAADP content was determined using a combination of the sea urchin egg homogenate bioassay and HPLC analysis of nucleotides (10).
Sea urchin egg homogenate bioassay.
Homogenates from Lytechinus pictus eggs were prepared as described previously, with minor modification (10). Briefly, the eggs were obtained by injection of 0.5 M KCl into the coelomic cavity, and collected in artificial sea water. The jelly coats were washed from the eggs by several passages through an 80-mm mesh silk. The eggs were then washed once in artificial sea water, twice in Ca2+-free sea water containing 1 mM EGTA, twice in Ca2+-free water without EGTA, and once in GluIM. A suspension (25%, wt/vol) was prepared by homogenization with 4–5 strokes in a Dounce homogenizer with a type A pestle. The homogenate was then centrifuged for 10–12 s at 13,000 g at 4°C, and the supernatant was collected and stored in 1-ml aliquots at −70°C. Frozen homogenates were thawed in a 17°C water bath and diluted to 1.25% (vol/vol) with GluIM containing 2 U/ml creatine kinase, 4 mM phosphocreatine, 1 mM ATP, 3 μg/ml oligomycin, and 3 μg/ml antimycin.
HPLC analysis of nucleotides.
The synthesis of NAADP and cADPR by mammalian tissue extracts was verified by HPLC analysis, performed by anion-exchange chromatography using an AG MP-1 column (Bio-Rad) eluted with a nonlinear gradient of trifluoroacetic acid, as described previously (10). The nucleotides were detected by UV absorption at 254 nm. The authenticity of the NAADP and cADPR produced were confirmed by co-elution with NAADP and cADPR standards and by the sea urchin egg homogenate bioassay. The NAADP and cADPR used on our study were at least 99% pure as determined by HPLC analysis.
Western blot analysis.
Human myometrial cell and HL60 extracts were incubated in lysis buffer containing 0.05% IGEPAL-CA 630, 20 mM EDTA, 20 mM NaCl, and 20 mM Tris, pH 7.0. After 10% SDS-PAGE, protein was electroblotted onto PVDF membrane, blocked with 5% nonfat milk for 1 h and probed with 1/100 dilution of mouse monoclonal antibody against human CD38 (catalog no. SC-7325, Santa Cruz) for 4 h. The immunoreactive bands were detected using a 1/20,000 dilution of horseradish peroxidase-conjugated anti-mouse IgG (catalog no. SC-2020, Santa Cruz) as a secondary antibody and an enhanced chemiluminescence detection system. Western blot analysis of mice tissues was performed with the antibody SC-7049 antibody from Santa Cruz. This antibody reacts with mouse CD38.
Detection of cADPR levels in cells and tissues by cycling assay.
Mouse tissue or human cells were frozen in liquid N2, pulverized into a powder, and extracted with 10% trichloroacetic acid (TCA) at 4°C. TCA was removed with water-saturated ether. The aqueous layer containing the cADPR was removed and adjusted to pH 8 with 1 M Tris. To remove nucleotides, except cADPR, a mixture containing hydrolytic enzymes was added to the samples with the following final concentrations: 0.44 U/ml pyrophosphatase, 12.5 U/ml alkaline phosphatase, 0.0625 U/ml NADase, 2.5 mM MgCl2, and 20 mM sodium phosphate, pH 8.0. The detection of cADPR was performed by some modification of the cycling method described recently (26).
All other reagents, of the highest purity grade available, were supplied from Sigma (St. Louis, MO), except when stated otherwise. The lysosomal inhibitors GPN, bafilomycin A (BafA1), monesis, and the Ca2+ ATPase inhibitors TG and cyclopiazonic acid (CPA) were of the highest purity available.
The reported experiments were repeated at least three times, data are expressed as means ± SE or SD. Student’s t-test was used to evaluate statistical significance; P values <0.05 were considered significant.
NAADP-induced Ca2+-releasing system in myometrial cells.
We have previously described the presence of the NAADP Ca2+-releasing system in smooth muscle cells (29, 42). Here we found that human myometrial cells also contain the NAADP system. Microinjection of micromolar concentrations of NAADP into myometrial cells leads to the development of a cytoplasmic rise in intracellular free Ca2+, as described before for pulmonary smooth muscle cells (Fig. 1A) (6, 29). However, as described before, the microinjection of high concentrations of NAADP (1 mM, HNAADP) had no effect on intracellular Ca2+ transients in myometrial cells (Fig. 1A). In fact, it has been proposed that the NAADP receptor from mammalian cells can be inactivated by high concentrations of NAADP itself (7).
To further characterize the NAADP system we used human myometrial cell microsomes. We observed that NAADP can trigger Ca2+ release from microsomes. The NAADP-induced Ca2+ release in microsomes was inhibited by the lysosomal inhibitor GPN, which causes lysis of lysosomes (Fig. 1, B and C), and by another lysosomal inhibitor BafA1 (data not shown). In contrast, GPN had no effect on the Ca2+ transients induced by cADPR or inositol 1,4,5-trisphosphate (IP3; Fig. 1C and data not shown). The NAADP-induced Ca2+ release in microsomes was not affected by the sarcoplasmic reticulum (SR) Ca2+ ATPase inhibitor TG (Fig. 1B). In contrast, pretreatment of microsomes with TG lead to a complete inhibition of the IP3 and cADPR-induced Ca2+ release (data not shown). NAADP-induced Ca2+ release in microsomes was not affected by the IP3 inhibitor (xestospongin C) or by the cADPR inhibitor (8-Br-cADPR) (Fig. 1D). It has been previously shown that NAADP-induced Ca2+ release can be inhibited by L-type Ca2+ channel blockers and by high (mM) or low (<1 nM) concentrations of NAADP itself (3, 36). The channels activated by NAADP show a bell-shaped activation curve, where suboptimal or supraoptimal concentrations of NAADP are inhibitory to channel activation and optimal NAADP concentrations vary from one cell type to another. Here, we found that preincubation of human myometrial microsomes with nifedipine, a L-type channel blocker, or 1 mM NAADP (HNAADP) leads to a complete inhibition of the subsequent Ca2+ release induced by 1 μM NAADP (Fig. 1D). In contrast, high concentrations of NAADP had no effect upon the Ca2+ release induced by IP3 or cADPR (Fig. 1E). These data indicate that the NAADP system is present in myometrial cells and that it is independent of the RyR channel. Furthermore, it indicates that the NAADP Ca2+ stores are derived from lysosomes.
Lysosomal Ca2+ stores in myometrial cells: role in histamine-induced Ca2+ transients.
As previously described and demonstrated above, the NAADP Ca2+ stores are derived from lysosomal-like structures (18, 29, 41). To determine the role of lysosomes and NAADP on agonist-stimulated Ca2+ transients in human myometrial cells, we first characterized the lysosomal Ca2+ stores in these cells. Using a double stain with the lysosomal marker lysotracker red and with the SR marker BODIPY-TG, we observed that myometrial cells contain both SR and lysosomal like structures (Fig. 2A). These lysosomal like structures occur with a mostly perinuclear distribution in contrast with the more diffuse distribution of the SR (Fig. 2A). Furthermore, we confirmed that the lysotracker red stain was specific for lysosomes because the lysosomal inhibitor GPN can completely abolish the stain obtained with lysotracker (Fig. 2C). In contrast, TG had no effect on the lysotracker stain (data not shown). The reverse was also true, staining of SR was unaffected by GPN (Fig. 2C) while being abolished by unlabeled TG (data not shown). To further assess the presence of lysosomal Ca2+ stores in myometrial cells, we determined the effects of GPN and TG on intracellular Ca2+ in intact human myometrial cells. We observed that both GPN and TG caused a transient increase in intracellular Ca2+ (Fig. 2B). To determine the role of the SR Ca2+-ATPase on the presence of Ca2+ in lysosomes and also to address any overlap between the SR and the lysosomal Ca2+ stores, we determined the effect of TG on the lysosomal Ca2+ content. We found that incubation of myometrial cells with TG leads to a transient increase in intracellular Ca2+ consistent with depletion of SR Ca2+ stores (Fig. 3A). However, after the intracellular free Ca2+ returns to its baseline further addition of TG fails to promote more Ca2+ release, indicating that the SR were completely depleted by the first addition of TG (Fig. 3A). In contrast, preincubation with GPN had no effect upon the TG-induced Ca2+ transients, indicating that the lysosomal Ca2+ stores are independent from the SR stores (Fig. 3B). All of these experiments were carried out in the absence of extracellular Ca2+, so the observed increases in intracellular Ca2+ had to be derived from intracellular stores. We also observed that the Ca2+ release induced by TG can be prevented by preincubation with another SR Ca2+-ATPase inhibitor CPA but not with BafA1, an inhibitor of the lysosomal H+-ATPase, or GPN (Fig. 3C). In contrast, the Ca2+ release induced by GPN was abolished by pretreatment with both GPN and BafA1 but not with CPA or TG (Fig. 3D).
To further determine the role of lysosomal Ca2+ stores in myometrial cells, we studied the effect of lysosomal Ca2+ inhibitors on agonist-induced Ca2+ transients. We first characterized the role of different intracellular and extracellular signaling pathways on the Ca2+ release induced by oxytocin, histamine, and ACh (Fig. 4). The Ca2+ release induced by histamine was only slightly inhibited by removal of extracellular Ca2+ and almost abolished by depletion of the SR stores with TG (Fig. 4B). These data indicate that intracellular Ca2+ mobilization and not extracellular Ca2+ influx play a major role in histamine induced Ca2+ transients. Furthermore, histamine-induced Ca2+ release was inhibited by the IP3 antagonist xestospongin C (Fig. 4, A and B), but not by ryanodine. In contrast, we observed that the oxytocin-induced Ca2+ transients in myometrial cells can be almost completely abolished by removal of extracellular Ca2+ (Fig. 4A). These data indicates that both extracellular Ca2+ influx and intracellular Ca2+ release play a role in oxytocin-induced Ca2+ transients. Ca2+ responses induced by ACh were not significantly reduced by the omission of extracellular Ca2+ and were inhibited by inhibitors of both IP3 receptor (xestospongin C) and cADPR-RyRs pathway (8-Br-cADPR and ryanodine) (Fig. 4A), these data indicate that the ACh-induced Ca2+ release is dependent on intracellular Ca2+ stores regulated by cADPR and IP3.
We further determined the role of the lysosomal Ca2+ store on oxytocin and histamine-induced Ca2+ transients. Using inhibitors of the lysosomal Ca2+ accumulation, we observed that both the histamine- and oxytocin-induced Ca2+ transients in myometrial cells were dependent on lysosomal stores (Fig. 4, B and C). In Fig. 4B, we determined the effect of treatment with several lysosomal inhibitors upon the histamine-induced Ca2+ release, including GPN, BafA1, and monesin. This data clearly indicates that lysosomal Ca2+ contributes to Ca2+ transients induced by certain agonists in myometrial cells. To confirm the specificity of the lysosomal inhibitors, we determined their effects on ACh-induced Ca2+ transients in myometrial cells, observing that they had no effect on ACh-induced Ca2+ transients (Fig. 4C).
Our data provides a chance to understand the role of lysosomal Ca2+ stores in RyR-dependent and RyR-independent Ca2+ transients. Furthermore, if one assumes that lysosomal Ca2+ release is only mediated by NAADP it is possible to extrapolate our findings to the coupling between NAADP and other channels. It has been previously described that NAADP-induced Ca2+ transients may be mediated by direct stimulation of RyRs or by a mechanism coupled with the NAADP-dependent Ca2+ stores and the Ca2+-induced Ca2+ release (CICR) system mediated by RyRs (20, 24, 25, 28). In fact, it has been proposed that NAADP-induced Ca2+ transients are completely dependent on the expression of RyR in cells (20, 24, 25, 28). Here, we observed that NAADP can work in a RyR-dependent and -independent manner. Oxytocin-mediated Ca2+ transients appear to be dependent on both IP3 and cADPR-RyR function (Fig. 4A). In contrast, inhibitors of the RyR-cADPR system had no effect upon the histamine induced Ca2+ transients in myometrial cells (Fig. 4A). Furthermore, the histamine induced Ca2+ transients can be inhibited by the IP3 inhibitor xestospongin C (Fig. 4, A and B). These data indicate that the lysosomal Ca2+ stores can work in a RyR-independent manner and that they may be coupled to the IP3 channel. It is possible that the NAADP and the IP3 system can be coupled by a CICR mechanism because the IP3 system also behaves as a CICR (16).
NAADP as a second messenger.
To further determine whether NAADP is a second messenger in myometrial cells, we determined NAADP levels in cells and examined the effect of histamine on the intracellular accumulation of NAADP. We observed that NAADP is a nucleotide present in myometrial cells and that stimulation of cells with histamine leads to a severalfold increase in intracellular levels of NAADP (Fig. 5). We further characterized the time course and the coupling between histamine and NAADP accumulation (Fig. 5, inset). Stimulation of myometrial cells with histamine led to a rapid increase in NAADP levels that peaked at 30 s and declined back to basal levels in 1 min (Fig. 5, inset). Furthermore, the histamine-induced NAADP accumulation was inhibited by the H1 receptor antagonist diphenhydramine (Fig. 5). We also observed that the NAADP accumulation induced by histamine appears to be Ca2+ and calmodulin dependent, as the accumulation of NAADP was inhibited by the Ca2+ chelator BAPTA and the calmidazolium, a calmodulin antagonist (Fig. 5). These data were surprising because the only enzyme known to generate NAADP in mammalian cells, CD38, appears not to be Ca2+ or calmodulin dependent (data not shown). These data led us to explore the role of the enzyme CD38 on the in vivo generation of NAADP.
CD38 is not necessary for histamine-induced NAADP accumulation in myometrial cells.
It has been previously shown that the enzyme CD38 is capable of generating NAADP in vitro (1, 11). CD38 catalyzes the so-called base-exchange reaction, where the nicotinamide from NADP is substituted by nicotinic acid-generating NAADP. To date, CD38 and the base-exchange reaction are the only enzyme and reaction known to be able to generate NAADP (1, 10). CD38 is also the enzyme responsible for the synthesis of the second messenger cADPR (1, 11). We observed that extracts prepared from the tissues of CD38 knockout mice are not capable of synthesis of NAADP (by the base-exchange reaction) or cGDPR in vitro (Fig. 6). cGDPR is a marker of the activity of the ADPribosyl cyclase from CD38 (11), in previous studies (2, 10, 11), we have shown that synthesis of cADPR from NAD is nearly abolished in several tissues from CD38 knockout mice. Furthermore, we determined the levels of NAADP and cADPR present in situ in tissues from both wild-type and knockout mice. Not surprisingly, the presence of intracellular levels of cADPR is essentially eliminated in CD38 knockout tissues (Fig. 7), the only tissue where cADPR levels were detected in equal amounts in both wild-type and CD38 knockout mice was in the brain (data not shown). However, the in vivo cellular content of NAADP was not reduced by the absence of CD38 (Fig. 7). The data from Figs. 8 and 9 indicate that neither CD38 nor the base-exchange reaction is necessary for the in vivo generation of NAADP.
We further determined the role of the CD38 enzyme upon the agonist (histamine)-induced intracellular in vivo accumulation of NAADP. Using myometrial cells cultured from CD38 wild-type and CD38 knockout mice, we determined the role of this enzyme on the histamine-induced accumulation of NAADP in vivo. As shown in Fig. 8, inset, CD38 is normally expressed in myometrial cells from wild-type mice but is absent in myometrial cells from CD38 knockout mice. Furthermore, the absence of CD38 does not prevent the in vivo generation and accumulation of NAADP in response to histamine (Fig. 8); as expected, histamine-induced Ca2+ transients are not altered by the absence of CD38 (Fig. 8B). In fact, myometrial cells from the CD38 knockouts appear to have a higher content of NAADP than wild-type cells in both the basal and agonist-stimulated states (Fig. 8). At present, the mechanism behind this is not known, but one possibility is compensation for the absence of the second-messenger cADPR. These data indicate that CD38 is not necessary for basal and agonist-induced generation of NAADP in vivo.
We also determined the effect of overexpression of the enzyme CD38 upon intracellular levels of NAADP in cells. Treatment of HL60 cells with retinoic acid (RA) has been shown to increase the expression of CD38, and subsequently the synthesis and intracellular levels of cADPR (Fig. 9, A–C) (34). Furthermore, overexpression of CD38 with RA leads to an increase of the in vitro generation of NAADP catalyzed by the base-exchange reaction (Fig. 9D). In contrast, the intracellular levels of NAADP were not increased by overexpression of CD38 with RA (Fig. 9E). Our data presents a new aspect of the NAADP second messenger system, indicating that CD38 is not the enzyme responsible for the in vivo generation of this nucleotide. Our experiments open a new avenue for the discovery of the in vivo pathway responsible for the synthesis of NAADP.
The present study demonstrates that the NAADP system is present and functional in human myometrial cells, and may play a role as a second messenger transducing the generation of [Ca2+]i transients in response to histamine and oxytocin. Furthermore, we clearly indicated that the acidcalciossomes (the NAADP-sensitive Ca2+ stores) are present in myometrial cells and play a role in intracellular Ca2+ homeostasis. In contrast with other authors (20, 24, 25, 28), that proposed that the NAADP-system is dependent on the RyR, we provide evidence that the NAADP system can also work in a RyR-independent way, however, more direct work needs to be done to further determine the role of the RyR on the NAADP functions. Recently, extracellular effects of NAADP have also been described (27); however, in our studies, extracellular application of NAADP has not been explored and the possible role of lysosomal and IP3 channels upon extracellular effect of NAADP in myometrial cells cannot be assumed without experimental evidence.
The majority of the experiments in the first part of our study were obtained with the use of pharmacological inhibitors. In these regard, our conclusions have the limitations of any studies that depend heavily on the use pharmacological inhibitors. However, the inhibitors used here have been extensively used by us and others and have been shown to have an acceptable specificity for such studies (19, 29, 41).
In another aspect of our study we determined the role of the enzyme CD38 on the in vivo accumulation of NAADP. We (8, 11, 12, 15) previously described the synthesis of NAADP in several tissues including brain, liver, spleen, heart, and kidney glomeruli. Synthesis of NAADP can be catalyzed in vitro by a NAD(P)ase, analog to the lymphocyte antigen CD38 (8, 11, 12, 15), in a reaction called the base-exchange reaction (Fig. 12; see Ref. 13). The enzyme catalyzes the exchange of nicotinamide for nicotinic acid on the molecule of NADP+, generating NAADP (Fig. 10; see also Ref. 13).
For these reasons, it is important to consider other theoretical pathways for the synthesis of NAADP in vivo (Fig. 10). Conceivably, NAADP might be generated by deamination of NADP+ (Fig. 10) or phosphorylation of NAAD+. The latter is a particularly attractive hypothetical route because NAAD+ is a compound present in cells and NAADP might be then catalyzed by NAD+ kinase with ATP as a 2′-phosphate donor (Fig. 10). These alternative synthetic pathways ought to be explored in future studies. Notably, Lerner et al. (32) characterized a human NAD+ kinase in vitro and found no evidence that it could synthesize NAADP by phosphorylation of NAAD+. Nevertheless, these data do not completely exclude the possibility that NAAD+ phosphorylation might perhaps occur in vivo, for example, if putative intracellular cofactors and/or other physiological conditions exist in vivo that are required to enable the reaction. They also do not exclude the possibility that other isoforms might catalyze the reaction. Therefore, the postulated NAAD+ phosphorylation pathway seems unlikely at this point but cannot be ruled out.
Despite the limitations discussed, the base-exchange reaction is the only pathway currently described for the synthesis of NAADP in biological systems (14). In this regard, an important observation is that enzymes with ADP ribosyl cyclase activity (capacity for synthesis of cADP ribose) are also able to catalyze the synthesis of NAADP through the base-exchange reaction (14). In fact, the mammalian version of ADP-ribosyl cyclase, CD38, is capable of generating both NAADP and cADPR (14). This observation led to the proposal of cross talk between these two signaling pathways. However, as discussed above, it is still unknown whether the base-exchange reaction occurs under physiological conditions. Using CD38 knockout mice, we determined that CD38 is the major enzyme responsible for the base-exchange reaction in mouse tissues in vitro (11, 12). However, in that study, the capacity for synthesis of NAADP by the base-exchange reaction in cells did not correlate with the presence of NAADP-induced Ca2+ release in the same cells. As a result, this discrepancy raised doubts about the role of the base-exchange reaction as the physiological route for the synthesis of NAADP. Our present data clearly indicates that there is no correlation between the expression of CD38 and the in vivo intracellular content of NAADP in basal and agonist induced NAADP accumulation. It is important for future work to explore and determine the true pathway responsible for the in vivo generation of this new intracellular second messenger.
This study was supported by The Mayo Foundation and by an American Heart Association grant (to E. N. Chini).
↵* S. Soares and M. Thompson contributed equally to this study.
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