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Signal Transduction Laboratory, Department of Anesthesiology (H.B., M.T., W.Z., S.F., E.N.C.), and Cell Imaging and Physiology Laboratory, Department of Physiology and Biophysics (Y.S.H., C.B.M., Y.S.P., G.S.), Mayo Clinic, Rochester, Minnesota 55905
Address all correspondence and requests for reprints to: Eduardo Nunes Chini, Signal Transduction Laboratory, Department of Anesthesiology, Mayo Clinic and Foundation, Rochester, Minnesota 55905. E-mail: chini.eduardo{at}mayo.edu.
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Abstract |
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regulation of oxytocin-induced [Ca2+]i transients, CD38 cyclase activity, and CD38 expression in human myometrial cells. We found that oxytocin-induced [Ca2+]i transients were significantly increased by 50 ng/ml TNF. Similarly, CD38 mRNA levels, CD38 expression, and cyclase activity were increased by TNF, thus increasing cADPR levels. We propose that a complex interaction between multiple signaling pathways is important for the development of intracellular Ca2+ transients induced by oxytocin and that TNF may contribute for the myometrium preparation for labor by regulating the cADPR-signaling pathway. The observation that the cADPR-signaling pathway is important for the development of intracellular Ca2+ transients in human myometrial cells raises the possibility that this signaling pathway could serve as a target for the development of new therapeutic strategies for abnormal myometrial contraction observed during pregnancy. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResults and DiscussionReferences |
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Intracellular Ca2+ ([Ca2+]i) regulation is a key factor in the modulation of uterine contraction (1, 2, 3, 4). To date, the mechanisms regulating [Ca2+]i homeostasis in human myometrial cells have not been completely elucidated (3, 4). Understanding the signaling pathways regulating agonist-stimulated [Ca2+]i transients in myometrial cells is imperative for the development of new therapeutic approaches to treat pathophysiological myometrial contraction.
Oxytocin is a naturally occurring peptide responsible for myometrial contraction during labor (1, 2, 3, 4). Oxytocin is also frequently used as a pharmacological agonist to increase uterine contraction during dysfunctional labor (1). Oxytocin-induced uterine contraction is initiated by an increase in [Ca2+]i (1, 3, 4). In myometrium, both influx of extracellular Ca2+ and mobilization of Ca2+ from intracellular stores are important for the generation of oxytocin-stimulated [Ca2+]i transients in myometrial cells (1, 3, 5, 6).
In human myometrium, both inositol 1,4,5-trisphosphate (IP3) and ryanodine receptor (RyR) channels are present (7). However, to date, only the IP3 signaling pathway has been extensively studied in human myometrial cells (4). The precise role of RyR channel-mediated Ca2+ release in human myometrium has not been completely elucidated (4, 8, 9, 10, 11). In addition, the intracellular mechanisms regulating RyR channel gating in human myometrium have not been examined. Cyclic ADP-ribose (cADPr) is an endogenous activator of the RyR channel in mammalian cells (for review, see Refs. 12, 13, 14, 15). In previous studies in human myometrium, we found that both cADPR and the bifunctional enzyme CD38 (capable of both synthesis and degradation of cADPR) are present and functional in human myometrium (16, 17).
In the present study, we explored the role of the cADPR signaling pathway in oxytocin-induced [Ca2+]i transients. In addition, we investigated the modulation of the CD38-cADPR-signaling pathway by cytokines such as TNF. We found that the cADPR system is crucial for the oxytocin-induced [Ca2+]i transients in human myometrial cells. Furthermore, we demonstrate that TNF regulates CD38 at the transcriptional level, increasing oxytocin-induced [Ca2+]i transients. We propose that pharmacological approaches targeting the cADPR-signaling pathway may lead to new therapies for dysfunctional and preterm labor.
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Materials and Methods |
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-smooth muscle actin and negative for keratin. For experiments, cells were made quiescent by replacing the growth medium with SmBM without serum or growth factors. Cell medium was again replaced with SmBM containing testing agents solubilized in 0.1% dimethylsulfoxide or water added to the final concentrations.
Isolation of microsomes
Microsomal fractions were isolated (at 0–4 C) from human myometrial tissue. Myometrium, minced with a razor blade, was suspended in a buffer containing 300 mM sucrose, 10 mM HEPES, 0.1 mM EDTA, and 0.5 mM phemylmethylsulfonyl fluoride (pH 7.4) and homogenized by a Polytron homogenizer. The homogenate was centrifuged for 10 min at 2,000 x g, and the pellet was discarded. The supernatant was further centrifuged at 20,000 x g for 20 min, and the supernatant thus obtained was ultracentrifuged at 100,000 x g for 1 h. The pellet was resuspended in a small volume of homogenizing buffer by Dounce homogenizer. The microsomes were either used fresh for measurement of Ca2+ release or divided into aliquots, quickly frozen, and stored at -70 C for measurement of [3H]-ryanodine binding. Storage at -70 C preserved the [3H]-ryanodine binding capacity.
Ryanodine binding and labeling
[3H]-ryanodine binding was performed using a filtration method (18). In brief, microsomal fractions (100–200 µg protein) were incubated in a medium containing (final concentrations) 600 mM KCl, 100 µM EGTA, 0.2 mM phemylmethylsulfonyl fluoride, and 25 mM HEPES (pH 7.2) and saturating concentrations of [3H]-ryanodine (180 nM) for 30 min at 35 C. Reaction was stopped by applying sample to a glass fiber filter and washing under vacuum three times with ice-cold water. The radioactivity remaining in the filter was determined with standard scintillation counter techniques. Fluorescent labeling of RyR was performed using BODIPY-TR-ryanodine antibody (B 13802; Molecular Probes, Eugene, OR). Human myometrial cells were plated on collagen-coated coverslips and grown as stated above. On reaching quiescence, the cells were rinsed in PBS, fixed in 4% paraformaldehyde for 15 min, and rinsed again in PBS three times for 5 min each. Cells were permeabilized in PBS containing 0.1% Triton X-100 for 30 min, followed by three washings in PBS for 5 min. The cells were then incubated in PBS containing 1% BSA, with or without 100 µM ryanodine, for 30 min, followed by incubation with 2 µM BODIPY-TR-ryanodine for 60 min. Coverslips were rinsed three times in PBS for 5 min, allowed to air dry, and mounted on glass microscope slides. The cells were then imaged on an Fluoview confocal imaging station (575 nm excitation and 613 nm emission; Olympus, Tokyo, Japan).
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, 4 mM pi, 25 µg/ml leupeptin, 20 µg/ml aprotinin, 100 µg/ml soybean trypsin inhibitor, and 3 µM fluo-3. Using an F-2000 spectrofluorimeter (Hitachi, Indianapolis, IN), fluo-3 fluorescence was monitored at 490 nm excitation and 535 nm emission in a 250-µl cuvette, while in a 37 C circulation water bath, 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 [Ca2+]i imaging
Detailed techniques for real-time confocal imaging of [Ca2+]i in smooth muscle cells have been described previously by our laboratory (19, 20). Briefly, human myometrial cells were plated on 15-mm glass coverslips coated with rat tail collagen type 1 at a concentration of 2.5 x 104 cells/coverslip, grown until approximately 70% confluent, and made quiescent as stated previously. Coverslips with attached cells were incubated in SmBM containing 5 µM fluo 3-AM (Molecular Probes) and 1 µM pluronic acid at 37 C for 60 min, rinsed briefly in HBSS, and then placed on an open slide chamber (Warner Instruments, Hamden, CT). Cells were visualized using an Odyssey XL real-time confocal system (Noran Instruments, Middleton, WI) attached to a microscope (Nikon, Tokyo, Japan) and equipped with an Ar-Kr laser (488 nm excitation and 515 nm emission). An Olympus x40, 1.3 numerical aperture, oil-immersion objective lens was used for imaging, with image size set to 640 x 480 pixels (0.06 µm2/pixel). Optical section thickness was set to 1 µm. Based on previous calibrations of [Ca2+]i, a fixed combination of laser intensity (30% of maximum) and photomultiplier gain (1800 from a maximum of 4096) was set a priori to ensure that pixel intensities within regions of interest ranged from 25 to 255 gray levels. Eight regions of interest of 5 x 5 pixels (1.5 µm2) were defined within the cells and release of [Ca+2]i was initiated by addition of HBSS containing 1 µM oxytocin. The gray level data were then converted to nanomoles Ca+2 on the basis of a previously described calibration procedure (19, 20).
Effect of antagonists on [Ca+2]i response to oxytocin
Human myometrial cells were preexposed to various concentrations of 8-Br-cADPR, a selective inhibitor of the cADPR receptor; nicotinamide, an inhibitor of ADP-ribosyl cyclase; or 10 µM dantrolene, a RyR channel inhibitor for 60 min. The [Ca+2]i response to 1 µM oxytocin was then evaluated as described previously.
Cyclase activity
Activity of the ADP-ribosyl cyclase was performed using the nicotinamide guanine dinucleotide (NGD) technique as described previously (21). Enzyme 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 fluorometric assay at 300 nm excitation and 410 nm emission (21). In key experiments, results were also confirmed with the use of nicotinamide adenine dinucleotide as the natural substrate of the enzyme.
HPLC analysis of nucleotides
The synthesis of cADPR by human myometrium was verified by HPLC analysis and performed by anion-exchange chromatography using an AG MP-1 (Bio-Rad Laboratories, Hercules, CA) column eluted with a nonlinear gradient of trifluroacetic acid, as described previously (22). The nucleotides were detected by UV absorption at 254 nm. The authenticity of cADPR produced was confirmed by coelution with standard compounds and the sea urchin egg bioassay, as described previously (22).
Detection of cADPR levels in myometrial tissue
One gram of human myometrium tissue was frozen in liquid N2, pulverized into a powder, and extracted with 5% trichloroacetic acid (TCA) at 4 C. TCA was removed with water-saturated ether as described previously (17). The aqueous layer containing the cADPR was removed and adjusted to pH 8 with 20 mM sodium phosphate. To remove nucleotides other than cADPR, a mixture containing hydrolytic enzymes was added to the samples, with the following final concentrations: 0.44 U/ml nucleotide pyrophosphatase, 12.5 U/ml alkaline phosphatase, 0.0625 U/ml NADase, 2.5 mM MgCl2, and 20 mM sodium phosphate (pH 8.0). Incubation proceeded overnight at 37 C. The enzyme mixture hydrolyzes all nucleotides (including NAD+) in the samples, except for cADPR, which is resistant to these enzymes (17). Enzymes were removed by filtration with Centricon-3 filters, and samples were recovered in the filtrate after centrifugation at 4 C and 3000 x g for 30 min (17). The detection of cADPR was performed with some modifications to the cycling method described recently (22). In brief, 0.1 ml cADPR standard or nucleotides extracted from human myometrium samples were incubated with 50 µl cycling reagent containing 0.3 µg/ml ADP-ribosyl cyclase, 30 mM nicotinamide, 100 mM sodium phosphate (pH 8), 2% ethanol, 100 µg/ml alcohol dehydrogenase, 20 µM resazurin, 10 µg/ml diaphorase, 10 µM flavin mononucleotide, and 0.5 mg/ml BSA. Increase in the resorufin fluorescence (with excitation at 544 nm and emission at 590 nm) was measured using a Hitachi F-2000 fluorometer. The results shown are means ± SE from at least three independent measurements. The recovery rate of exogenous cADPR detected by this method, after TCA extraction of standard concentrations of cADPR, was in the range of 75–80%.
Immunoprecipitation and Western blot
Human myometrial cells and myometrium extracts were incubated in lysis buffer containing 0.05% IGEPAL-CA 630, 20 mM EDTA, 20 mM NaCl, 20 mM Tris (pH 7.0), 10% (vol/vol) protein G Sepharose, and a 1:100 dilution of mouse monoclonal antibody against human CD38 (SC 7325; Santa Cruz Biotechnology, Santa Cruz, CA) for 4 h at 4 C. After 7.5% SDS-PAGE, protein was electroblotted onto polyvinyl difluoride membrane, which was further blocked with 5% nonfat milk overnight and probed with 1:100 dilution of goat polyclonal antibody against human CD38 (catalog no. SC 7048; Santa Cruz Biotechnology) for 4 h. The immunoreactive bands were detected using a 1:20,000 dilution of horseradish peroxidase-conjugated antigoat IgG (SC 2020; Santa Cruz Biotechnology) as secondary antibody and an enhanced chemiluminescence detection system.
Measurement of CD38 mRNA expression and RT-PCR
mRNA was isolated using an Invitrogen kit (FastTrack 2.0 kit, K1593–03) according to the manufacturer’s instructions. cDNA was synthesized at 42 C for 50 min using 0.05 µg oligo (dt) and 50 U SuperScript II reverse transcriptase (11904-018; Life Technologies, Inc., Grand Island, NY). For PCR, a 2 µl-aliquot of each cDNA solution was added to a reaction medium containing 0.2 mM deoxynucleotide triphosphate, 50 mM MgCl2, 0.25 U Taq polymerase, and 0.2 µM CD38 primers. Reactions were performed in a DNA thermal cycler, with 40 cycles at 94 C for 45 sec, at 55 C for 60 sec, and at 72 C for 90 sec. CD38 primers were: 5'-ACCCCGCCTGGAGCCCTATG-3' and 5'-GCTAAAACAACCACAGCGACTGG-3'. As housekeeping mRNA, glyceraldehyde-3-phosphate dehydrogenase (GAPDH) primers were used under the same conditions as described for CD38.
Flow cytometric analysis and immunocytochemistry of myometrial CD38
Flow cytometric analysis was performed by incubating intact myometrial cells with fluorescein isothiocyanate (FITC)-labeled monoclonal antibody against CD38 (SC7325-FITC; Santa Cruz Biotechnology) in a 1:100 dilution and further analyzed in a FACScan fluorescence cytometer (Becton Dickinson, Lincoln Park, NJ). For immunocytochemistry, cells plated on coverslips were permeabilized with 0.1% triton X-100 for 30 min and stained for either ryanodine receptor-channel, using a BODIPY-TR-ryanodine (B13802; Molecular Probes) and an antihuman CD38 monoclonal antibody FITC-labeled (SC7325-FITC; Santa Cruz Biotechnology).
Force generation in human uterine muscle strips
In summary, uterine smooth muscle strips were dissected, 0.3- to 0.5-mm-wide strips of human myometrium mounted in a Güth muscle research system (Scientific Instruments, Heidelberg, Germany), and samples were mounted in a quartz tissue cuvette between length and force transducers by stainless steel microforceps. Signals were recorded via a data acquisition board (AT-MIO-16-L9; National Instruments Corp., Austin, TX) and software (Labview; National Instruments) running on a personal computer. The strips were perfused initially at 1 ml/min with physiological saline solution aerated with 95% O2-5% CO2. After stabilization and testing with oxytocin, the strips were incubated with 5 mM nicotinamide for 1 h before further addition of 1 µM oxytocin. Each strip was its own control.
Materials
The human myometrium used in all experiments was obtained from healthy premenopausal women undergoing elective hysterectomy. Lytechinus pictus and Aplysia californica were obtained from Marinus, Inc. (Long Beach, CA). BODIPY-TR-ryanodine and Fluo-3 were purchased from Molecular Probes. All other reagents (of the highest available purity grade) were supplied by Sigma Chemical Co. (St. Louis, MO).
The reported experiments were repeated at least three to six times, as appropriate. Data are expressed as means ± SD. The paired t test was used to evaluate statistical significance; P < 0.05 was considered significant.
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Results and Discussion |
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, human myometrial cells can be labeled with bodipy ryanodine, and its binding can be blocked by excess nonlabeled ryanodine (data not shown). Perinuclear distribution of bodipy ryanodine is consistent with the location of the RyR in the sarcoplasmic reticulum of the myometrial cells. Ryanodine binding sites are also present in human myometrial microsomes. Binding of radioactive ryanodine to microsomes was determined and can be blocked by cold ryanodine, cADPR, and ruthenium red (Fig. 1B). In addition, the effect of cADPR on ryanodine binding can be blocked by the cADPR inhibitor 8-Br-cADPR (Fig. 1B). These experiments confirmed the presence of RyR in myometrial cells (8, 9, 10, 11, 23) and indicate that cADPR interacts with the RyR in human myometrial cells and may play a role as the endogenous regulator of the RyR.
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). Furthermore, 8-Br-cADPR does not inhibit the IP3-induced Ca2+ release in human myometrial microsomes (Fig. 1C).
Role of the cADPR-signaling pathway in oxytocin-induced Ca2+ release in human myometrial cells
Agonists such as oxytocin play critical roles in the coordination of uterine contraction during labor. IP3-triggered [Ca2+]i transients have been shown to play roles in oxytocin-induced myometrial contraction (1, 3, 4, 5, 6); however, inhibition of phospholipase C does not completely abolish the [Ca2+]i transient induced by oxytocin (3, 4, 5, 6). These results strongly suggest that another intracellular signaling pathway is involved in the regulation of uterine [Ca2+]i homeostasis in response to agonists. In fact, it has been previously shown that the RyR channel is important for the propagation of oxytocin-induced Ca2+ waves in cultured myometrium (8, 9, 10, 11). Here, we also found that the RyR is important for the oxytocin-induced Ca2+ transients. The RyR inhibitors dantrolene and ruthenium red (at concentrations of 30 µM) can impair the oxytocin-induced Ca2+ transient by about 43 and 50%, respectively.
The intracellular signaling pathway that regulates the RyR channel in response to uterotonic agonists in human myometrium is not yet known. Because cADPR has been implicated as a second messenger, responsible for the regulation of RyR channel in several mammalian cells (12, 13, 14, 15), we explored the role of this nucleotide in oxytocin-induced [Ca2+]i increase. Single-cell confocal imaging showed that the oxytocin-induced Ca2+ transient is dependent on the intact extracellular Ca2+ influx pathway (Fig. 2A). In addition, oxytocin-induced Ca2+ transients were also dependent on the Ca2+ release from internal stores. The use of inhibitors of both the cADPR and IP3 systems indicates that oxytocin-induced Ca2+ release appears to be dependent on both intracellular signaling systems (Fig. 2, B and C). A complex coordination of extracellular Ca2+ and intracellular messengers appears to be fundamental for the oxytocin-induced Ca2+ transients in human myometrial cells. Similar interactions between Ca2+ pools have been described previously (19, 20, 24). We postulated that the cADPR system regulates the Ca2+-induced Ca2+ release mediated by the RyR and augments the intracellular Ca2+ transient possibly initiated by extracellular Ca2+ influx or activation of the IP3-receptor.
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augments oxytocin-induced Ca2+ transients via increase of intracellular levels of cADPR induces cellular expression of CD38 and increases synthesis of cADPR (25). Furthermore, it has been proposed that increased intracellular levels of cADPR in TNF-treated cells could increase agonist-stimulated Ca2+ release (25, 26). In fact, TNF-treated mesangial and airway smooth muscle cells experience an increase in agonist-stimulated Ca2+ transient that is likely due to intracellular accumulation of cADPR (25, 26). In the present study, the use of TNF-stimulated human myometrial cells further demonstrated the influence of the cADPR system on oxytocin-stimulated intracellular Ca2+ transients.
As shown in Fig. 3, A and B, treatment of cultured human myometrial cells with TNF led to an increase in CD38 cyclase activity of about 500%. Time dependence of the TNF stimulation of cyclase indicates that its effect is mediated byincreased expression of the enzyme (Fig. 3B). Furthermore, TNF was more potent then other cytokines tested (Fig. 3C). Treatment of cells with TNF for 24 h led to a 450% increase in intracellular levels of cADPR (Fig. 3D).
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, the effect of TNF on CD38 cyclase activity is dependent on a genomic mechanism that leads to increased expression of the enzyme. First, the effect of TNF on CD38 cyclase activity was inhibited by the DNA synthesis inhibitor actinomycin D and cycloheximide, an inhibitor of protein synthesis (Fig. 4A). Furthermore, we observed an increased expression of immunoreactive CD38 in cells treated with TNF. Immunoprecipitation and Western blot analysis showed a 3-fold increase of CD38 enzyme in TNF-treated cells when compared with nontreated cells (Fig. 4B, lanes 3 and 4, and Fig. 4C). Cycloheximide and actinomycin D inhibited the effect of TNF on the activity of the ADP-ribosyl cyclase (Fig. 4C). These data were confirmed by immunostaining the cultured human myometrial cells with FITC-labeled monoclonal antibody for CD38 (Fig. 4, D and E). Furthermore, flow cytometric analysis of intact myometrial cells showed a 5-fold increase in CD38-positive cells treated with 50 ng/ml TNF (data not shown).
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-treated human myometrial cells (Fig. 4F); however, the increase in CD38 mRNA expression by TNF was abolished by actinomycin D (data not shown). In contrast, TNF had no effect on the mRNA levels of the constitutive enzyme GAPDH. These data clearly indicate that TNF treatment leads to increased expression of CD38 and increased intracellular cADPR levels in human myometrial cells. Additional experiments revealed that in TNF-treated cells, oxytocin-induced Ca2+ transient was approximately 1.5 times greater than normal (Fig. 5). The cADPR inhibitor 8-Br-cADPR blocked this increase (Fig. 5, inset).
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-treated and control cells can be severely impaired by treating the cells with nicotinamide (Fig. 6, A and B). These data further confirm the role of the cADPR system in oxytocin-induced Ca2+ transients in human myometrial cells. We also observed that treatment of human myometrial strips with nicotinamide led to a 30–50% inhibition of oxytocin-induced contraction in vitro (Fig. 6C).
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We also demonstrated that TNF and interleukins can affect ADP-ribosyl cyclase activity and CD38 expression in human myometrial cells, thus increasing levels of cADPR. TNF-induced alterations in cADPR increase oxytocin-induced [Ca2+]i transient and may play important roles in the process that culminates in successful delivery.
During human pregnancy the myometrium is relatively quiescent until close to term (1, 28, 29). The precise sequence of events preceding the initiation of intense contractile activity in human labor is still unknown and is likely due to an orchestration of many pathways. It is possible that cytokines may modulate the expression of components of the cADPR pathway in vivo and, in this way, increase contractility.
During pregnancy, the myometrium undergoes various structural and biochemical changes (1, 28, 29). During the first two stages of pregnancy, hyperplasia and hypertrophy occur. It is during the third stage of pregnancy, however, that the balance in the myometrium starts to shift from relaxant to contracting pathways. Mounting evidence indicates that the parturition process represents an inflammatory response (30, 31, 32, 33). When term labor approaches, leukocytes, macrophages, neutrophils, and T-lymphocytes infiltrate the myometrium. Moreover, it has been shown that proinflammatory cytokines are expressed in myometrium in association with labor (30, 32). Recent studies have demonstrated that the onset of term labor induces elevated production of IL-1ß, IL-6, IL-8, and TNF by placental endothelial cells and increases their concentrations in the amniotic fluid (31).
In any case, our findings clearly indicate a role for cADPR in oxytocin-induced Ca2+ transients in human myometrial cells. Development of new therapeutic strategies that target the cADPR system may be key to the generation of new specific therapies for dysfunctional uterine contractions.
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Footnotes |
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Abbreviations: (Ca2+)i, Intracellular Ca2+; cADPR, cyclic ADP-ribose; FITC, fluorescein isothiocyanate; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; HBSS, Hanks’ balanced salt solution; IP3, inositol 1,4,5-trisphosphate; NGD, nicotinamide guanine dinucleotide; RyR, ryanodine receptor; SmBM, smooth muscle cell basal medium; TCA, trichloroacetic acid.
Received June 25, 2003.
Accepted for publication October 8, 2003.
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TopAbstractIntroductionMaterials and MethodsResults and DiscussionReferences |
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