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1 Department of Physiology and Neuroscience
2 Department of Cell Biology
3 Department of Cardiology, New York University School of Medicine, New York, NY 10016, USA
4 Departments of Cell Biology and of Biochemistry, University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd, Dallas, TX 75390-9039, USA
5 Department of Medicine, Mc1027, amb m172, University of Chicago, 5841 South Maryland Avenue, Chicago, IL 60637
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Abstract |
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(Received 29 March 2005;
accepted after revision 26 April 2005;
first published online 28 April 2005)
Corresponding author G. G. Holz: Medical Sciences Building Room 442, 550 First Avenue, New York, NY 10016, USA. Email: holzg01{at}popmail.med.nyu.edu
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Introduction |
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Coincidence detection in biological systems is a phenomenon in which two or more complementary signals interact synergistically to generate a cellular response. Neither signal is an adequate stimulus in the absence of its complement. The type VIII isoform of adenylyl cyclase expressed in ß cells acts as a molecular coincidence detector because it is stimulated not only by GS GTP-binding proteins, but also by Ca2+/calmodulin (Pipeleers et al. 1985; Schuit & Pipeleers, 1985; Delmeire et al. 2003). Coincidence detection also exists when the activity of an effector molecule is governed by multiple second messengers. This may be the case for intracellular Ca2+ release channels (ryanodine receptors, RYR; inositol 1,4,5-trisphosphate (IP3) receptors, IP3-R), the opening of which is reported to be facilitated by Ca2+ and cAMP (Marx et al. 2000; Bruce et al. 2003). Because GLP-1 stimulates cAMP production, and because ß cell glucose metabolism stimulates influx of Ca2+ through voltage-dependent Ca2+ channels (VDCCs), it is predicted that GLP-1 and glucose should interact synergistically to gate Ca2+ release channels from a closed to open state. Indeed, second messenger coincidence detection of this type might explain the unusual interaction of GLP-1 and glucose to mobilize an intracellular source of Ca2+ in the ß cell (Gromada et al. 1995; Bode et al. 1999; Holz et al. 1999; Kang et al. 2001, 2003; Kang & Holz, 2003; Sasaki et al. 2002; Tsuboi et al. 2003; Dyachok & Gylfe, 2004).
Here we demonstrate that Ex-4 acts via cAMP, protein kinase A (PKA), and the Epac family of cAMP-regulated guanine nucleotide exchange factors (cAMPGEFs; also known as Epac1 and Epac2; Holz, 2004a) to sensitize Ca2+-induced Ca2+ release (CICR) mediated by the RYR and IP3-R. Sensitization allows CICR to be triggered by the uncaging of Ca2+ in INS-1 cells or mouse ß cells loaded with a photolabile Ca2+ chelator (NP-EGTA; Ellis-Davies et al. 1994). Because the uncaging of Ca2+ fails to stimulate CICR in the absence of cAMP-elevating agents, it is concluded that there exists in ß cells a process of second messenger coincidence detection whereby intracellular Ca2+ release channels monitor a simultaneous increase of cAMP and Ca2+ concentrations. Some of these findings have been published in preliminary form (Kang et al. 2005).
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Methods |
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Islets were isolated from male C57BL/6 mice fed ad libitum (20–25 g body weight; Charles River Laboratories, Inc., Wilmington, MA, USA). The mice were anaesthetized by inhalation of CO2 (100%; 2–3 min exposure), and were killed by cervical dislocation. Surgical procedures for removal of the pancreas were performed in accordance with NYU School of Medicine policies governing the ethical use of mice for experimentation (IACUC Protocol no. 040602-01). After digestion of the pancreas with collagenase P (Roche Applied Science, Indianapolis, IN, USA; 2 mg ml–1 dissolved in RPMI 1640 medium), batches of 150–200 islets were subjected to mild trypsinization and were dispersed by trituration in a Ca2+-free saline in order to generate a single cell suspension. Isolated cells were then allowed to adhere to glass coverslips (25CIR-1; Fisher Sci.) coated with concanavalin A (type V; Sigma-Aldrich, St Louis, MO, USA). Primary cultures were maintained in a humidified incubator (95% air, 5% CO2) at 37°C in RPMI 1640 supplemented with 10% FBS, 100 units ml–1 penicillin G, and 100 µg ml–1 streptomycin. ß cells were identified on the basis of their large diameter and granular appearance. INS-1 cells (passage numbers 70–90) were maintained in RPMI 1640 containing 10 mM Hepes, 11.1 mM glucose, 10% FBS, 100 units ml–1 penicillin G, 100 µg ml–1 streptomycin, 2.0 mM L-glutamine, 1.0 mM sodium pyruvate, and 50 µM 2-mercaptoethanol (Asfari et al. 1992). INS-1 cells were passaged by trypsinization and subcultured once a week. All reagents for cell culture were obtained from Invitrogen-Life Technologies (Rockville, MD).
Measurement of [Ca2+]i
The fura–2 loading solution consisted of standard extracellular saline (SES) containing (mM): 138 NaCl, 5.6 KCl, 2.6 CaCl2, 1.2 MgCl2, 10 Hepes, 11.1 D–glucose and supplemented with 1 µM fura–2 AM (Molecular Probes Inc., Eugene, OR, USA), 2% FBS, and 0.02% Pluronic F-127 (w/v; Molecular Probes Inc.). Cells were exposed to the fura–2 loading solution for 20–30 min at 22°C. Experiments were performed in SES at 32°C using a TE300 inverted microscope (Nikon, Melville, NY, USA) equipped with a temperature-controlled stage (Medical Systems Corp., Greenvale, NY, USA) and a 100x Nikon UVF oil immersion objective (NA 1.3). Microfluorimetry was performed ratiometrically at 0.5 s intervals using a video imaging system outfitted with an intensified CCD camera (IonOptix Corp., Milton, MA, USA). A rotating mirror delivered excitation light at 340 or 380 nm. The emitted light was measured at 510 nm, and the average of 29 frames of imaging data was used to calculate numerator and denominator values for determination of 340/380 ratios after background subtraction. [Ca2+]i was calculated according to established methods (Grynkiewicz et al. 1985). Calibration of raw fura–2 fluorescence values was performed as described (Kang & Holz, 2003) using fura–2 [K+]5 salt dissolved in calibration buffers from Molecular Probes Inc. (Calcium Calibration Kit 1 with Mg2+). Values of Rmin and Rmax were 0.20 and 7.70. Components of the imaging system are illustrated (Supplementary Fig. 1).
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Cells were bathed for 60 min at 23°C in SES containing cell-permeable caged Ca2+ (NP-EGTA-AM, 5 µM, Molecular Probes, Inc.) or caged IP3 (ci-IP3/PM, 10 µM). The loading solution also contained 1 µM fura-2-AM, 2% FBS, and 0.02% Pluronic F-127. ci-IP3/PM is a photolabile caged IP3 related in structure to cm-IP3/PM (Li et al. 1998). It was synthesized in nine steps from myo-inositol in the laboratory of W-H Li, using a procedure similar to that used for making cm-IP3/PM (W-H Li, manuscript in preparation). In ci-IP3/PM, 2- and 3-hydroxyl groups of myo-inositol are protected by an isopropylidene group; whereas in cm-IP3/PM, 2- and 3-hydroxyl groups are protected by a methoxymethylene group. Like cm-IP3/PM, ci-IP3/PM is cell permeable, and once inside cells, induces Ca2+ release from intracellular stores upon UV flash photolysis (W-H Li, unpublished results; Wagner et al. 2004). Uncaging of caged compounds was achieved using a flash photolysis system (Model JML-C2, Rapp OptoElectronic, Hamburg, Germany). The excitation light of 80 J intensity and 600 µs duration was filtered using a short-pass filter (cut-off 390 nm), and was delivered to the specimen by way of the microscope's objective (Supplementary Fig. 1). The intensity and duration of the flash were minimized so that no measurable photo-bleaching of fura–2 was observed. The intensity of 340 and 380 nm excitation light for detection of fura–2 was also reduced so as to be so low as to produce no measurable uncaging of caged compounds.
Measurement of endoplasmic reticulum [Ca2+] using YC3.3-er
INS-1 cells were transiently transfected with a plasmid encoding yellow cameleon 3.3-er (YC3.3-er) (Griesbeck et al. 2001) using Lipofectamine 2000 (Invitrogen-Life Tech.). Cells on glass coverslips were placed within a microperfusion chamber mounted on an inverted microscope (TE-2000, Nikon) and were visualized with a 40x objective (Hara et al. 2004). The excitation light (440 nm) was attenuated 50–90% using neutral density filters. Changes in fluorescence emission intensities at 535 nm (citrine; FRET acceptor) and 485 nm (enhanced cyan fluorescent protein, ECFP; FRET donor) were monitored using emission filters mounted on a computer-controlled filter wheel (Lambda 10–2 Optical Filter Changer, Sutter Instruments, Novato, CA, USA). Images (50–100 ms exposure time) were captured with a 16-bit Cascade 650 digital camera (Photometrics, Tucson, AZ, USA) at 2–10 s intervals, and were analysed using MetaMorph/MetaFluor software (Universal Imaging Corp., Downington, PA, USA). Data were expressed as the background subtracted ratios of the FRET acceptor and FRET donor emission intensities monitored at 535 and 485 nm, respectively. Cells were superfused at 32°C in saline of high pH-buffering capacity consisting of (mM): 138 NaCl, 5.6 KCl, 2.6 CaCl2, 1.2 MgCl2, 25 NaHCO3, 10 Hepes-NaOH (pH 7.40), and 11.1 glucose (Varadi & Rutter, 2004).
Confocal microscopy for measurement of [Ca2+]i
Cells plated on 0.15 mm glass coverslips were incubated for 20 min at room temperature in SES containing 5 µM fluo-4 AM (Molecular Probes). The [Ca2+]i was imaged using a laser scanning confocal microscope (Leica DM IRE2; Leica Microsystems Heidelberg GmbH) equipped with a 63x water immersion objective (NA 1.2). Fluo-4 was excited at 488 nm using an argon laser. The emitted light passed through a 500 nm dichroic filter for detection using a fluo-4 emission filter (Leica TCS SP2; Leica Microsystems Heidelberg GmbH). Images of the x–y optical sections were recorded with a resolution of 512 pixels line–1 at 400 Hz. Raster point size was 0.1 µm, with an overall lateral resolution of 0.16 µm. For each data set, 20 x–y scans of a small cluster of cells were acquired. After the third section was scanned, a test solution was applied via a micropipette. Image analysis was performed using Leica Confocal Software (Version 2.5; Leica Microsystems).
Microinjection of heparin
Low molecular weight heparin (MW 5000 kDa; Calbiochem) was dissolved in buffer containing (mM): 110 KCl, 10 NaCl, 2 MgCl2, 20 Hepes, 5 KH2PO (297 mOsmol, pH 7.2). The heparin was injected from Femtotip II needles into individual ß cells using a Transjector 5246 microinjection system (Eppendorf, Hamburg, Germany) mounted on a Nikon TE200 inverted microscope. The injection pressure was 100 hPa, the compensation pressure was 33 hPa, and the duration of injection was 0.2 s. In order to identify ß cells injected with heparin, the injection solution also contained fluorescein (1 mg ml–1; Sigma-Aldrich). The injected cells were then loaded with fura 2 AM and caged compounds. Prior to imaging of Ca2+, cells injected with heparin and fluorescein were visualized using an EYFP filter set.
Epac constructs and transfection protocol
Human wild-type Epac1 (GenBank Accession No. AAF103905) (de Rooij et al. 1998) and dominant negative Epac1 (R279E) in pcDNA3.1 were obtained from Dr X. Cheng (Galveston, TX) (Qiao et al. 2002). Mouse wild-type Epac2 (cAMPGEFII; Accession No. AB021132) and dominant negative Epac2 (G114E, G422D) in pSR
were provided by Dr S. Seino (Kobe, Japan) (Ozaki et al. 2000; Kashima et al. 2001). Epac1 and Epac2 cDNAs were subcloned into pCMV2-FLAG (Sigma-Aldrich, USA) to insert the FLAG epitope at the exchange factor's N-terminus. Epac constructs were introduced into INS-1 cells using LipofectAMINE Plus (Invitrogen-Life Tech.). Cells transfected with wild-type or dominant negative Epac were identified by cotransfection with pEYFP-N1 (Clonetech, Palo Alto, CA, USA). A 1: 4 molar ratio of pEYFP-N1 relative to Epac plasmid was used in all transfections (Kang et al. 2001). EYFP fluorescence was monitored in fura-2-loaded cells 2 days post-transfection, using 513 nm excitation and 527 nm emission filters. Once an EYFP-positive cell was identified as having been transfected, the filter set was manually switched to a fura-2 filter set, allowing ratiometric determinations of [Ca2+]i. Control experiments demonstrated that less than 1% crossover existed between fura-2 and EYFP when using filter sets selective for each reporter (see Supplementary Figs 2–4).
Detection of recombinant Epac immunoreactivity
Whole cell lysates of transfected INS-1 cells expressing recombinant Epac were dissolved in 1x Laemmli's sample buffer, boiled for 5 min, centrifuged to remove unsolubilized material, and resolved by SDS-PAGE using 4% stacking and 12% resolving gels. The resolved Epac proteins were transferred to Immobilon-P PVDF membrane (Millipore, Bedford, MA, USA) by electrophoretic transfer (120 V, 1 h). Western immunoblot analyses were performed using mouse anti-FLAG monoclonal primary antiserum (Sigma-Aldrich; 1: 1000 dilution) in combination with goat antimouse polyclonal secondary antiserum (1: 5000 dilution) conjugated to horseradish peroxidase (Sigma-Aldrich).
CRE-Luc reporter assay
Luciferase activity was measured in lysates of INS-1 cells transfected with CRE-Luc (Stratagene, La Jolla, CA, USA) as described (Chepurny et al. 2002, Chepurny & Holz, 2002). This construct allows expression of luciferase to be regulated by cAMP response elements (CREs) located within a minimal promoter. Monolayers of INS–1 cells were exposed to test substances for 4 h, lysed and assayed for luciferase-catalysed photoemissions using a luciferase assay kit (Promega, Madison, WI, USA) and a luminometer allowing automated application of solutions containing ATP and luciferin (Model TR-717, Perkin Elmer Applied Biosystems, Foster City, CA, USA). Experiments were carried out in triplicate. Statistical analyses were performed using an ANOVA test combined with Fisher's PLSD test.
Sources of reagents and application of test substances
Exendin-4, forskolin, caffeine, ryanodine, and thapsigargin were from Sigma-Aldrich. H-89 was from Calbiochem (San Diego, CA, USA). 8-pCPT-cAMP, 6-Bnz-cAMP, and 8-pCPT-2'-O-Me-cAMP were from BioLog Life Science (Bremen, Germany). Test solutions dissolved in SES were added to the bath solution or were applied to individual cells from glass ‘puffer’ micropipettes (type 1B150-6; World Precision Institute Inc., Sarasota, FL, USA) using a pressure ejection system (PicoSpritzer II, General Valve Corp., NJ, USA) as described (Holz et al. 1993).
Statistical analyses of CICR
Population studies were performed at the single-cell level in order to determine the percentage of cells exhibiting CICR under conditions in which cells were treated with pharmacological agents added directly to the bath and ‘puffer’ pipette solutions. A coverslip with adherent cells served as a control, while a ‘sister’ culture served as the test. At least 10 cells (coverslip)–1 were selected one at a time in random order to determine basal [Ca2+]i and CICR amplitude. CICR in INS-1 cells was defined as a transient increase of [Ca2+]i, the duration of which did not exceed 30 s when measured at the 10% amplitude cut-off (Fig. 1C). An additional requirement was that the increase of [Ca2+]i for INS-1 cells must have exceeded 200 nM when measured at the 50% amplitude cut-off (Fig. 1C). Because CICR in mouse ß cells tended to be of smaller amplitude, these criteria were modified so that the increase of [Ca2+]i must have exceeded 125 nM when measured at the 50% amplitude cut-off. Each experiment was performed in triplicate, and statistical analyses were performed using the ANOVA test combined with Fisher's PLSD test.
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Results |
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To assess the properties of CICR in INS-1 cells, we performed UV flash photolysis to uncage Ca2+ from NP-EGTA, a photolabile Ca2+ chelator. Prior to the loading of cells with NP-EGTA, the resting [Ca2+]i was 135 ± 65 nM (mean ± S.D.; n = 20 cells). After loading with NP-EGTA, there was a statistically significant reduction of [Ca2+]i to 91 ± 5 nM (P < 0.001; n = 20 cells). A 600 µsec flash of UV light produced a small increase of [Ca2+]i in these cells loaded with NP-EGTA (122 ± 26 nM increase; n = 30 cells) (Fig. 1A). This increase of [Ca2+]i recovered to its original baseline and was fully repeatable (Fig. 1A). No such increase of [Ca2+]i was measured in cells not loaded with NP-EGTA (n = 20 cells; data not shown). It may be concluded that the increase of [Ca2+]i depicted in Fig. 1A resulted from the uncaging of Ca2+.
When the GLP-1-R agonist exendin-4 (Ex-4, 1 nM) was applied to single INS-1 cells loaded with NP-EGTA, no increase of [Ca2+]i was measured (Fig. 1B). In contrast, a large and transient increase of [Ca2+]i (CICR) was observed when Ex-4 was applied simultaneous with the uncaging of Ca2+ (Fig. 1B). As summarized in Table 1A, this sensitizing action of Ex-4 to promote CICR was observed in 17 of 24 cells tested. The mean amplitude of the Ca2+ spike measured under these conditions was 1110 ± 176 nM, and the mean duration measured at the 50% amplitude cut-off was 5.1 ± 1.3 s (n = 15 cells). Sensitization of CICR was also observed when UV flash photolysis was performed in the presence of cAMP-elevating agent forskolin (2 µM) (Fig. 1D). This action of forskolin was repeatable and was observed in 23 of 30 cells tested.
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cAMP and Ca2+ may interact to generate CICR in INS-1 cells. To evaluate this possibility, we examined whether cell-permeant cAMP analogues sensitize the CICR mechanism of INS-1 cells in a manner analogous to that described for Ex-4 and forskolin. When NP-EGTA-loaded INS-1 cells were exposed to 8-pCPT-cAMP (100 µM), an activator of both PKA and Epac, the uncaging of Ca2+-triggered CICR (Fig. 2A, Table 1A). This action of 8-pCPT-cAMP was reduced but not blocked by treatment with H-89 (10 µM), a PKA inhibitor (Fig. 2A; inset). CICR was also observed when cells were exposed to 8-pCPT-2'-O-Me-cAMP (100 µM), a cAMP analogue active at Epac only (Fig. 2B), or 6-Bnz-cAMP (100 µM), an analogue active at PKA only (Fig. 2C). As predicted, 8-pCPT-2'-O-Me-cAMP remained effective in cells treated with H-89, whereas the action of 6-Bnz-cAMP was nearly abrogated (cf. Fig. 2B and C; insets). Given that INS-1 cells express mRNA corresponding to Epac1 and Epac2 (Leech et al. 2000), such findings are expected if the sensitizing action of cAMP is mediated not only by PKA but also by Epac. To confirm the efficacy of H-89 as an inhibitor of PKA, the activity of a PKA-regulated luciferase reporter (CRE-Luc) was assessed in transfected INS-1 cells. Under conditions of 8-pCPT-cAMP treatment, both the basal and stimulated activities of CRE-Luc were inhibited by H-89 (Fig. 2D).
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We next sought to ascertain if it is PKA or Epac that mediates the cAMP-dependent mobilization of Ca2+ by Ex-4. To this end, the action of Ex-4 was evaluated after treatment of INS-1 cells with H-89 or after transfection with dominant negative isoforms of Epac that do not bind cAMP (Holz, 2004a). Population studies demonstrated that H-89 exerted partial inhibitory effects when evaluating its ability to suppress CICR. Whereas a low concentration of H-89 (1 µM) was without effect, a higher concentration (10 µM) reduced the percentage of cells responding to Ex-4 by 60% (Fig. 3A). The ineffectiveness of 1 µM H-89 in this assay of CICR is notable, because this concentration of H-89 reduced CRE-Luc activity by 33% (Fig. 2D). Therefore, there may exist a PKA-independent signalling pathway by which Ex-4 exerts its sensitizing action. This is likely to be the case because the action of Ex-4 was inhibited by overexpression (see immunoblot, Fig. 3B) of dominant negative (DN) FLAG epitope-tagged Epac1 (Fig. 3C) or Epac2 (Fig. 3D). These DN Epacs incorporate inactivating amino acid substitutions within their cAMP-binding domains (Holz, 2004a). Importantly, overexpression of wild type (WT) FLAG-Epac1 or FLAG-Epac2 failed to confer such an inhibitory effect (Figs 3C and D).
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A 10–15 min pretreatment of INS-1 cells with ryanodine (10 µM) reduced the percentage of INS-1 cells exhibiting CICR under conditions of UV flash photolysis. This was the case when the uncaging of Ca2+ was performed in the presence of Ex-4 (1 nM) or 8-pCPT-cAMP (100 µM) (Fig. 4A and B; Table 1A). Caffeine (1 mM) mimicked the action of Ex-4 (Fig. 4C), and the action of caffeine was also inhibited by ryanodine (Fig. 4D; Table 1A). To validate that an intracellular source of Ca2+ was mobilized as a consequence of CICR, the actions of caffeine and forskolin were examined under conditions in which the SES was nominally Ca2+ free. Under such conditions, caffeine (1 mM) and forskolin (2 µM) allowed for the appearance of CICR (Fig. 5A and B). The source of Ca2+ mobilized by caffeine and forskolin included thapsigargin-sensitive Ca2+ stores, because the actions of both agents were abrogated by pretreatment with thapsigargin (1 µM) (Fig. 5C and D).
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A high concentration of caffeine (10 mM) produced a transient increase of [Ca2+]i in INS-1 cells not loaded with NP-EGTA. This action of caffeine was fast in onset and recovered to baseline within 10–20 s (Fig. 6A). The increase of [Ca2+]i was measured in the cytoplasm and also the nucleus (Fig. 6B). The source of Ca2+ mobilized included the endoplasmic reticulum (ER), as demonstrated using INS-1 cells expressing a cameleon Ca2+ reporter (YC3.3-er) targeted to the ER. When 10 mM caffeine was administered to these cells, a transient decrease of 535/485 nm emission ratio was detected (Fig. 6C). This signifies a decrease of ER calcium concentration ([Ca2+]ER) (Griesbeck et al. 2001). The [Ca2+]ER of INS-1 cells was also lowered when these cells were exposed to thapsigargin (Fig. 6D).
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To examine how cAMP-elevating agents influence CICR in mouse ß cells, UV flash photolysis was performed under conditions of NP-EGTA loading identical to that described for INS-1 cells. Prior to loading, the resting [Ca2+]i was 104 ± 29 nM (n = 25 cells). This value decreased to 87 ± 14 nM (n = 22 cells) in NP-EGTA-loaded cells (P < 0.001; n = 20 cells). The uncaging of Ca2+ in ß cells generated a small increase of [Ca2+]i but did not initiate CICR (Fig. 7A; 55 ± 11 nM increase; n = 17 cells). In these same cells, application of Ex-4 (1 nM), forskolin (2 µM), or the Epac-selective cAMP analogue 8-pCPT-2'-O-Me-cAMP (100 µM) failed to alter resting [Ca2+]i (Fig. 7A–C). However, all three cAMP-elevating agents allowed for the appearance of CICR under conditions of UV flash photolysis (Fig. 7A–C; Table 1B). 8-pCPT-2'-O-Me-cAMP (100 µM) was highly effective in this assay. It allowed for the appearance of CICR in 8 of 15 cells tested. Similar to findings obtained with INS-1 cells, no sensitization of CICR by forskolin was observed when mouse ß cells were treated with thapsigargin (Fig. 7D).
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To assess whether CICR sensitized by forskolin resulted from activation of RYR, the action of ryanodine was examined. A role for RYR is indicated because pretreatment with ryanodine (10 µM) rendered forskolin less effective in the assay of ß cell CICR reported here (Table 1B). Furthermore, caffeine (1 mM), a sensitizer of RYR, allowed for the appearance of CICR under conditions of UV flash photolysis. This action of caffeine was also diminished by ryanodine (Table 1B).
Because ryanodine-resistant CICR was sometimes observed (Table 1B), a sensitizing action of cAMP at the IP3-R may also exist in ß cells. Therefore, we assessed the efficacy of an IP3-R inhibitor (low molecular weight heparin, 100 mg ml–1; Ehrlich et al. 1994) administered intracellularly under conditions of NP-EGTA loading and UV flash photolysis. Administration of heparin led to a reduction in the percentage of forskolin-treated ß cells exhibiting CICR (Fig. 8A). Moreover, CICR was nearly abolished after combined treatment with heparin and ryanodine, thereby demonstrating that these two inhibitors of Ca2+ release channels acted in an additive manner (Fig. 8B).
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Discussion |
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Prior studies demonstrate that cAMP-elevating agents mobilize Ca2+ in ß cells and insulin-secreting cell lines (Islam et al. 1998; Holz et al. 1999; Kang et al. 2001, 2003; Kang & Holz, 2003; Dyachok & Gylfe, 2004; Dyachok et al. 2004). Ca2+ mobilized in this manner acts as a direct stimulus for exocytosis of insulin (Kang & Holz, 2003; Kang et al. 2003; Dyachok & Gylfe, 2004; reviewed by Holz, 2004b). Thus, exocytosis in ß cells is not simply dependent on influx of Ca2+ through VDCCs, but is also dependent on the interaction of Ca2+ and cAMP to promote CICR. In this regard, new studies provide evidence for the existence of a highly Ca2+-sensitive pool (HCSP) of secretory granules in ß cells (Wan et al. 2004; Yang & Gillis, 2004). These granules are released under conditions in which the [Ca2+]i increases to low micromolar levels. Since the mechanism of cAMP-dependent Ca2+ mobilization reported here generates an increase of [Ca2+]i in the 0.5–2 µM range, it may serve as an adequate stimulus for exocytosis of the HCSP.
Second messenger coincidence detection in ß cells
Here we describe an experimental strategy by which the existence of cAMP-regulated CICR in ß cells may be validated. The strategy relies on the use of caged Ca2+ and UV flash photolysis to produce a small increase of [Ca2+]i. Under these conditions, the uncaging of Ca2+ triggers CICR when Ca2+ release channels are sensitized by cAMP-elevating agents. Of particular note is our demonstration that the uncaging of Ca2+ fails to generate CICR in the absence of cAMP-elevating agents. This key observation leads us to conclude that a simultaneous increase of intracellular Ca2+ and cAMP concentrations is a necessary prerequisite for the initiation of CICR in ß cells. Fundamentally, this is a process of second messenger coincidence detection, and it results from dual stimulatory actions of Ca2+ and cAMP at Ca2+ release channels. These Ca2+ release channels correspond to RYR and the IP3-R, as demonstrated by antagonism of CICR following application of ryanodine or heparin.
IP3-R-mediated Ca2+ mobilization is also demonstrated through our use of ci-IP3/PM, a membrane-permeant caged IP3 that activates the IP3-R in a selective manner. Under conditions in which ß cells are treated with forskolin, the uncaging of IP3 stimulates CICR. This action of forskolin might reflect cAMP-dependent sensitization of the IP3-R. Alternatively, cAMP may sensitize RYR to stimulatory effects of Ca2+ released from activated IP3 receptors. If so, heparin would act at the IP3-R to prevent the small rise of [Ca2+]i that acts as an initiator of CICR mediated by RYR. Indeed, the mobilization of Ca2+ by IP3 in pancreatic acinar cells is reported to trigger CICR mediated by ryanodine receptors (Straub et al. 2000; Ashby et al. 2002). Thus, it will be of interest to ascertain whether there also exist functional interactions between IP3 receptors and RYR in ß cells.
Coincidence detection may explain context specificity
Prior studies of ß cells demonstrate that CICR occurs in a context-specific manner (Lemmens et al. 2001; Bruton et al. 2003). CICR is only observed when ß cells are equilibrated in elevated concentrations of extracellular glucose. Metabolism of glucose by ß cells promotes the filling of ER Ca2+ stores (Maechler et al. 1999; Tengholm et al. 1999), and it also produces an increase of [Ca2+]i (Henquin, 2000). Therefore, under conditions in which ß cells are exposed to glucose, two necessary conditions are met. ER Ca2+ stores are full and the cytosolic Ca2+ concentration is elevated sufficiently to allow CICR to be generated when Ca2+ release channels are sensitized by cAMP. Context specificity of this type is likely to be of physiological significance, because it may explain, at least in part, why the insulin secretagogue action of GLP-1 is strictly dependent on exposure of ß cells to glucose (Kieffer & Habener, 1999).
RYR as a determinant of ß cell function
Because insulin-secreting cells express RYR, albeit at low levels (Takasawa et al. 1998; Gamberucci et al. 1999; Holz et al. 1999; Islam, 2002; Lee et al. 2002; Mitchell et al. 2003; Beauvois et al. 2004; Johnson et al. 2004a,b), it is not surprising that caffeine, a sensitizer of RYR, recapitulates the Ca2+-mobilizing action of Ex-4 reported here. This action of caffeine is inhibited by ryanodine, thereby confirming the existence of CICR mediated by RYR. Such findings are consistent with prior studies demonstrating ryanodine and caffeine-sensitive mobilization of Ca2+ in ß cells or ß cell lines (Islam et al. 1992, 1998; Gromada et al. 1995; Gamberucci et al. 1999; Maechler et al. 1999; Holz et al. 1999; Kang et al. 2001, 2003; Kang & Holz, 2003; Mitchell et al. 2003; Sasaki et al. 2002; Varadi & Rutter, 2002; Bruton et al. 2003; Tsuboi et al. 2003; but see Dyachok & Gylfe, 2004). The source of Ca2+ mobilized may include the ER because we find that caffeine and forskolin fail to promote CICR in thapsigargin-treated cells. Moreover, caffeine releases ER Ca2+, as measured in INS-1 cells expressing YC3.3-er.
RYR is implicated in the regulation of [Ca2+]i, exocytosis, apoptosis, and endosome function in ß cells (Islam et al. 1992, 1998; Takasawa et al. 1998; Holz et al. 1999; Lemmens et al. 2001; Quesada et al. 2002; Bruton et al. 2003; Kang & Holz, 2003; Johnson et al. 2004a,b). Because not all studies support these contentions (Tengholm et al. 1998, 1999, 2000), there may exist differences in the levels of expression of RYR when comparing strains of mice or when making comparisons across species lines. The failure of prior studies to detect RYR in ß cells might also be explained by the use of an experimental strategy that relies on treatment of cells with diazoxide and verapamil (Dyachok & Gylfe, 2004; Dyachok et al. 2004). These agents disrupt CICR by virtue of their ability to inhibit Ca2+ influx (Meissner, 2002). In contrast, human ß cells not treated with these agents exhibit an increase of [Ca2+]i in response to a low concentration of ryanodine, thereby demonstrating the presence of RYR in this cell type (Johnson et al. 2004a). Similarly, human ß cells not treated with diazoxide or verapamil exhibit an increase of [Ca2+]i in response to GLP-1, an action inhibited by a high concentration of ryanodine (Holz et al. 1999).
New studies demonstrate the presence of immunologically detectable RYR in human and mouse ß cells (Johnson et al. 2004a,b). Although the molecular nature of ß cell RYR remains to be determined, it may exhibit features somewhat different from the isoforms described to date. Indeed, splice variants of RYR1 and RYR2 mRNA are expressed in islets (Lee et al. 2002; Okamoto et al. 2004). Sequence variations of this type may explain why some have found it difficult or impossible to detect RYR mRNA by RT-PCR of purified preparations of mouse ß cells (Beauvois et al. 2004).
Potential IP3-R-mediated signalling properties of the GLP-1-R
Actions of GLP-1 mediated by the IP3-R are also of interest. Although activation of the GLP-1-R fails to stimulate IP3 production in islets or INS-1 cells (Fridolf & Ahren, 1991; Zawalich et al. 1993; Kang et al. 2003), cAMP can, under certain conditions, facilitate IP3-R function. This action of cAMP is PKA mediated (Bruce et al. 2003), and is reported to be operational in mouse ß cells (Liu et al. 1996; Dyachok et al. 2004). Mobilization of Ca2+ from IP3-R-regulated Ca2+ stores may explain, at least in part, how GLP-1 stimulates exocytosis in ß cells (Dyachok et al. 2004). It may also play some role in the stimulation of ß cell mitochondrial ATP production (Tsuboi et al. 2003) and mitogen-activated protein (MAP) kinase signalling (Arnette et al. 2003). Therefore, it is noteworthy that we demonstrate IP3-R-mediated CICR sensitized by a cAMP-elevating agent and which is blocked by treatment of ß cells with heparin, an inhibitor of IP3-R function.
It is also important to note that Ca2+ exerts stimulatory effects on IP3 production in insulin-secreting cells (Roe et al. 1993; Gromada et al. 1996). This action of Ca2+ is most likely a consequence of its ability to activate phospholipase C (Berridge et al. 2003). If such an action of Ca2+ were to exist in ß cells loaded with NP-EGTA, the uncaging of Ca2+ might raise levels of IP3, thereby stimulating the IP3-R. CICR triggered in this manner might be facilitated under conditions in which the IP3-R is sensitized by cAMP-elevating agents. Because evidence also exists for ‘atypical’ mechanisms of Ca2+ release in the ß cell (Johnson & Misler, 2002; Masgrau et al. 2003; Mitchell et al. 2003; Beauvois et al. 2004), actions of cAMP-elevating agents described here may not be restricted to RYR or the IP3-R.
Epac-mediated sensitization of CICR
Prior studies of multiple cell types demonstrate that cAMP acts via PKA to sensitize RYR and the IP3-R (Marx et al. 2000; Bruce et al. 2003). Here we report that the action of cAMP may also be Epac-mediated. A cAMP analogue selective for Epac sensitizes the CICR mechanism of ß cells, whereas transfection of INS-1 cells with dominant negative Epac diminishes CICR. Although prior studies implicate Epac2 in this process (Kang et al. 2001, 2003; Tsuboi et al. 2003), we are the first to demonstrate that Epac1 may also be a contributing factor. One established downstream effector of Epac is the small-molecular weight GTPase Rap. Although the signalling properties of Rap are not fully understood, evidence exists that it promotes protein kinase-mediated phosphorylation independently of PKA (Holz, 2004a). Therefore, cAMP-dependent sensitization of intracellular Ca2+ release channels might require formation of an activated Epac/Rap signalling complex. Such an action of Epac would complement its ability to interact with secretory granule-associated proteins (Kashima et al. 2001; Shibasaki et al. 2004) and to regulate fast Ca2+-dependent exocytosis in the ß cell (Eliasson et al. 2003).
Conclusion
Findings presented here suggest that RYR and IP3-R Ca2+ release channels mediate the Ca2+ mobilizing action of GLP-1 in ß cells. Both types of Ca2+ release channels act as cAMP and Ca2+ coincidence detectors, because their opening is facilitated by a simultaneous increase of intracellular cAMP and Ca2+ concentrations. PKA and Epac-mediated sensitization of CICR demonstrated here may provide a new explanation for how GLP-1 interacts with ß cell glucose metabolism to stimulate insulin secretion.
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