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Endocrinology, doi:10.1210/en.2004-0165
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Endocrinology Vol. 145, No. 10 4635-4644
Copyright © 2004 by The Endocrine Society

Mechanisms of Arginine-Vasopressin-Induced Ca2+ Oscillations in ß-Cells (HIT-T15): A Role for Oscillating Protein Kinase C

Michael Schaefer, Harald Mischak, Susanne Schnell, Anne Griese, Roman Iakubov, Gabriele Riepenhausen and Christof Schöfl

Institut für Pharmakologie (M.S.), Charité-Universitätsmedizin Berlin, 14195 Berlin, Germany; and Abteilung für Nephrologie (H.M.) and Abteilung für Gastroenterologie, Hepatologie, und Endokrinologie (S.S., A.G., R.I., G.R., C.S.), Medizinische Hochschule Hannover, 30623 Hannover, Germany

Address all correspondence and requests for reprints to: Dr. Christof Schöfl, Abteilung für Gastroenterologie, Hepatologie, und Endokrinologie, Medizinische Hochschule Hannover, 30623 Hannover, Germany. E-mail: schoefl.christof{at}mh-hannover.de.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
We examined the role of protein kinase C (PKC) for the generation of arginine-vasopressin (AVP)-linked Ca2+ oscillations in ß-cells (HIT-T15). Activation of PKC by phorbol-12,13-dibutyrate (PDBu) reduced the frequency and finally abolished AVP-induced Ca2+ oscillations. The PKC inhibitors Gö 6976, Ro-32-0432, or chelerythrine converted Ca2+ oscillations to a plateau-like rise in cytosolic free Ca2+, and PKC down-regulation reduced the percentage of cells exhibiting AVP-induced Ca2+ oscillations. Several mechanisms were identified by which PKC could exert feedback on the AVP-linked Ca2+ oscillator. PDBu, but not the PKC inhibitors, inhibited AVP-stimulated inositol 1,4,5-trisphosphate production and mobilization of internal Ca2+. Ca2+ influx through voltage-sensitive Ca2+ channels was attenuated by PDBu and PKC inhibitors, indicating complex PKC-dependent regulation of voltage-sensitive Ca2+ channels involving stimulatory as well as inhibitory components. Furthermore, AVP caused oscillatory translocation of yellow fluorescent protein (YFP)-tagged PKC{alpha} and PKCßI to the plasma membrane, which paralleled the Ca2+ oscillations in single cells. Repetitive translocation of YFP-PKC{alpha} and -PKCßI could also be elicited by repetitive release of caged Ca2+. By contrast, AVP-stimulated translocation of YFP-PKC{epsilon} was monophasic, not synchronized with Ca2+ oscillations, and could not be mimicked by release of caged Ca2+. In conclusion, undisturbed activation of PKCs is a necessary intermediate to generate or maintain AVP-induced Ca2+ oscillations in pancreatic ß-cells. The data further suggest that classical PKCs, predominantly by inhibition of inositol 1,4,5-trisphosphate production, provide the negative feedback required for AVP-induced Ca2+ oscillations to occur that is mediated by their repetitive activation by oscillating Ca2+ concentrations.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
NEUROTRANSMITTERS AND HORMONES, like acetylcholine, arginine-vasopressin (AVP), or bombesin, that activate receptors coupled to the Ca2+-phosphoinositide (PI) pathway cause a rise in cytosolic free Ca2+ ([Ca2+]i) and stimulate insulin secretion from normal and transformed ß-cells in the presence of glucose (1, 2, 3, 4, 5, 6). The generation of Ca2+ signals by phospholipase C (PLC)-linked hormones requires the formation of inositol 1,4,5-trisphosphate (IP3) from PLC-mediated breakdown of phosphatidylinositol 4,5-bisphosphate. IP3 binds to specific receptors in the endoplasmic reticulum, thereby mobilizing [Ca2+]i. In addition, influx of Ca2+ from the outside is necessary for sustained PLC-linked Ca2+ signals, which in the excitable ß-cells predominantly occurs through voltage-sensitive Ca2+ channels (VSCCs) (4, 7, 8, 9). In HIT-T15 cells and primary ß-cells, PLC-linked agonists at low, near physiological concentrations cause Ca2+ oscillations whose frequency is determined by the extracellular agonist concentration, whereas the amplitude remains constant (4, 7, 10, 11, 12). This indicates that the cytosolic Ca2+ signal evoked by PLC-linked agonists might be primarily frequency encoded. As transient rises in [Ca2+]i increase, the rate of exocytosis from single HIT-T15 and primary ß-cells, a functional role of PLC-linked Ca2+ oscillations, can be assumed (13, 14). Whereas the basic features of PLC-linked Ca2+ oscillations have been elucidated in recent years, the mechanisms underlying the PLC-linked Ca2+ oscillations are still unknown. Various positive and negative feedback mechanisms have been described and used to model transient Ca2+ oscillations in a number of cell types (15, 16). Several lines of evidence mainly from nonexcitable cells suggest that protein kinase C (PKC), which is coactivated on stimulation of receptors coupled to the Ca2+-PI pathway, may control PLC-linked Ca2+ oscillations via a negative feedback loop (15, 16, 17, 18, 19, 20, 21).

Several PKC isoenzymes exist, which are classified in subfamilies based on their differential sensitivity toward stimuli (22). The conventional (c) PKC isoenzymes consist of the {alpha}-, ßI-, ßII-, and {gamma}-isoforms that are dually activated by Ca2+ and diacylglycerol (DAG). Novel (n) PKCs, like PKC{delta}, PKC{epsilon}, PKC{eta}, and PKC{theta}, are activated by DAGs, whereas atypical (a) PKC isoforms such as PKC{zeta} or PKC{iota}/{lambda} are unresponsive to both Ca2+ and DAG. Definitive evidence for a role of PKC(s) in the generation of PLC-linked Ca2+ oscillations is limited to a very few examples (19, 20, 21). The elements of the Ca2+-PI signaling pathway, however, differ among cell types (23), and mechanisms described in one particular cell or in heterologous cell systems expressing recombinant membrane receptors cannot be simply transferred to other cell types. In the case, for example, of metabotropic glutamate receptor 5-induced Ca2+ oscillations, which is one of the most intensively studied systems, the data so far are controversial (19, 20, 24). Because there is little information about the processes controlling the dynamics of Ca2+ oscillations elicited by agonists activating the Ca2+-PI pathway in pancreatic ß-cells, we explored a potential role of PKCs for the generation of PLC-linked Ca2+ oscillations in excitable pancreatic ß-cells (HIT-T15) using AVP as the agonist.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
HIT-T15 cell culture
HIT-T15 cells were grown in RPMI 1640 medium containing 10 mM glucose supplemented with 10% fetal calf serum (vol/vol), 100 U/ml penicillin, and 100 µg/ml streptomycin at 37 C in 5% CO2 and 95% air (vol/vol). All experiments were performed with cells from passages 65–86.

Western blotting
Extracts of HIT-T15 cells treated with or without phorbol-12-myristate-13-acetate (PMA) (1 µM) for 15 min were prepared by adding a lysis buffer (50 mM Tris/HCl, pH 7.5; 10 mM EGTA, 2 mM EDTA, 3 mM dithiothreitol, and 1 mM phenylmethylsulfonylfluoride) and subsequent sonication on ice. The broken cells were centrifuged at 100,000 x g for 60 min at 4 C. The supernatant was designated the cytosolic fraction and the pellet the membrane fraction. The pellet was resuspended by sonication in Tris-lysis buffer supplemented with Nonidet P-40 (1% vol/vol). The cytosolic and membrane fractions were subjected to SDS-PAGE and electrophoretically transferred to a polyvinylidene fluoride membrane (Millipore, Eschborn, Germany) using a semidry blotting chamber (Bio-Rad Laboratories, Munich, Germany). Blots were probed with isozyme-specific polyclonal antibodies. The antibodies for PKC{alpha}, PKC{gamma}, PKC{epsilon}, PKC{delta}, and PKC{zeta} were obtained from Invitrogen (Karlsruhe, Germany), and antibodies for PKCßI, PKCßII, and PKC{iota} from Santa Cruz Biotechnology (Santa Cruz, CA). The specificity of the interaction was assessed by either use of the isoform-specific blocking peptide provided by the manufacturer or comparison with the expression in rat brain. The secondary antibody was a goat antirabbit IgG conjugated to alkaline phosphatase, which was used at a dilution of 1:5000 and visualized by enhanced chemiluminescence using 3-(4-methoxyspiro{1,2-dioxetane-3,2'-(5'-chloro)tricyclo[3,3.1.11,7]decan}-4-yl)phenylphosphate (Calbiochem, Bad Soden, Germany) as a substrate.

Measurement of IP3
HIT-T15 cells (10–30 x 106 cells) grown in petri dishes (20 cm2) were preincubated for 30 min in a medium containing 130 mM NaCl, 4.7 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 1.5 mM CaCl2, 10 mM glucose, 20 mM HEPES, 0.1% BSA (wt/vol), and 10 mM LiCl, aerated with 100% O2 (vol/vol) (pH 7.4) at 37 C. Cells were washed and incubated for 20 sec with or without the respective compounds, and intracellular IP3 was determined using a receptor competition assay kit (Amersham, Braunschweig, Germany) as described (25).

Expression of fluorescent PKC fusion proteins
Constructs encoding human PKC isoenzymes C-terminally fused to green fluorescent (GFP) or yellow fluorescent protein (YFP) were used as described earlier (26). HIT-T15 cells were grown in 35-mm dishes and transiently transfected with the constructs (2 µg plasmid DNA per dish) and 4 µl of a Fugene 6 transfection reagent (Roche Molecular Biochemicals, Mannheim, Germany) according to the manufacturer’s instructions. After 24 h, transfected cells were seeded on glass coverslips and used for confocal microscopy or digital videoimaging experiments the following day.

Measurement of [Ca2+]i and fluorescence imaging
HIT-T15 cells cultured on coverslips were loaded with 5 µM fura 2/acetoxymethyl ester (AM) for 30 min at 37 C. The loading buffer was as follows: 130 mM NaCl, 4.7 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 1.5 mM CaCl2, 10 mM glucose, 20 mM HEPES, 2% BSA (wt/vol), and 0.1% pluronic acid (wt/vol), gassed with 100% O2 (vol/vol) (pH 7.4). After loading, the coverslips were washed, mounted in a temperature-controlled superfusion chamber (37 C), and placed on the stage of an Axiovert IM 135 equipped with a x40/1.3 Achrostigmat oil immersion objective (Carl Zeiss, Göttingen, Germany). The chamber was superfused with the same buffer as used for fura 2 loading with 0.1% BSA (wt/vol) and without pluronic acid. The flow rate was 0.75–2 ml/min. [Ca2+]i was measured in cells of average size and healthy appearance (round in shape, no membrane blebs). Fura 2 fluorescence from a single cell was recorded with a dual-excitation spectrofluorometer system (Deltascan 4000, Photon Technology Instruments, Wedel, Germany). [Ca2+]i was calculated according to the following formula: [Ca2+]i = KD x B x (R – Rmin)/(Rmax – R), where KD = 224 nM (27), R is the ratio of fluorescence intensities excited at 340 and 380 nm, Rmax, Rmin, and B are constants that were determined in the superfusion chamber from solutions containing fura 2 (1 µM) and various concentrations of free Ca2+ (data not shown).

For coimaging of fluorescent proteins (YFP) and [Ca2+]i, HIT-T15 cells transiently transfected with the respective construct were incubated with fura 2/AM and washed before the experiment as described above. Imaging was performed by exciting the probe with a monochromator (Polychrome II, Till-Photonics. Martinsried, Germany) through a x40/1.3 F-Fluar objective (Zeiss), and images were recorded with a cooled CCD camera (Imago; Till-Photonics). A dichroic mirror (502 nm inflection point) with extended reflectivity (320–500 nM) was combined with a 512-nm long-pass filter. Cells were alternately excited at 340, 358, 380, 450, and 480 nm, and calibration of [Ca2+]i and monitoring of the relative plasma membrane association of the respective YFP-fused PKC isoenzyme were done applying a spectral fingerprinting method as described previously (28).

For coimaging changes in the membrane potential and [Ca2+]i, HIT-T15 cells were coincubated for 15 min at 37 C and another 15 min at ambient temperature with loading buffer supplemented with the potentiometric dye di-8-ANEPPS (10 µM) and fura 2/AM (1 µM). Coverslips were rinsed with loading buffer and imaged as described for coimaging of [Ca2+]i and fluorescent PKC constructs with the exception that the excitation wavelengths were set to 340, 380, 450, and 490 nm. In all experiments, fura 2 fluorescence was very low (less than 0.05% of the fluorescence of di-8-ANEPPS excited at 450 nm), assuring that fura 2 fluorescence did not bleed into di-8-ANEPPS fluorescence excited at 450 or 490 nm. Mean fluorescence intensities were calculated over regions of interest covering single cells, corrected for background signals, and expressed as ratios F340 nm/F380 nm and F480 nm/F440 nm representing fluctuations of [Ca2+]i and the membrane potential, respectively. To assess changes in the di-8-ANEPPS fluorescence during complete depolarization, cells were superfused with a high K+-buffer (loading buffer containing 80 mM KCl and only 50 mM NaCl) at the end of each experiment.

Confocal laser-scanning microscopy and photolysis of caged Ca2+
A LSM 510 inverted confocal laser-scanning microscope (Zeiss) was used for confocal imaging and photolysis of caged Ca2+. YFP was excited at 488 nm through a Plan-Apochromat x63/1.4 objective. In some experiments, fura 2 was loaded and alternately excited with the 351- and 364-nm lines of a UV argon laser through a Plan-Neofluar x40/1.3 objective. For intracellular loading of caged Ca2+, cells were incubated for 30 min at 20 C in loading buffer supplemented with o-nitrophenyl-EGTA/AM (10 µM; Molecular Probes, Eugene, OR). For photolysis of caged Ca2+, YFP was imaged with a dual-reflectivity beam splitter (364 and 488 nm), and photolysis was achieved by brief pulses (20–100 msec) of 364-nm laser light that was locally applied at maximal intensity through the control of the LSM software (Zeiss). The tube current of the UV laser was adjusted to obtain nonsaturating photolysis of caged Ca2+, resulting in [Ca2+]i signals, which were just sufficient to induce brief translocation pulses of classical PKC{alpha} or -ßI (typically around 200–400 nM as tested in fura 2-loaded cells with ionomycin and various concentrations of extracellular free Ca2+ buffered in 5 mM EGTA).

Materials
Fura 2/AM and di-8-ANEPPS were purchased from Molecular Probes; verapamil was provided by Knoll (Ludwigshafen, Germany); RPMI 1640, penicillin, and streptomycin were from Invitrogen; collagenase was from Roche Molecular Biochemicals; thapsigargin, PMA, phorbol-12,13-dibutyrate (PDBu), 4{alpha}-phorbol-12,13-didecanoate (4{alpha}-PDD), chelerythrine chloride, Gö 6976, and Ro-32-0432 were from Calbiochem. All other reagents were from Sigma (Deisenhofen, Germany) or Merck (Darmstadt, Germany). Stock solutions were prepared in water or as follows: AVP (100 µM in 0.01 N HCl), thapsigargin (5 mM in dimethylsulfoxide), Gö 6976, Ro-32-0432, chelerythrine chloride, PMA and PDBu (1 mM in dimethylsulfoxide).

Statistics
Unless representative tracings are shown, values are means ± SEM. Statistical analysis was performed using Student’s t test for paired or unpaired data when two samples were compared. Multiple comparisons were assessed by ANOVA followed by Student-Newman-Keuls test. P < 0.05 was considered as significantly different.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Activation of PKC by phorbol esters inhibits AVP-induced Ca2+ oscillations
In the presence of glucose (10 mM) [Ca2+]i amounted to 174 ± 5 nM (n = 73 cells) in HIT-T15 cells. When applied at submaximally effective concentrations, AVP (1 nM; Figs. 1Go and 2Go) induced Ca2+ oscillations as reported previously (4, 29, 30). In the presence of AVP (1 nM), oscillatory [Ca2+]i spikes peaked at 572 ± 21 nM (n = 86) with a mean frequency of 0.84 ± 0.06 min–1 (n = 24 cells). Both amplitude and frequency of the Ca2+ transients in response to the same agonist concentration varied from cell to cell (Figs. 1Go and 2Go). This presumably reflects heterogeneity of single cells regarding membrane receptors or elements of the Ca2+-PI signaling pathway. Because there was no synchronization among neighboring cells, we concluded that the observed regenerative Ca2+ responses were triggered independently from transcellular signaling through connexins or electrical coupling between cells (31, 32, 33, 34). The phorbol esters PMA (0.01–1 µM; data not shown) and PDBu (10–100 nM; Fig. 1AGo) reduced the frequency and finally stopped the AVP-induced Ca2+ oscillations (n = 8 cells). This effect was concentration dependent at the level of a single cell and fully reversible in all cells tested (Fig. 1AGo). Despite 10-fold higher concentrations, the control compound 4{alpha}-PDD (1 µM) had no effect on the AVP-induced Ca2+ oscillations (n = 3 cells; Fig. 1BGo).



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  		FIG. 1. Effect of the phorbol esters on AVP-induced Ca2+ oscillations. A, Concentration-dependent effect of PDBu on ongoing Ca2+ oscillations (n = 6 cells). B, 4{alpha}-PDD (1 µM), an inactive control compound, was applied on oscillating HIT-T15 cells (n = 3). Bars indicate the presence of the respective compounds in the superfusion medium. Representative tracings are shown.

 


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  		FIG. 2. Effect of PKC antagonists on AVP-induced Ca2+ oscillations in HIT-T15 cells. A–C, Treatment with Gö 6976, Ro-32-0432, or chelerythrine abolished the AVP-induced Ca2+ oscillations and transformed them into a plateau-like rise in [Ca2+]i. D–F, AVP-induced Ca2+ responses before and after pretreatment with Gö 6976 (D) or Ro-32-0432 (E) or in vehicle-treated control cells (F). Bars indicate the presence of the respective compounds in the superfusion medium. Depicted are representative tracings of three or four cells showing similar responses.

 
PKC inhibitors prevent AVP-induced Ca2+ oscillations
The specific PKC inhibitor chelerythrine chloride (10 µM) and the PKC inhibitors Gö 6976 (100 nM) and Ro-32-0432 (1 µM), which are regarded as selective inhibitors of cPKC isoenzymes, stopped the AVP-induced Ca2+ oscillations and transformed the Ca2+ signal into a plateau-like rise in [Ca2+]i (Fig. 2Go, A–C). After pretreatment for 5 min with Gö 6976 (100 nM) or Ro-32-0432 (1 µM), the generation of Ca2+ oscillations was prevented, and AVP (1 nM) caused a nonoscillatory biphasic Ca2+ response with a peak followed by a long-lasting plateau (Fig. 2Go, D and E). Down-regulation of phorbol ester-sensitive cPKCs and nPKC{epsilon} by long-term pretreatment with PDBu (1 µM for 24 h) reduced the number of cells exhibiting AVP-induced Ca2+ oscillations from 21 of 39 cells (54%) in controls to 4 of 39 cells (10%) in the pretreated cells (P < 0.001).

PKC activation inhibits IP3 formation and internal Ca2+ mobilization by AVP
IP3-mediated mobilization of internal Ca2+ underlies the generation of PLC-linked Ca2+ signals. We therefore sought to determine whether activation of PKC by phorbol esters could interfere with either IP3 formation or IP3-mediated mobilization of internal Ca2+. A pretreatment (5 min) with PDBu (100 nM) abolished the AVP-induced Ca2+ response in Ca2+-free medium (Fig. 3BGo) and completely blocked AVP (10 nM)-induced IP3 formation (Fig. 3CGo). The PKC inhibitors Gö 6976 (100 nM) and Ro-32-0432 (1 µM) had no effect on the mobilization of internal Ca2+ by AVP (10 nM). In the absence of external Ca2+, AVP (10 nM) increased [Ca2+]i by 405 ± 62 nM (n = 7, Fig. 3AGo). After 5 min pretreatment with Gö 6976 (100 nM) or Ro-32-0432 (1 µM), the amplitude of the remaining Ca2+ signal was 69 ± 8% (n = 15) and 74 ± 11% (n = 15), compared with untreated controls, which was statistically not significant.



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  		FIG. 3. Effect of the phorbol ester PDBu on the AVP-induced Ca2+ mobilization and formation of IP3. A, Mobilization of internally stored Ca2+ by AVP (10 nM) in a Ca2+-free superfusion medium supplemented with EGTA (2.5 mM) (n = 18 cells). B, Effect of pretreatment with PDBu (100 nM for 5 min) on the AVP (10 nM)-induced Ca2+ mobilization (n = 6 cells). Bars indicate the presence of the respective agents in the superfusion medium. Representative tracings are shown (A and B). C, Effects of PDBu pretreatment (100 nM for 5 min) on basal and stimulated IP3 formation in HIT-T15 cells. Incubations were stopped 20 sec after the addition of agonist (AVP, 10 nM) or solvent (control). IP3 concentrations were determined with a receptor competition assay. Values are means ± SEM of three independent experiments determined in triplicates. **, Significant difference, compared with unstimulated cells (P < 0.01).

 
PKC modulates Ca2+ influx through VSCCs
Simultaneous measurements of [Ca2+]i and changes in the membrane potential by di-8-ANEPPS in single HIT-T15 cells revealed a membrane hyperpolarization in response to AVP after the initial increase in [Ca2+]i as evidenced by an increase in the ratio of di-8-ANEPPS fluorescence excited at 490 and at 450 nm by 1.1 ± 0.2% (n = 35 cells). During ongoing AVP-induced Ca2+ oscillations, however, there was no significant correlation between changes in membrane potential and the occurrence of individual Ca2+ transients (–0.02 ± 0.02%, n = 51 transients). Based on the changes in the di-8-ANEPPS fluorescence signal on maximal depolarization by 80 mM K+ (–2.1 ± 0.29%, n = 35), the sensitivity of the method for detecting changes of membrane potential can be estimated to be in the range of about 2–5 mV. These data imply that the PLC-dependent Ca2+ oscillator in excitable ß-cells is driven independently of changes in the membrane potential. Nevertheless, Ca2+ influx through VSCCs is required for the sustained generation of AVP-induced Ca2+ oscillations in ß-cells (4, 30). Hence, we investigated a potential role of PKC in the control of voltage-sensitive Ca2+ influx. PDBu (100 nM) or PMA (100 nM) but not 4{alpha}-PDD (100 nM) increased [Ca2+]i by 46 ± 12 nM (n = 6, P < 0.001). This increase in [Ca2+]i required the presence of extracellular Ca2+ and could be blocked by verapamil (50 µM), which demonstrates enhanced Ca2+ influx through VSCCs by activation of classical phorbol ester receptors (Fig. 4AGo). Repetitive membrane depolarization elicited by 20-sec pulses of K+ (45 mM) every 120 sec caused parallel rises in [Ca2+]i presumably through Ca2+ influx via VSCCs (Fig. 4Go, B and C). Inhibition of PKC activity by Ro-32-0432 (1 µM) or Gö 6976 (100 nM) reversibly decreased the amplitude of K+-triggered Ca2+ signals by 33 ± 6% (P < 0.01, n = 5) or 42 ± 11% (P < 0.01, n = 5, Fig. 4BGo), respectively. These data are consistent with an augmentation of voltage-sensitive Ca2+ influx by PKC-dependent processes. PDBu (100 nM) but not 4{alpha}-PDD (100 nM, not shown), however, also reduced the amplitude of the K+ (45 mM)-induced increases in [Ca2+]i by 29 ± 4% (P < 0.001, n = 6, Fig. 4CGo). We conclude that VSCCs are involved in the maintenance of PLC-dependent [Ca2+]i oscillations by mediating sustained Ca2+ influx and are regulated by classical or nPKCs.



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  		FIG. 4. Effects of PDBu and the PKC inhibitor Gö 6976 on Ca2+ influx through VSCCs. A, Effects of the VSCC blocker verapamil on the PDBu-induced increase in [Ca2+]i (n = 6 cells). B and C, Repetitive membrane depolarization elicited by 20-sec pulses of K+ (45 mM) every 120 sec in the superfusion medium causes parallel rises in [Ca2+]i. The effects of treatment with either the PKC inhibitor Gö 6976 (100 nM) (n = 5 cells) (B) or the PKC activator PDBu (100 nM) (n = 6 cells) (C) on K+ (45 mM)-induced increases in [Ca2+]i are tested. Note the reduced peak [Ca2+]i during application of the modulators. Bars indicate the presence of the respective agents in the superfusion medium. Representative tracings are shown.

 
PKC isoenzymes expressed and activated by phorbol esters in HIT-T15 cells
Western blotting was performed to determine which PKC isoforms are expressed in HIT-T15 cells and which are affected by treatment with phorbol esters. The cPKC isoforms cPKC{alpha}, cPKCßI, cPKCßII but not cPKC{gamma} and the nPKC isoforms nPKC{epsilon} and nPKC{delta} as well as the atypical isoforms aPKC{zeta} and aPKC{iota} were expressed in HIT-T15 cells (see Fig. 6AGo, not shown nPKC{delta} and the aPKCs). The nPKC{eta}, nPKC{theta}, and aPKC{lambda} were not tested for. Stimulation of HIT-T15 cells with PMA (1 µM) for 15 min resulted in a redistribution of cPKC{alpha}, cPKCßI, cPKCßII, and nPKC{epsilon} from the soluble to the particulate fraction (Fig. 5AGo). Pretreatment with PMA (1 µM) for 24 h down-regulated cPKC{alpha}, cPKCßI, cPKCßII, and nPKC{epsilon} but not the aPKCs (not shown). To visualize the subcellular distribution and monitor the translocation of PKC isoenzymes in living cells, GFP-fused PKC isoenzymes were transiently expressed in HIT-T15 cells and imaged by confocal laser-scanning microscopy. All fluorescent PKC fusion proteins were mainly detectable in the cytosol of resting HIT-T15 cells as shown in Fig. 5BGo. Treatment with PMA (1 µM) but not with the inactive phorbol ester 4{alpha}-PDD (1 µM, data not shown) caused a translocation of PKC{alpha}-GFP, PKCßI-GFP, PKCßII-GFP, and PKC{epsilon}-GFP from the cytosol to the plasma membrane within several minutes (Fig. 5BGo).



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  		FIG. 6. AVP-induced Ca2+ oscillations and spatiotemporal translocation of fluorescent PKC fusion proteins. A and B, Living HIT-T15 cells transiently expressing cPKCßI-YFP (A) or cPKC{epsilon}-YFP (B) were stimulated with AVP (100 nM) for the indicated times. Before (control) and after stimulation with AVP, the distribution of PKC fusion proteins was imaged by confocal laser-scanning microscopy. C and D, AVP (1 nM)-induced translocations of YFP-fused cPKC{alpha} (n = 4 cells) (C) and nPKC{epsilon} (n = 3 cells) (D) were detected by spectrally resolving digital videomicroscopy. Ca2+ oscillations (gray lines) and PKC translocations (black lines) were simultaneously recorded in the same cells loaded with the Ca2+ indicator fura 2. YFP fluorescences over regions corresponding to the plasma membrane (Fmembrane) were divided by the cytosolic (Fcytosol) signals and normalized to the initial values. Representative results for each PKC isoform are depicted.

 


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  		FIG. 5. Expression and translocation of PKC isoenzymes by PMA in HIT-T15 cells. A, Western blot analysis of ß-cell cytosol or membrane fractions demonstrate the expression of PKC{alpha}, PKCßI, PKCßII, and PKC{epsilon}. Treatment of HIT-T15 cells with PMA (1 µM) for 5 min caused a redistribution of PKC{alpha}, PKCßI, PKCßII, and PKC{epsilon} from the cytosol to the membrane fraction. HIT-T15 cells were fractionated, and fractions were probed with isoenzyme-specific PKC antibodies as described in Materials and Methods. Immunoblot analyses were repeated three to four times using various cell passages with similar results. Specificity was verified with isozyme-specific blocking peptides. C, Cytosolic fraction; M, membrane fraction. B, Subcellular distribution of PKC fusion proteins transiently expressed in HIT-T15 cells before (control) and 7 min after treatment with PMA (1 µM). Living cells transiently transfected with constructs encoding the respective PKC fusion proteins were imaged by confocal laser-scanning microscopy. Representative cells of at least three independent experiments are shown.

 
AVP causes oscillatory translocation of cPKC isoenzymes entrained by Ca2+ oscillations in single HIT-T15 cells
Simultaneous measurements of [Ca2+]i and PKC translocation in AVP (1 nM)-stimulated HIT-T15 cells revealed repetitive translocations of YFP-tagged cPKC{alpha} (Fig. 6CGo) and cPKCßI (not shown). The PKC translocations closely paralleled the onset and peaks of secondary Ca2+ spikes with delay times of 1–5 sec. Typical thresholds for the [Ca2+]i-driven translocation of both classical PKCs were in the range of 150–250 nM. Repetitive release of caged Ca2+ also caused repetitive translocation of cPKC{alpha}-YFP and cPKCßI-YFP as observed by confocal laser scanning microscopy (Fig. 7Go, A and B). The shorter lag period of less than 200 msec after partial photolysis of intracellularly loaded caged Ca2+ presumably reflects the higher resolution of the confocal detection system and the faster rise of [Ca2+]i, compared with the more sinusoidal shape of hormonally induced regenerative Ca2+ spikes. By contrast, AVP (1 nM) induced a slow and monophasic translocation of nPKC{epsilon}, which was not synchronized with Ca2+ oscillations (Fig. 6DGo). The translocation of nPKC{epsilon} was weak and peaked about 40–120 sec (n = 4) after the addition of AVP. In some cells, the AVP-induced translocation of nPKC{epsilon}-YFP proceeded even during the falling phase of [Ca2+]i and the onset of the following spike (Fig. 6DGo). Furthermore, release of caged Ca2+ caused no translocation of nPKC{epsilon}-YFP (Fig. 7CGo). Thus, changes in [Ca2+]i are the driving force for repetitive translocation of cPKCs to the cell membrane of HIT-T15 cells.



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  		FIG. 7. Spatiotemporal plasma membrane translocation of classical but not novel PKC isoenzymes during photolysis of caged Ca2+ in single HIT-T15 cells. A and B, High-speed confocal imaging of PKC translocations during repetitive photolysis of caged Ca2+ (intracellularly loaded o-nitrophenyl-EGTA; 10 µM). Arrows indicate brief pulses of UV laser light causing partial photolysis of intracellularly loaded caged Ca2+. Depicted are ratios of fluorescence intensities measured over the plasma membrane (Fmembrane) and the adjacent cytosol (Fcytosol). Note the repetitive translocation of cPKC{alpha}-YFP and cPKCßI-YFP from the cytosol to the plasma membrane (n = 4 each). C, Same experiment as in A but with HIT-T15 cells expressing the nPKC{epsilon}-YFP fusion protein (n = 3 cells). Shown are representative tracings for each PKC isoenzyme.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Our results strongly indicate a crucial role of PKCs for the generation of repetitive Ca2+ oscillations by AVP in pancreatic ß-cells. First, the frequency of AVP-induced Ca2+ oscillations was reduced and finally the Ca2+ signal ceased by direct and constant activation of PKC activity by the phorbol esters PMA and PDBu. This effect of the phorbol esters was concentration dependent and reversible at the level of a single cell, and specificity was indicated by the lack of effect of the inactive control compound 4{alpha}-PDD. Second, exposure to the PKC inhibitors chelerythrine, Gö 6976, or Ro-32-0432 converted the Ca2+ oscillations into a nonoscillatory plateau-like rise in [Ca2+]i. Third, down-regulation of PKCs by chronic treatment with PDBu significantly reduced the percentage of cells that exhibited Ca2+ oscillations in response to AVP stimulation. Taken together, these data demonstrate that the undisturbed activation of classical and/or nPKCs through the PLC pathway is a necessary intermediate to generate or maintain AVP-induced Ca2+ oscillations in pancreatic ß-cells. The data further suggest dominant-negative feedback of PKC-dependent mechanisms on processes that are central to the generation of AVP-stimulated Ca2+ oscillations. This is similar to previous reports from nonexcitable cells such as hepatocytes, in which a role for PKC in the control of IP3-dependent Ca2+ oscillations via a negative feedback loop has been proposed (15, 17, 18, 35).

Phorbol ester treatment inhibits AVP-stimulated IP3 formation as described earlier for mouse islet cells (36), thereby preventing mobilization of Ca2+ from internal stores. Potential mechanisms involved that have been discussed in other cell types are the phosphorylation and uncoupling of the membrane receptor from Gq (19, 20) or phosphorylation of PLCß3, thereby preventing its activation by Gq (37). ß-Cells express both a variant of the V1b-receptor subtype, which has several phosphorylation sites for PKC (38, 39), and PLCß3 (40, 41). Either mechanism would inhibit IP3 formation and consecutive Ca2+ mobilization.

Unlike nonexcitable cell types, a major component of constitutive Ca2+ influx into excitable pancreatic ß-cells occurs through VSCCs (4, 7, 8, 9). Modulation of voltage-sensitive Ca2+ influx has a major impact on PLC-linked Ca2+ oscillations in ß-cells (9, 29, 30). We therefore explored whether PKCs may control AVP-dependent Ca2+ oscillations by interfering with voltage-sensitive Ca2+ entry. PDBu reversibly reduced high K+-induced Ca2+ influx through VSCCs, whereas the inactive control compound 4{alpha}-PDD was ineffective. Because voltage-sensitive Ca2+ influx may modulate IP3 production and/or the IP3-linked Ca2+-release process (42, 43, 44), inhibition of VSCCs could result in negative feedback on the AVP-driven Ca2+ oscillations. The regulation of voltage-sensitive Ca2+ influx, however, by PKCs in ß-cells appears to be more complex. Phorbol esters given alone increased [Ca2+]i by enhancing voltage-sensitive Ca2+ influx, and the PKC inhibitors Gö 6976 and Ro-32-0432 attenuated high K+-induced Ca2+ influx. These data are consistent with the finding that L-type currents are augmented by PKC in HIT-T15 cells (45). The majority of L-type channel transcripts in pancreatic ß-cells are of the Cav1.3 subtype (46, 47), which has been shown to be stimulated by PMA (48). In addition to L-type VSCCs, a low threshold-activated T-type current has been characterized in HIT-T15 cells (49), which could be inhibited by PKC-dependent mechanisms (50). Thus, in ß-cells, like in cardiac and smooth muscle cells (51), PKC-mediated control of VSCCs involves both stimulatory and inhibitory components.

Negative feedback by PKCs on the AVP-coupled Ca2+ oscillator presumably involves inhibition of IP3 formation as well as inhibition of voltage-sensitive Ca2+ influx. PDBu treatment decreased the overall Ca2+-influx through VSCCs by about 30%, which by itself appears to be insufficient to explain the observed effects of phorbol ester treatment on AVP-linked Ca2+ oscillations because in the presence of 10 µM nifedipine, K+-induced Ca2+ signals were reduced by about 90%, but AVP-induced [Ca2+]i oscillations remained intact in a subset of cells (4). Considering that PKCs may inhibit only T-type channels, the inhibitory action of PKCs on those VSCC subtypes would be underestimated, and a greater impact on the Ca2+ oscillations cannot be excluded. In any case, however, we observed, in HIT-T15 cells, an almost complete PKC-mediated inhibition of IP3 production and subsequent Ca2+ mobilization. Because constant or oscillatory formation of IP3 is a prerequisite for PLC-linked Ca2+ oscillations (52), inhibition thereof via PKC could provide the dominant inhibitory mechanism whereby PKCs could principally control the generation and maintenance of AVP-linked Ca2+ oscillations in pancreatic ß-cells (HIT-T15).

Because either PKC inhibition or permanent activation by phorbol esters disrupted ongoing AVP-induced [Ca2+]i oscillations, a rapid activation/deactivation kinetic of PKC appears essential. By Western blot analysis, we could demonstrate that HIT-T15 cells express cPKC{alpha}, cPKCßI, cPKCßII, nPKC{epsilon}, nPKC{delta}, aPKC{zeta}, and aPKC{iota}, which is consistent with previous reports from various other ß-cell lines (53, 54). AVP stimulation of HIT-T15 cells expressing YFP-fused PKC isoforms caused translocation of cPKC{alpha}, cPKCßI, and nPKC{epsilon} to the plasma membrane, indicating their activation in response to receptor stimulation.

By simultaneous measurement of [Ca2+]i and PKC translocation, we demonstrate repetitive translocations of cPKC{alpha} and cPKCßI in response to a PLC-linked agonist in ß-cells. The changes in [Ca2+]i appear to be the principal driving force behind the translocation of cPKCs to the cell membrane in HIT-T15 cells because the translocation of cPKC{alpha} and cPKCßI closely followed the Ca2+-transients with time delays of 1–5 sec and the repetitive release of caged Ca2+ was sufficient to mimic the effects of AVP-stimulated Ca2+-transients on translocation of cPKC{alpha}-YFP and cPKCßI-YFP. The thresholds for global Ca2+ changes to trigger translocation were estimated to be around 200 nM for cPKC{alpha} and cPKCßI. Recently Mogami et al. (55) reported a significantly higher threshold of about 400 nM for global Ca2+ changes to cause translocation of cPKC{alpha}-GFP in the insulin-secreting cell line INS-1, and they demonstrate that, despite similar global [Ca2+]i, stimulation of a muscarinic receptor was less efficient than a tetraethyl ammonium-induced depolarization in inducing cPKC{alpha} translocation in INS-1 cells. One should note that estimates based on a cytosolic indicator dye do not represent the subplasmalemmal Ca2+ concentration, which is decisive for the C2 domain-driven association of cPKCs to the plasma membrane. An attempt to measure the subplasmalemmal [Ca2+]i in pancreatic ß-cells using a recombinant plasma membrane-targeted probe revealed that Ca2+ concentrations in the subplasmalemmal compartment may be significantly higher than global [Ca2+]i, especially when VSCCs are activated (56). Although we could exclude acute opening of VSCCs during the AVP-linked [Ca2+]i oscillations in HIT-T15 cells, part of the [Ca2+]i signal may still be provided by other Ca2+-permeable entry pathways such as store-operated or second messenger-gated conductances. In rat basophilic leukemia cells, a store-operated conductance has been shown to support a sustained translocation of the Ca2+-binding C2 module of cPKC{gamma} on G protein-coupled receptor stimulation (57). Thus, similar conductances may allow a significant amount of Ca2+ to enter the subplasmalemmal space and thereby promote translocation of cPKCs in ß-cells, even under conditions in which global increases in [Ca2+]i appear modest.

Novel PKCs are activated by DAG but, unlike cPKCs, they are unresponsive to changes in [Ca2+]i. AVP caused a slow and monophasic translocation of nPKC{epsilon}, which was not synchronized with Ca2+ oscillations. This is similar to the translocation pattern of nPKC{theta} in acetylcholine-stimulated INS-1 cells (55). However, in contrast to nPKC{theta}, which in INS-1 cells translocated in response to voltage-sensitive Ca2+ influx, nPKC{epsilon} could not be recruited to the plasma membrane just by the release of caged Ca2+ in HIT-T15 cells. This may indicate that the route of Ca2+ elevation is decisive for activation of nPKCs by Ca2+, which is thought to occur indirectly via enhanced DAG formation through stimulatory effects of Ca2+ on PLC activity (55). An alternative explanation could come from cell line- or isoenzyme-specific activation of nPKCs because nPKC{delta} in contrast to nPKC{theta} did not translocate in response to voltage-sensitive Ca2+ influx in MIN6 cells (56). Because of the nonsynchronized appearance of PKC{epsilon} translocation during oscillatory [Ca2+]i responses in AVP-stimulated HIT-T15 cells, a regulatory role of nPKC{epsilon} in triggering the regenerative Ca2+ responses appears unlikely. The synchronized activation and deactivation kinetics, however, of cPKCs and the sensitivity of [Ca2+]i oscillations toward inhibitors of cPKC isoforms point to a regulatory role of cPKCs in triggering and maintaining AVP-induced regenerative [Ca2+]i oscillations in HIT-T15 cells.

Our findings support a simplified mechanistic model in which AVP receptor stimulation causes PLC-mediated breakdown of phosphatidylinositol 4,5-bisphosphate with formation of IP3 and DAG. IP3 mobilizes intracellular Ca2+ leading to an increase in [Ca2+]i, which together with DAG triggers membrane translocation and activation of cPKCs. Phosphorylation by cPKCs of proteins involved in IP3 formation and triggering the release of internal Ca2+ provides negative feedback, thereby terminating the Ca2+ rise. A fall in [Ca2+]i and possibly DAG shuts off the cPKCs with relocation to the cytosol. Recovery from cPKC-mediated negative feedback, i.e. dephosphorylation, is then required for the next Ca2+ transient to be triggered. The frequency of the Ca2+ transients is, therefore, primarily set by the time needed for recovery from cPKC-induced phosphorylation. To work, oscillating activity of cPKCs is required in this model. As observed experimentally, interference with the cPKC activation pattern either by inhibition or constant activation results in the termination of the Ca2+ oscillations.

Frequency-modulated Ca2+ oscillations in response to PLC-linked agonists are a fundamental signaling mechanism in excitable as well as nonexcitable cells controlling cellular responses including hormone secretion, mitochondrial metabolism, and proliferation. In recent years data from various systems that demonstrate that, by changing the frequency and/or the amplitude of Ca2+ oscillations, distinct and differential regulation of cellular functions is possible accumulated (58, 59, 60). Conventional PKCs that are activated by Ca2+ and DAG have been suggested as devices to selectively decode distinct Ca2+ signaling patterns and translate them into distinct cellular responses (61). Our data now demonstrate that, in excitable pancreatic ß-cells, cPKCs may not only be involved in decoding cytosolic Ca2+ signals but also represent an integral component of the PLC-linked Ca2+ oscillator itself.


   Footnotes
 
This work was supported by grants from the Deutsche Forschungsgemeinschaft (Scho 466/2-1 and Scha 941/1).

Abbreviations: a, Atypical; AM, acetoxymethyl ester; AVP, arginine-vasopressin; c, conventional; [Ca2+]i, cytosolic free Ca2+; DAG, diacylglycerol; GFP, green fluorescent protein; IP3, inositol 1,4,5-trisphosphate; n, novel; PDBu, phorbol-12,13-dibutyrate; 4{alpha}-PDD, 4{alpha}-phorbol-12,13-didecanoate; PI, phosphoinositide; PKC, protein kinase C; PLC, phospholipase C; PMA, phorbol-12-myristate-13-acetate; VSCC, voltage-sensitive Ca2+ channel; YFP, yellow fluorescent protein.

Received February 9, 2004.

Accepted for publication July 1, 2004.


   References
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 Introduction
 Materials and Methods
 Results
 Discussion
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