Decoding of Short-lived Ca2+ Influx Signals into Long Term Substrate Phosphorylation through Activation of Two Distinct Classes of Protein Kinase C*,

Hideo MogamiDagger §, Hui Zhang, Yuko SuzukiDagger , Tetsumei UranoDagger , Naoaki Saito||, Itaru Kojima, and Ole H. Petersen**DaggerDagger

From the Dagger  Department of Physiology, Hamamatsu University School of Medicine, 1-20-1 Handayama, Hamamatsu 431-3192, Japan, the ** Medical Research Council (MRC) Secretory Control Research Group, Physiological Laboratory, University of Liverpool, Liverpool L69 3BX, United Kingdom, the  Department of Cell Biology, Institute for Molecular and Cellular Regulation, Gunma University, 3-39-15 Showa-machi, Maebashi 371-8512, Japan, and the || Laboratory of Molecular Pharmacology, Biosignal Research Center, Kobe University, Rokkodai-cho 1-1, Nada-ku, Kobe 657-8501, Japan

Received for publication, October 17, 2002, and in revised form, December 23, 2002

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

In electrically excitable cells, membrane depolarization opens voltage-dependent Ca2+ channels eliciting Ca2+ influx, which plays an important role for the activation of protein kinase C (PKC). However, we do not know whether Ca2+ influx alone can activate PKC. The present study was conducted to investigate the Ca2+ influx-induced activation mechanisms for two classes of PKC, conventional PKC (cPKC; PKCalpha ) and novel PKC (nPKC; PKCtheta ), in insulin-secreting cells. We have demonstrated simultaneous translocation of both DsRed-tagged PKCalpha to the plasma membrane and green fluorescent protein (GFP)-tagged myristoylated alanine-rich C kinase substrate to the cytosol as a dual marker of PKC activity in response to depolarization-evoked Ca2+ influx in the DsRed-tagged PKCalpha and GFP-tagged myristoylated alanine-rich C kinase substrate co-expressing cells. The result indicates that Ca2+ influx can generate diacylglycerol (DAG), because cPKC is activated by Ca2+ and DAG. We showed this in three different ways by demonstrating: 1) Ca2+ influx-induced translocation of GFP-tagged C1 domain of PKCgamma , 2) Ca2+ influx-induced translocation of GFP-tagged pleckstrin homology domain, and 3) Ca2+ influx-induced translocation of GFP-tagged PKCtheta , as a marker of DAG production and/or nPKC activity. Thus, Ca2+ influx alone via voltage-dependent Ca2+ channels can generate DAG, thereby activating cPKC and nPKC, whose activation is structurally independent of Ca2+.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Since the first molecular cloning and sequencing of a bovine brain protein kinase C (PKC)1 (1), PKC has been one of the most extensively studied enzymes in eukaryotic cells. We now know that PKC plays a pivotal role in a myriad of cellular functions. Ten isoforms of PKC have been identified so far and have been classified into three categories based on structural differences in the regulatory domain: conventional PKC (cPKC; PKCalpha , PKCbeta I, PKCbeta II, and PKCgamma ), novel PKC (nPKC; PKCdelta , PKCepsilon , PKCeta , and PKCtheta ), and atypical PKC (PKCzeta and PKClambda ) (2, 3). The C1 and C2 regions in the regulatory domain are responsible for diacylglycerol (DAG) and Ca2+ binding, respectively. DAG comes mainly from plasma membrane phosphatidylinositol 4,5-bisphosphate (PIP2) hydrolysis. This is caused by phospholipase C (PLC) activation, following agonist binding to a G protein-coupled receptor. In excitable cells, cytosolic Ca2+ signals are generated either by Ca2+ influx through voltage-dependent Ca2+ channels (VDCCs) and/or by Ca2+ release from the endoplasmic reticulum through inositol 1,4,5-trisphosphate (IP3) receptors upon binding of IP3, the other product of PIP2 hydrolysis (4, 5). DAG and Ca2+ activate the first family of cPKCs that have both the C1 and C2 regions. The second family of the novel PKCs is also activated by DAG, but in a Ca2+-independent manner because of the absence of the functional C2 region. The third family of the atypical PKCs can be activated by phosphoinositide-dependent kinase 1 in a Ca2+-independent manner (6).

We focused this study on PKCalpha and PKCtheta as representatives of cPKC and nPKC, respectively, to probe further into the mechanisms underlying the Ca2+ signaling-induced activation of cPKC and nPKC. To this end, we employed INS-1 cells, an insulin-secreting cell line established from a rat insulinoma (7), as a model system in which VDCCs are the main pathways for the generation of Ca2+ signals. Key observations from other laboratories have shown that PKCalpha is activated within the physiological Ca2+ concentration range in the presence of both DAG and phosphatidylserine (8) and that depolarizing K+ concentrations evoke an increase in the IP3 concentration in rat pancreatic islets, suggesting PLC-mediated production of DAG (9). Two important questions arise: 1) Can depolarization-evoked Ca2+ influx through the opening of VDCCs activate cPKC? and 2) Can Ca2+ influx also activate nPKC, if the Ca2+ influx results in production of DAG? Furthermore, in INS-1 cells, as in normal insulin-secreting cells, glucose-induced oscillations in membrane potential elicit repetitive openings of VDCCs (10). This is responsible for oscillations in the cytosolic Ca2+ concentration ([Ca2+]i), which causes pulsatile insulin secretion (11). Ca2+ oscillations play an essential role in exocytotic secretion in neuroendocrine cells (12). It is therefore important to know how depolarization-evoked Ca2+ oscillations are decoded into long term physiological modifications, such as insulin secretion, through PKC activation.

Recent advances in the use of green fluorescent protein (GFP) has allowed us to investigate PKC activity in intact living cells by monitoring translocation of GFP-tagged PKC (13, 14). Inactive PKCs are located in the cytosol. Upon activation, following PIP2 hydrolysis, they translocate from the cytosol to other cellular locations, such as the plasma membrane. By simultaneously measuring the cytosolic Ca2+ concentration and the translocation of GFP-PKCgamma in astrocytes, it has been shown that there is a marked temporal correlation between glutamate-elicited Ca2+ spikes and PKC translocation (15). A current model for activation of cPKC (14) proposes that: 1) the [Ca2+]i elevation recruits cPKC to the plasma membrane via the C2 region, 2) the site on the enzyme where the pseudosubstrate inhibitory region in the regulatory domain is occupied at rest is exposed and becomes available for substrate binding, and (3) full activation of the enzyme takes place when DAG tightly tethers the enzyme to the plasma membrane via the C1 region. The sequence of these events suggests that translocation and activation of cPKC may not always correspond. This raises a final question: How do we know when the pseudosubstrate inhibitory region is removed (i.e. when exactly does activation of cPKC take place)?

To address these questions, we monitored translocation of PKCalpha -GFP, as markers for cPKC, in response to depolarization-evoked Ca2+ influx through VDCCs in INS-1 cells. The Ca2+ influx resulted in translocation of PKCalpha -GFP to the plasma membrane. We also assessed the phosphorylation state of the PKC substrate myristoylated alanine-rich C kinase substrate (MARCKS) (16) as another marker of PKC activity, by monitoring translocation of GFP-tagged MARCKS (MARCKS-GFP) with DsRed-tagged PKCalpha (PKCalpha -DsRed). When phosphorylated by PKC, MARCKS translocates from the plasma membrane to the cytosol (17). Translocation of MARCKS-GFP to the cytosol took place as soon as PKCalpha -DsRed translocated to the plasma membrane upon stimulation of Ca2+ influx. These results indicate that the Ca2+ influx can generate DAG, because cPKC is activated by Ca2+ and DAG. We showed this in three different ways by demonstrating: 1) Ca2+ influx-induced translocation of GFP-tagged C1 domain of PKCgamma , 2) Ca2+ influx-induced translocation of GFP tagged pleckstrin homology domain (GFP-PHD), and 3) Ca2+ influx-induced translocation of PKCtheta -GFP as a marker of DAG production. The depolarization-evoked increase in DAG concentration was estimated from in situ calibration to be 1.90 ± 0.02 µM. We have demonstrated for the first time that depolarization-evoked Ca2+ influx can generate DAG, thereby activating cPKC and nPKC. We also observed that MARCKS remained phosphorylated through PKC activation as long as the depolarization-evoked Ca2+ oscillations continued. Our results show that short-lived Ca2+ signals can be transduced via PKC activation into long term phosphorylated MARCKS.

    EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Plasmid Construction

PKCalpha -pEGFP, PKCtheta -pEGFP, pEGFP-N2, and pDsRed1-N1 were obtained from Clontech Lab, Inc. (Palo Alto, CA). To attain brighter fluorescence of MARCKS-GFP, the GFP of MARCKS-GFP (17) was replaced with pEGFP-N2. pEGFP of PKCalpha -pEGFP was replaced with pDsRed1-N1. A GFP-tagged C1 region of PKCgamma (C12-GFP) was produced from a DNA clone of lambda CKRgamma 1, which was subcloned into an expression plasmid for mammalian cells, pTB701 (18). A cDNA fragment of PKCgamma for C1 region with an EcoRI site in the 5' terminus and a BglII in the 3' terminus was produced by PCR using pTB701 as a template. The sense and antisense primers used were 5'-TTGAATTCGCCATGGTGAAGAGCCACAAGTTCACC-3' and 5'-TTAGATCTGTCCACGCCGCAAAGGGAGGG-3', respectively. A PCR product for C1 region of PKCgamma was subcloned into the EcoRI site and the BglII site in GFP containing pTB701 (18). The PCR product was verified by sequencing. A GFP-tagged pleckstrin homology domain of PLCdelta 1 (GFP-PHD) was donated by Dr. Hirose (Tokyo University, Tokyo, Japan) (19).

Cell Culture and Transfection

INS-1 cells (7), insulin-producing cells, were a gift from Dr. Sekine (Tokyo University).The cells were grown in 100-mm culture dishes at 37 °C and 5% CO2 in a humidified atmosphere. The culture medium was RPMI 1640 with 10 mM glucose supplemented with 10% fetal bovine serum, 1 mM sodium pyruvate, and 50 µM mercaptoethanol. For fluorescence imaging, the cells were cultivated on a coverslip at 50% confluency 2 days before transfection. A plasmid of the GFP- or DsRed-tagged proteins was transfected into the cells by lipofection using TransITTM-LT1 (Mirus, Madison, WI). The experiments were performed 2 days after transient transfection.

Solutions

The standard extracellular solution contained 140 mM NaCl, 5 mM KCl, 1 mM MgCl2, 2.5 mM CaCl2, 3 mM glucose, and 10 mM Hepes-NaOH (pH 7.3). The solutions for membrane depolarization contained 105 mM NaCl, 40 mM KCl, 1 mM MgCl2, 2.5 mM CaCl2, 3 mM glucose, and 10 mM Hepes-NaOH (pH 7.3) or contained 120 mM NaCl, 5 mM KCl, 1 mM MgCl2, 2.5 mM CaCl2, 20 mM tetraethylammonium chloride, 3 mM glucose, and 10 mM Hepes-NaOH (pH 7.3). In some experiments, CaCl2 was not included (Ca2+-free solution). The cells, placed on a glass coverslip attached to an open perifusion chamber, were continuously perifused from a gravity-fed system. The experiments were performed in the standard extracellular solution at room temperature, unless otherwise stated.

Materials

Fura2/AM, 12-O-tetradecanoylphorbol-13-acetate (TPA), 1,2-dioctyl-sn-glycerol (DiC8) was purchased from Sigma. U73122 was obtained from Calbiochem (La Jolla, CA).

Imaging Experiments

Epifluorescence Microscopy-- The fluorescence images were captured using a Olympus inverted microscope (60×, water immersion objective, and 60×) equipped with a cooled (-50 °C) coupled charge device digital camera (ORCA-II and ORCA-ER; Hamamatsu Photonics, Hamamatsu, Japan) and recorded and analyzed on a Aquacosmos imaging station (Hamamatsu Photonics). The excitation light source was 150 W xenon lamp with a Polychrome I monochromator (T.I.L.L. Photonics GmbH., Planegg, Germany). GFP fluorescence was excited at 488 nm for high time resolution of GFP-tagged PKCs digital imaging. We measured the fluorescence intensity of the GFP (DsRed)-tagged proteins (PKCs, MARCKS, PHD, and C1 domain) in the cytosol of the cell, excluding the nucleus, and/or at the plasma membrane, as a marker of translocation. These values (F) were normalized to each initial value (F0), so that the relative fluorescence change was referred to as the "ratio F/F0." For simultaneous measurements of the relative change in fluorescence intensity of the GFP-tagged PKCs in the cytosol and [Ca2+]i, GFP fluorescence was excited at 488 nm, whereas Fura2 was excited at wavelengths alternating between 340 and 380 nm. We put a short pass filter of 330-495 nm to reduce background fluorescence in the light pass between a dichroic mirror of 505 nm and an emission filter of 535/45 nm band pass. The cells transiently expressing GFP-tagged PKCs were loaded with 2 µM Fura2/AM in the standard extracellular solution for 30 min at room temperature. The cells were washed twice and used within 2 h. The Fura2 ratio was calibrated using exposure to 10 µM ionomycin and 10 mM Ca2+ or 10 mM EGTA in the Fura2-loaded cells without transfection of the GFP-tagged PKCs. A dissociation constant of 150 nM for Ca2+ and Fura2 at room temperature was used. For simultaneous measurement of relative fluorescence change in intensity of MARCKS-GFP and PKCalpha -DsRed using a dual band for fluorescein isothiocyanate and TRITC, GFP fluorescence was excited at 488 nm, whereas PKCalpha -DsRed fluorescence was excited at 558 nm. To reduce cross-talk between them, an emission filter wheel was used, and alternate emission filters of 535/45-nm band pass and 605/50-nm long pass were synchronously set with excitation filters for GFP and DsRed.

TIRFM (Evanescent Wave)-- To obtain high signal-to-noise ratio over the conventional epifluorescence microscopy (see supplemental figure), we installed a TIRFM unit (Olympus) into the same imaging system mentioned above. The incidental light was introduced from the objective lens for TIRFM (Olympus NA = 1.45, 60×). GFP and Fura Red were excited by a 488-nm laser, and each emitted light was collected through 535/45 and 645/75 nm, respectively. For simultaneous measurement of relative fluorescence change in intensity of C12-GFP and [Ca2+]i in the Fura Red-loaded cells, we used a W-view Optics (Hamamatsu Photonics), a branching optics that splits the incident light into using a Dichroic mirror of 550 nm, so that two separate images of the GFP and Fura Red fluorescence can be produced.

    RESULTS
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Membrane Depolarization Induces Transient Translocation of PKCalpha Following [Ca2+]i Elevation-- We first examined the distribution of PKCalpha -GFP with the help of high time resolution digital imaging. Fig. 1A shows the rapid and reversible translocation of PKCalpha -GFP in response to a depolarizing K+ concentration (40 mM), which evoked Ca2+ influx through opening of VDCCs. The relative changes in the fluorescence intensities of PKCalpha -GFP in the cytosol and at the plasma membrane are plotted in Fig. 1B, as a function of time. PKCalpha -GFP translocated from the cytosol to the plasma membrane, and this can be seen by the reciprocal changes in the two parameters (Fig. 1B) (n = 6). Thus, either parameter can be used as a marker of PKCalpha -GFP plasma membrane translocation. We chose to employ the relative fluorescence change in the cytosol as a marker of translocation. Next, we simultaneously measured [Ca2+]i and PKCalpha translocation in Fura2-loaded and PKCalpha -GFP-expressing INS-1 cells. As seen in Fig. 1C, a depolarizing K+ concentration induced a transient translocation of PKCalpha -GFP to the plasma membrane following a transient [Ca2+]i elevation (n = 8). Thus, the temporal profile was similar to that observed when PKCalpha -GFP measurements were carried out alone, suggesting successful dissection of Ca2+ and GFP signals.


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Fig. 1.   Depolarization-evoked Ca2+ influx induces translocation of PKCalpha in PKCalpha -GFP transiently expressing INS-1 cells. A, images of PKCalpha -GFP translocation induced by 40 mM K+ at high time resolution (sampling interval, 0.25 s). Panels a-e were taken at the times indicated by the arrows in B. The bar represents 10 µm. Maximal translocation was observed at 2 s (panel c) after stimulation. The subcellular distribution of PKCalpha -GFP in panel a was indistinguishable from that in panel e. B, the entire time course of PKCalpha -GFP translocation induced by the 40 mM K+ solution. The regions of interests were at the plasma membrane (PM) and in the cytosol (white boxes in A). C, simultaneous measurements of [Ca2+]i and PKCalpha translocation in Fura2-loaded and PKCalpha -GFP-expressing INS-1 cells. ratio (PKCcyt) represents the relative fluorescence change in intensity of PKCalpha -GFP in the cytosol. The translocation of PKCalpha -GFP had ceased (back to the prestimulation level) before [Ca2+]i returned to the resting level.

Threshold Value of [Ca2+]i for PKCalpha Translocation-- We loaded INS-1 cells with Fura2, without expressing PKCalpha -GFP, to accurately estimate [Ca2+]i. In the standard extracellular solution containing 2.5 mM CaCl2 and 3 mM glucose, more than 50% of the Fura2-loaded INS-1 cells displayed spontaneous cytosolic Ca2+ oscillations (20). The peak [Ca2+]i was no more than 400 nM. When the same cells were depolarized by the K+ channel blocker tetraetheylammonium (TEA) (20 mM), the Ca2+ oscillations became more pronounced (21), and the peak [Ca2+]i was in the range of 600-800 nM (Fig. 2A) (n = 10). To avoid cross-talk between Ca2+ and GFP signals, exactly the same protocol as in Fig. 2A was applied to PKCalpha -GFP-expressing cells without Fura2 loading. No translocation of PKCalpha -GFP took place in the standard extracellular solution, whereas oscillatory translocations of PKCalpha -GFP started immediately after introduction of the TEA-containing solution (Fig. 2B) (n = 5), suggesting a threshold value of [Ca2+]i of more than 400 nM for PKCalpha translocation. The temporal profile of the TEA-evoked PKCalpha -GFP translocations (Fig. 2, C and D) (n = 5) was similar to that observed in response to the membrane depolarization evoked by a high K+ concentration (Fig. 1B).


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Fig. 2.   Threshold value of [Ca2+]i for translocation of PKCalpha in INS-1cells. A, spontaneous and TEA-evoked Ca2+ oscillations. Three different cells displayed spontaneous oscillatory changes in [Ca2+]i under 400 nM. The amplitude of the Ca2+ oscillations became larger (>400 nM) during TEA stimulation. B, two PKCalpha -GFP expressing cells, not loaded with Fura2, showed oscillatory translocations of PKCalpha -GFP in the presence of TEA, but not in the standard solution. C, images of PKCalpha -GFP translocation induced by 20 mM TEA at high time resolution (sampling interval, 0.25 s). Panels a-e were taken at the times indicated by the arrows in D. the bar represents 10 µm. Maximal translocation was observed 2 s (panel d) after stimulation. The subcellular distribution of PKCalpha -GFP in panel a was indistinguishable from that in panel e. D, the entire time course of PKCalpha -GFP translocation. The regions of interests were at the plasma membrane (PM) and in the cytosol (white boxes in C).

Depolarization-evoked Translocation of PKCalpha Depends on Ca2+ Influx but Not on Ca2+ Mobilization-- Fig. 3A shows that upon removal of external Ca2+, the TEA-induced Ca2+ oscillations and the PKCalpha -GFP translocations were abolished (n = 5), indicating that both are totally dependent on Ca2+ influx. We then tested whether PKCalpha can be fully activated in the presence of DAG at physiological Ca2+ concentrations. Using PKCalpha -GFP translocation to the plasma membrane as a marker of activation, short exposure to a combination of TEA and the diacylglycerol analogue DiC8 (100 µM) induced sustained PKCalpha activation, despite the fact that [Ca2+]i quickly returned to the steady resting level upon removal of the stimulation (Fig. 3B) (n = 8). This suggests that even a single TEA-evoked Ca2+ spike (within the physiological Ca2+ concentration range) can fully activate PKCalpha in the presence of a sufficient amount of DAG (8).


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Fig. 3.   Translocation of PKCalpha is dependent on Ca2+ influx through VDCCs. Simultaneous monitoring of the translocation of PKCalpha -GFP and [Ca2+]i in Fura2-loaded cells with expression of PKCalpha -GFP is shown. ratio (PKCcyt) represents relative fluorescence change in intensity of PKCalpha -GFP in the cytosol. A, repetitive translocations of PKCalpha -GFP were synchronous with Ca2+ oscillations in the presence of TEA. Both were abolished upon removal of external Ca2+. B, combined application of 100 µM DiC8 and TEA resulted in sustained translocation of PKCalpha -GFP, although [Ca2+]i had returned to the prestimulation level upon removal of the stimulation.

To compare the effects of Ca2+ influx and IP3-mediated Ca2+ mobilization on PKCalpha -GFP translocation, we tested, in the same cells, the actions of TEA and acetylcholine (ACh) (using a supramaximal concentration (100 µM), which might produce sufficient DAG to activate PKCalpha (see Figs. 7B and 9A)). As shown in Fig. 4A, the TEA-evoked translocation of PKCalpha -GFP was much more substantial than that induced by ACh, although the bulk [Ca2+]i elevations produced by the two agents were nearly equal (n = 11). For more accurate evaluation of the cytosolic Ca2+ elevation caused specifically by IP3-induced store release, we removed the Ca2+ influx component due to capacitative Ca2+ entry through store-operated Ca2+ channels in the plasma membrane (22, 23). This was simply done by removing external Ca2+ during stimulation with ACh (100 µM) (Fig. 4B). The result was similar (n = 5) to that shown in Fig. 4A, suggesting that there was little effect of store-operated Ca2+ influx on PKCalpha translocation. Reversing the sequence of events (applying TEA first and then subsequently ACh) gave similar results to those shown in Fig. 4 (data not shown).


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Fig. 4.   Ca2+ influx through VDCCs, but not Ca2+ mobilization induced by ACh, is important for translocation of PKCalpha . Simultaneous monitoring of the translocation of PKCalpha -GFP and [Ca2+]i in Fura2-loaded cells with expression of PKCalpha -GFP is shown. A, a supramaximal concentration of ACh (100 µM) induced a small translocation of PKCalpha -GFP, whereas TEA-evoked depolarization induced a marked PKCalpha -GFP translocation, although the magnitude of the [Ca2+]i elevation induced by ACh was virtually the same as that elicited by TEA and lasted longer. B, the same concentration of ACh (100 µM) in the absence of external Ca2+ also induced a very small translocation. It should be noted that the third TEA-evoked translocation was much larger than the second ACh-elicited translocation.

PKCalpha , Activated by Depolarization-evoked Ca2+ Influx, Can Phosphorylate Its Substrate, MARCKS-- One line of evidence has shown that [Ca2+]i elevations drive translocation of cPKC to the plasma membrane (13, 14). However, we do not know whether cPKC can be activated by depolarization-evoked Ca2+ influx alone. To answer this question, we employed a GFP-tagged MARCKS as another marker of PKC activity (17), which is a putative and direct substrate for PKC (16), as well as PKCalpha -DsRed. We co-transfected both of them into INS-1 cells. When activated PKC phosphorylates the plasma membrane-anchored MARCKS, then phosphorylated MARCKS translocates from the plasma membrane to the cytosol. Thus, simultaneous monitoring of PKCalpha -DsRed and MARCKS-GFP allows us to test whether depolarization-evoked Ca2+ influx can activate cPKC. Fig. 5 (A and B) shows translocations of MARCKS-GFP and PKCalpha -DsRed induced by TEA, indicating that depolarization-evoked Ca2+ influx can activate PKCalpha . Phosphorylated MARCKS only slowly and gradually returned to the plasma membrane (~2.5 min) in contrast to the rapid temporal profile of PKCalpha translocation (~30 s) (Fig. 5B) (n = 6). Ca2+ oscillation-driven translocations of PKCalpha kept MARCKS phosphorylated in the cytosol until termination of the repetitive PKCalpha translocations (Fig. 5C) (n = 8).


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Fig. 5.   Ca2+ oscillation-driven translocation of PKCalpha -DsRed results in phosphorylation of MARCKS-GFP. Simultaneous monitoring of the translocations of PKCalpha -DsRed and MARCKS-GFP is shown. A, superimposed panels a-f were taken at the times indicated by the arrows in B. The bar represents 10 µm. The regions of interests were at the plasma membrane and in the cytosol (white box). Panel a, MARCKS-GFP (green) anchored at the plasma membrane, whereas PKCalpha -DsRed (red) was in the cytosol. A short exposure to TEA resulted in a dramatic reverse change in the red and green colors; MARCKS that had been phosphorylated was translocated to the cytosol as soon as PKCalpha translocation to the plasma membrane had taken place (panels b and c). MARCKS was still in the cytosol as merged color of yellow can be seen in panels d and e. The last image (panel f) shows the same distribution of PKCalpha -DsRed and MARCKS-GFP as in panel a. B, the entire time course of the translocations of PKCalpha -DsRed and MARCKS-GFP shown in A. C, trains of oscillatory PKCalpha -DsRed translocations kept MARCKS in the cytosol. The ratio represents relative fluorescence change in intensity of PKCalpha -DsRed and MARCKS-GFP at the same region of interest in the cytosol (B and C).

Depolarization-evoked Ca2+ Influx Induces Translocation of PKCtheta , Despite the Absence of the Functional C2 Domain for Ca2+ Binding-- As seen in Fig. 5 (A and B), we now know that depolarization-evoked Ca2+ influx can activate PKCalpha . This observation prompted us to explore whether depolarization-evoked Ca2+ influx can generate DAG, because Ca2+ and DAG are required for activation of cPKC (2). It has been shown that K+-induced membrane depolarization increases IP3 production in insulin-secreting rat pancreatic islets (9). Taken together, these observations indicate that there should be production of DAG in response to depolarization-evoked Ca2+ influx. To test this hypothesis, we employed PKCtheta -GFP as a marker of nPKC activity as well as DAG production because nPKC is activated by DAG alone in a Ca2+-independent manner (2). TPA caused a rapid and sustained translocation of PKCtheta -GFP in the complete absence of Ca2+ influx (Fig. 6, A and B) (n = 6). The simultaneous measurements of PKCtheta -GFP translocation and cytosolic Ca2+ concentration, shown in Fig. 6C, confirm that extracellular Ca2+ is not required for PKCtheta -GFP translocation (n = 4). We then applied a depolarizing K+ concentration to PKCtheta -GFP-expressing cells. Fig. 7A clearly shows that this stimulus induced a gradual translocation of PKCtheta -GFP to the plasma membrane, which was reversible upon removal of the high K+ solution (n = 12). The amplitude of the translocation induced by K+-induced depolarization was comparable with that elicited by ACh (Fig. 7B). It should be noted that ACh still continued to induce translocation of PKCtheta -GFP after [Ca2+]i had returned to the resting level, verifying that 100 µM ACh generated enough DAG to sustain the translocation until termination of the stimulus (n = 8) (Figs. 7B and 9B). The amplitude of the TEA-evoked translocation was also comparable with that induced by ACh, and the translocation was not synchronous with the TEA-evoked Ca2+ spikes (Fig. 7C) (n = 8).


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Fig. 6.   PKCtheta -GFP translocation as a representative of nPKC is induced by TPA in the absence of Ca2+. A and B, images of PKCtheta -GFP translocation induced by 100 nM TPA at high time resolution (sampling interval, 0.25 s) and the entire time course of the PKCtheta -GFP translocation. The regions of interests were at the plasma membrane (PM) and in the cytosol (white boxes in B). The panels a-e in B were taken at the times indicated by the arrows in A. The bar in B represents 10 µm. C, simultaneous monitoring of PKCtheta -GFP translocation and [Ca2+]i in Fura2-loaded cells with expression of PKCtheta -GFP. ratio (PKCcyt) represents relative fluorescence change in intensity of PKCtheta -GFP in the cytosol. Translocation of PKCtheta -GFP immediately started following application of 100 nM TPA in the absence of external Ca2+, whereas spontaneous Ca2+ oscillations had no effect on the translocation. Simultaneous monitoring of PKCtheta -GFP translocation and [Ca2+]i in Fura2-loaded cells with expression of PKCtheta -GFP (A-C).


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Fig. 7.   Depolarization-evoked Ca2+ influx and ACh induce PKCtheta -GFP translocation. Simultaneous monitoring of PKCtheta -GFP translocation and [Ca2+]i in Fura2-loaded cells with expression of PKCtheta -GFP is shown. ratio (PKCcyt) represents relative fluorescence change in intensity of PKCtheta -GFP in the cytosol. A, relatively slow translocation of PKCtheta -GFP induced by a depolarizing K+ concentration (40 mM). PKCtheta -GFP continued to be translocated until cessation of the stimulation. Thereafter, PKCtheta -GFP started to return to the cytosol. At the same time, [Ca2+]i started to decrease toward the prestimulation level. B, ACh gradually and continuously translocated PKCtheta -GFP until removal of ACh, when [Ca2+]i had already returned to the prestimulation level. C, TEA-evoked repetitive Ca2+ spikes also induced translocation of PKCtheta -GFP, with a temporal profile similar to that induced by ACh.

Depolarization-evoked Ca2+ Influx Translocates GFP-tagged Pleckstrin Homology Domain of PLCdelta 1 (GFP-PHD) and GFP-tagged C1 Domain of PKCgamma (C12-GFP)-- To fully corroborate the above evidence that the Ca2+ influx generates DAG and activates PKCtheta , we performed further experiments using GFP-PHD and C12-GFP. First, GFP-PHD allows us to visualize IP3 production by translocating from the plasma membrane to the cytosol because of the 20-fold higher affinity for IP3 than for PIP2 (19), such that we can assess indirectly the simultaneous production of DAG upon PIP2 hydrolysis by a Ca2+-dependent PLC activation. As shown in Fig. 8A, depolarization-evoked Ca2+ influx resulted in the relatively transient translocation of PHD-GFP, whereas the translocation was sustained during ACh stimulation (n = 11), indicating DAG production. Second, to directly monitor the plasma membrane DAG levels, we also employed translocation of the C12-GFP as a DAG sensor (14, 15) using TIRFM. This has a ~10-fold higher signal-to-noise ratio (see "Experimental Procedures") than conventional epifluorescence microscopy in terms of the fluorescent protein translocation near the plasma membrane (for example PKC-GFPs) (15). We loaded the C12-GFP-expressing cells with Fura Red to define the relationship between the [Ca2+]i change and DAG production in response to the Ca2+ influx. Depolarization-evoked Ca2+ influx clearly translocated C12-GFP to the membrane (Fig. 9A) after [Ca2+]i had risen to the peak (Fig. 9B), in contrast to the sustained ACh-induced translocation of C12-GFP following the transient [Ca2+]i elevation. Stimulation with 40 mM K+ also resulted in a C12 translocation of an undiminished magnitude in 5 µM thapsigargin-pretreated C12-GFP-expressing cells, indicating little contribution of either Ca2+- or IP3-induced Ca2+ release to the C12 translocation (data not shown). As long as both high K+ and ACh stimulation continued, C12-GFP remained at the plasma membrane. The translocation induced by Ca2+ influx was inhibited by the PLC inhibitor U73122 (n = 4), suggesting that DAG production is mediated by Ca2+-dependent PLC activation (Fig. 9D).


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Fig. 8.   Depolarization-evoked Ca2+ influx and ACh induce GFP-PHD translocation. A, the entire time course of GFP-PHD translocation induced by the high K+ solution (40 mM) and 100 µM ACh. The regions of interests were at the plasma membrane (PM) and in the cytosol (white boxes in B). ratio (PKCcyt) represents relative fluorescence change in intensity of GFP-PHD in the cytosol. GFP-PHD translocation was transient with stimulation of high potassium, whereas ACh continued to translocate GFP-PHD until the removal of ACh. B, panels a-d were taken at the times indicated by the arrows in A. The bar represents 10 µm.


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Fig. 9.   Depolarization-evoked Ca2+ influx and ACh induce C12-GFP translocation. Shown are simultaneous measurements of [Ca2+]i and C12-GFP translocation in Fura Red-loaded and C12-GFP expressing INS-1 cells using TIRFM. The ratio represents the relative fluorescence change in intensity of C12-GFP at the plasma membrane and of [Ca2+]i. Note that rises in Ca2+ are reported as a fall in fluorescence. The region of interest for both parameters was at the plasma membrane (white box in C). A, the entire time course of simultaneous measurements of [Ca2+]i and C12-GFP translocation. C12-GFP translocation to plasma membrane induced by both high potassium and ACh has biphasic pattern. B, areas denoted by the dashed box in A are expanded. Translocation of C1-GFP started 9 s after the [Ca2+]i elevation. C, panels a-c were taken at the times indicated by the arrows in A. The bar represents 10 µm. D, effect of U73122 on C12-GFP translocation induced by high potassium. The cell was pretreated with U73122 for 15 min following the control experiment with 40 mM K+ stimulation. U73122 almost completely inhibited C12-GFP translocation (red) induced by high potassium stimulation in comparison with the control experiment (blue).

In Situ Calibration of Depolarization-evoked Increase in DAG Content-- We calibrated the depolarization-evoked increase in DAG concentration in single C12-GFP expressing cells. An extracellular solution containing the DAG analogue DiC8 was introduced at the end of each experiment, following 40 mM K+ stimulation, using TIRFM. Fig. 10 shows a representative experiment where application of three different concentrations of DiC8 (1, 3 and 10 µM) resulted in different quasi-steady state levels of C12 translocation. Because the concentration of DiC8 inside the cell, at each level, is thought to equilibrate with that outside the cell, the depolarization-evoked increase in DAG concentration was estimated (from the calibration curves of the three experiments) to be 1.90 ± 0.02 µM (mean ± S.D.).


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Fig. 10.   Calibration of depolarization-evoked increase in DAG concentration in C12-GFP expressing cells using TIRFM. A, cells were first treated with 40 mM K+ to establish the magnitude of the translocation of C12-GFP. Thereafter, the cells were perfused with the Ca2+-free extracellular solution containing 0.2 mM EGTA for the first 2 min followed by consecutive applications of DiC8 1 µM (0.35 µg/ml), 3 µM (1.05 µg/ml), and 10 µM (3.5 µg/ml). B, panels a-e were taken at the times indicated by the arrows in A. The region of interest represented by the dashed box was at the plasma membrane. The ratio represents relative fluorescence intensity of C12-GFP. The bar represents 10 µm.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Microdomains of Elevated [Ca2+] beneath the Plasma Membrane, but Not Elevation of the Bulk [Ca2+]i, Are Required for cPKC Translocation-- We have shown that there is a threshold value of the bulk [Ca2+]i at ~400 nM for PKCalpha translocation in the insulin-secreting INS-1 cells (Fig. 2A), which is consistent with data from other laboratories (24). More importantly, we have also demonstrated that Ca2+ influx is a much stronger stimulus for PKCalpha translocation than Ca2+ mobilization from intracellular stores, even when the amplitudes of the induced bulk [Ca2+]i elevations are similar (Fig. 4A). This finding suggests that microdomains of elevated [Ca2+] beneath the plasma membrane ([Ca2+]sub) generated by Ca2+ influx through VDCCs may play a pivotal role in cPKC translocation in excitable cells rather than the elevated bulk [Ca2+]i, which is also consistent with a recent report (25). In other words, there is a threshold value of [Ca2+]sub for translocation of cPKC. In neuroendocrine cells, the estimated value of [Ca2+]sub at the mouth of open VDCCs is several micromoles/liter (12, 26). This indicates that a local [Ca2+] of several micromoles/liter may be required for cPKC translocation.

Ca2+ mobilization induced by ACh most likely fails to translocate PKCalpha because the [Ca2+] rise at the critical sites is insufficient. This could be due to the distance between the plasma membrane and the IP3 channels in the ER, combined with the substantial Ca2+ buffering capacity in the cytosol (27). However, it could be argued that Ca2+ mobilization from the ER should result in store-operated Ca2+ entry (22, 23) and that ACh stimulation therefore also could be expected to cause local Ca2+ elevation beneath the plasma membrane. Nevertheless, it would appear (Fig. 4B) that the magnitude of Ca2+ influx through store-operated Ca2+ channels is insufficient to generate the threshold level of [Ca2+]sub needed. Because the entry sites from store-operated Ca2+ channels would be very close to Ca2+ uptake sites into the ER through the powerful Ca2+ ATPase pumps (23, 28-31), the net delivery of Ca2+ to the cytosol through store-operated Ca2+ channels may be less than through voltage-gated channels even if both channels have similar ranges of Ca2+ concentrations at the mouths of their pores. They could also possibly be separately located. Our finding that Ca2+ entry through VDCCs is sufficient to cause cPKC and nPKC translocation may be important in relation to the control of glucose-elicited insulin secretion, because it has been shown that the L-type VDCCs and the insulin-containing secretory granules are co-localized (26). Thus, local Ca2+ entry in the secretory domains could induce PKC activation important for stimulation of exocytosis (25).

Ca2+ Influx through VDCCs Is Both Necessary and Sufficient for Activation of cPKC-- To fully substantiate that Ca2+ influx through VDCCs is both necessary and sufficient for activation of cPKC, we employed the phosphorylation state of MARCKS as another marker of PKC activity. As seen in Fig. 5B, plasma membrane-anchored MARCKS is turned into phosphorylated MARCKS, thereby moving into the cytosol, as soon as PKCalpha is translocated to the plasma membrane by Ca2+ influx through VDCCs. In INS-1 cells, in which PKCalpha and PKCepsilon are predominantly expressed (32), endogenous cPKC and nPKC may move to the plasma membrane in the same manner as the exogenous examples. Thus, we have provided the first direct evidence showing that single Ca2+ spike-driven translocations of cPKC, whose duration is just 30 s long, enable MARCKS to be phosphorylated. The pseudosubstrate inhibitory region of cPKC has been already removed before the association with MARCKS. The cessation of MARCKS translocation and the return to the prestimulation level is very much slower than that of the cPKC translocation, because of sustained MARCKS phosphorylation. We do not know the exact mechanism, but it might result from the net effect of several factors such as diacylglycerol kinase (17), PKC, or a phosphatase that dephosphorylates MARCKS.

Depolarization-evoked Ca2+ Influx through VDCCs Can Generate DAG and Thereby Activate cPKC and nPKC, Whose Activation Is Structurally Independent of Ca2+-- nPKC, which lacks the functional C2 domain for Ca2+ binding, is activated either by DAG or TPA. This can be seen by the sustained PKCtheta translocation induced by TPA in the absence of external Ca2+ (Fig. 6B). However, depolarization-evoked Ca2+ influx through VDCCs can also induce gradual and continuous nPKC translocation to the plasma membrane during [Ca2+]i elevation (Fig. 7A). It is possible that some regions of PKCtheta other than the C1 domain can be associated with the plasma membrane. However, two additional experiments using GFP-PHD and C12-GFP have added further credence to the observations (Fig. 7). First, the Ca2+ influx-evoked translocation of GFP-PHD (Fig. 8A) indicates that DAG can be generated upon PIP2 hydrolysis mediated by a Ca2+-dependent PLC activation (33), although the amplitude of the PHD translocation may parallel the concentration not of DAG but of IP3 (19). The translocation of GFP-PHD induced by depolarization was relatively transient, whereas it was sustained during ACh stimulation. This suggests a relatively transient increase in DAG concentration (3). Second, the Ca2+ influx-evoked translocation of C12-GFP (Fig. 9A) as a DAG sensor, directly supports the view that DAG synthesis is induced by depolarization-evoked Ca2+ influx through VDCCs. The simplest explanation for this surprising observation could be that the Ca2+ influx can initiate DAG generation, by triggering PLC activation, thereby translocating and activating nPKC. The fact that the translocation of the C1 domain did not start until the [Ca2+]i had nearly reached its peak (Fig. 9B), taken together with the observation that sustained Ca2+ influx induced by 1 µM ionomycin kept C12-GFP at the plasma membrane (data not shown), suggests that there may be a threshold value of [Ca2+]sub for DAG synthesis. Conversely, DAG synthesis can be detected by monitoring the C1 domain translocation, if the amplitude of the C1 domain translocation parallels the amount of DAG synthesis. Therefore, we tried to estimate the increase in DAG content with in situ calibration (Fig. 10A), which gave a value of 1.90 ± 0.02 µM (mean ± S.D., n = 3). In a report from another laboratory, using a biochemical assay, it was calculated that the amount of accumulated DAG is 13 pmol/106 cells at 30 s in platelet-derived growth factor-stimulated Balb/c/3T3 cells (34). Given a cell volume of ~1 pl, the DAG concentration would be 13 µM, which is comparable with our data. As shown in Fig. 5, a DAG concentration of ~2 µM, induced by depolarization-evoked Ca2+ influx, may be sufficient to ensure that activated PKC can phosphorylate MARCKS. Monitoring of C12-GFP translocation has been the most sensitive way of detecting DAG synthesis beneath the plasma membrane so far (14, 15). As shown in Fig. 9A, the C1 domain biphasically translocated to the membrane during high K+ stimulation; the first phase was transient, and the second phase was sustained. This indicates continuous production of DAG. DAG synthesis can be mediated by PLC (Figs. 8A and 9D) (33) and/or phospholipase D activated by the [Ca2+]i rise and/or PKCs (3, 9, 35, 36). However, neither Ca2+- nor IP3-induced Ca2+ mobilization from the Ca2+ stores is important for DAG synthesis, because of the undiminished magnitude of the depolarization-evoked C12 translocation in the thapsigargin-pretreated cells.

Our data also have important implications for cPKC activation. If the amplitude of the nPKC translocation reflects the amount of DAG synthesized, depolarization-evoked Ca2+ influx could translocate as well as activate cPKC by generating [Ca2+]sub and DAG. Ca2+ signals per se, such as action potential-induced Ca2+ oscillations, could function as second messengers as well as operate as primary activators of cPKC and nPKC in neuronal, endocrine and muscle cells. In other words, Ca2+ signals and the two PKCs signals may not be segregated in certain conditions. Therefore, the roles of these signals in a myriad of cellular functions may overlap.

Short-lived Ca2+ Signals through PKC Activation Are Transduced into Long-lived Phosphorylated MARCKS; Ca2+ Oscillation-driven Activation of cPKC and nPKC May Modulate Long Term Physiological Phenomena-- Our finding that Ca2+ oscillation-evoked activation of both cPKC and nPKC can keep MARCKS phosphorylated (Fig. 5) has important implications for the control of long term physiological phenomena such as insulin secretion (37), long term potentiation (38), the redox state in the mitochondria (39), and the control of gene expression (40, 41). We can envisage that, as long as Ca2+oscillation-driven activation of a first kinase such as PKC continues in a "sinus-like" manner, then the first kinase is maintained in the phosphorylated state, which in turn leads to activation of a second kinase on a time scale of hours or days. In this way, not only Ca2+ oscillations but also Ca2+ oscillation-driven activation of both PKCs may modulate a long term physiological phenomenon. Thus, we should bear in mind that two classes of PKCs can be activated in conditions where Ca2+ oscillations take place.

    ACKNOWLEDGEMENTS

GFP-PHD was a kind gift from Dr. Hirose (Tokyo University). We thank Dr. Alexei Tepikin for critical reading of the manuscript.

    FOOTNOTES

* This work was supported by grants-in-aid for Scientific Research from the Ministry of Education, Science, Sports and Culture of Japan and grants from Takeda Science Foundation, the Yamanouchi Foundation for Research on Metabolic Disorders, the Naito Foundation, the Nissan Science Foundation, and the Suzuken Memorial Foundation. This work was also supported by a Program Grant from the Medical Research Council (United Kingdom).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The on-line version of this article (available at http://www.jbc.org) contains a supplemental figure.

§ To whom correspondence should be addressed. Tel.: 81-53-435-2249; Fax: 81-53-435-7020; E-mail: hmogami@hama-med.ac.jp.

Dagger Dagger MRC Research Professor.

Published, JBC Papers in Press, January 3, 2003, DOI 10.1074/jbc.M210653200

    ABBREVIATIONS

The abbreviations used are: PKC, protein kinase C; cPKC, conventional PKC; nPKC, novel PKC; DAG, diacylglycerol; PIP2, phophatidylinositol 4,5-bisphosphate; PLC, phospholipase C; VDCC, voltage-dependent Ca2+ channel; IP3, inositol 1,4,5-trisphosphate; GFP, green fluorescent protein; MARCKS, myristoylated alanine-rich C kinase substrate; -GFP, GFP-tagged; -DsRed, DsRed-tagged; PHD, pleckstrin homology domain; TPA, 12-O-tetradecanoylphorbol-13-acetate; DiC8, 1,2-dioctyl-sn-glycerol; TRITC, tetramethylrhodamine isothiocyanate; TIRFM, total internal reflection fluorescence microscopy; TEA, tetraetheylammonium; Ach, acetylcholine; ER, endoplasmic reticulum.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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