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Originally published In Press as doi:10.1074/jbc.M702746200 on September 12, 2007

J. Biol. Chem., Vol. 282, Issue 45, 32710-32718, November 9, 2007
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Temperature-dependent Modulation of CaV3 T-type Calcium Channels by Protein Kinases C and A in Mammalian Cells*Formula

Jean Chemin{ddagger}1, Alexandre Mezghrani{ddagger}, Isabelle Bidaud{ddagger}2, Sebastien Dupasquier§3, Fabrice Marger{ddagger}4, Christian Barrère{ddagger}, Joël Nargeot{ddagger}, and Philippe Lory{ddagger}

From the {ddagger}Département de Physiologie and the §Département d'Oncologie Moléculaire et Cellulaire, Institut de Génomique Fonctionnelle, CNRS UMR 5203, INSERM U661, Universités de Montpellier 1 et 2, Institut Fédératif de Recherche 3, Montpellier 34094, France

Received for publication, March 30, 2007 , and in revised form, September 12, 2007.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Modulation of low voltage-activated CaV3 T-type calcium channels remains poorly characterized compared with high voltage-activated CaV1 and CaV2 calcium channels. Notably, it is yet unresolved whether CaV3 channels are modulated by protein kinases in mammalian cells. In this study, we demonstrate that protein kinase A (PKA) and PKC (but not PKG) activation induces a potent increase in CaV3.1, CaV3.2, and CaV3.3 currents in various mammalian cell lines. Notably, we show that protein kinase effects occur at physiological temperature (~30–37 °C) but not at room temperature (~22–27 °C). This temperature dependence could involve kinase translocation, which is impaired at room temperature. A similar temperature dependence was observed for PKC-mediated increase in high voltage-activated CaV2.3 currents. We also report that neither CaV3 surface expression nor T-current macroscopic properties are modified upon kinase activation. In addition, we provide evidence for the direct phosphorylation of CaV3.2 channels by PKA in in vitro assays. Overall, our results clearly establish the role of PKA and PKC in the modulation of CaV3 T-channels and further highlight the key role of the physiological temperature in the effects described.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Voltage-gated Ca2+ channels (VGCCs)5 are unique among voltage-gated ion channels because the permeant Ca2+ ion also acts as an intracellular second messenger, triggering diverse cellular functions (1, 2). VGCCs are therefore implicated in neuronal and cardiac excitability as well as in muscle contraction, neurotransmitter release or hormone secretion, and gene expression (1–6). Thus, the modulation of VGCCs plays a pivotal role in the control of cardiac and brain activities.

VGCCs are divided into three families: the L-type channels (CaV1 family); the neuronal N-, P/Q-, and R-type channels (CaV2 family); and the T-type channels (CaV3 family) (7). Although the molecular mechanisms implicated in the modulation of high voltage-activated Ca2+ channels of the CaV1 and CaV2 families are beginning to be unraveled (mainly, protein kinases for CaV1 channels and beta{gamma}-subunits of G proteins and protein kinase C (PKC) for CaV2 channels) (6, 8), those implicated in low voltage-activated CaV3 T-type channel regulation remain debated (9). Some transduction pathways mediate either decreases or increases in native T-currents, depending on the tissues/species studied and/or on the recording conditions (9). This apparent complexity could be explained by the existence of three T-channels (CaV3.1 or {alpha}1G, CaV3.2 or {alpha}1H, and CaV3.3 or {alpha}1I), which include different splice variant isoforms with specific tissue patterns and developmental expression (10). Additional regulatory subunits of T-channels might also be involved in their modulation, but the native composition of T-channels remains unknown (10). Therefore, molecular studies on recombinant T-channels are required to clarify their modulation.

Recent studies on recombinant T-channels have highlighted the complexity of their regulation (9). On one hand, as for CaV2 channels, CaV3.2 currents are inhibited by beta{gamma}-subunits of G proteins in mammalian cells (11, 12). In contrast with CaV2 currents, this mechanism cannot be generalized because this modulation is restricted to CaV3.2 and specifically occurs with a Gbeta{gamma} dimer containing beta2-subunits (11, 12). On the other hand, as observed with CaV1 channels, CaV3 currents are increased by protein kinase activation (13–18). Yet again, kinase effects on T-currents appear complex (9) because protein kinase A (PKA)- and PKC-mediated T-current increase has been observed in Xenopus oocytes but not in mammalian cells expressing CaV3 currents (11, 16–19). In this context, it is interesting to note that protein kinase effects on native VGCCs were first described in heart frog cells and appear more robust in these cells compared with those observed in mammalian tissues (20–23). Thus, we reasoned that these differences in kinase modulation could involve the temperature at which the experiments were performed, i.e. mostly at room temperature. Although room temperature is within the physiological range for amphibian cells, it is far below that required by mammalian cells. In this study, we provide new data on PKA and PKC activation in the function of temperature that clearly establish the role of these two kinases, but not of protein kinase G (PKG), in the modulation of CaV3 T-currents in various mammalian cell lines.


Figure 1
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  		FIGURE 1.
PMA induces an increase in CaV3 T-currents in transiently transfected tsA-201 cells at 37 °C but not at room temperature (~22 °C). A and B, effect of 100 nM PMA or the control solution (Ctrl) on CaV3.1 currents when incubated for 10 min at 37 °C and at room temperature (RT), respectively, before electrophysiological experiments, which were performed at room temperature during the following 50 min in the presence of PMA or the control solution (see protocols in insets). C and D, summary of the data obtained for CaV3.1, CaV3.2, and CaV3.3 currents upon incubation of PMA at 37 °C and at room temperature, respectively. Currents were elicited by a –45-mV depolarization (200-ms duration for CaV3.1 and CaV3.2 currents and 450-ms duration for CaV3.3 currents) applied immediately after the whole-cell configuration from a holding potential of –80 mV. ***, p < 0.001.

 

   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture and Transfection Protocols—tsA-201 cells and a Chinese hamster ovary (CHO) cell line stably expressing CaV3.2 channels (CHO-CaV3.2; a generous gift from Dr. Emmanuel Bourinet) were cultivated in Dulbecco's modified Eagle's medium and Ham's F-12 medium, respectively, supplemented with GlutaMAX and 10% fetal bovine serum (Invitrogen). Neomycin (0.6 mg/ml; Invitrogen) was added to the CHO-CaV3.2 cell medium. tsA-201 cell transfection was performed using jetPEI (Qbiogen, Inc.) according to the manufacturer's protocol (4 µl of jetPEI for ~1.5 µg of DNA/35-mm Petri dish) with a DNA mixture containing 0.5% of a green fluorescent protein (GFP) plasmid and 99.5% of one of the pcDNA3 plasmid constructs that code for the human CaV3.1a, CaV3.2, and CaV3.3 T-channel isoforms (24). CaV2.1-hemagglutinin (HA) (25) and CaV2.3 (rat brain E-II) (26) were transfected under the same conditions with a mixture containing the {alpha}2{delta}1- and beta4-subunits at a 2:1:1 ratio. Two days after, cells were then dissociated with Versen (Invitrogen) and plated at ~35 x 103 cells/35-mm Petri dish. Electrophysiological recordings were performed the following day.

Electrophysiological Recordings—Macroscopic currents were recorded at room temperature (~22 °C) by the whole-cell patch clamp technique using an Axopatch 200B amplifier (Axon Instruments). The extracellular solution contained 135 mM NaCl, 20 mM tetraethylammonium chloride, 2 mM CaCl2, 1 mM MgCl2, and 10 mM HEPES (pH adjusted to 7.4 with KOH, ~330 mosM). Borosilicate glass pipettes have a typical resistance of 1.5–2.5 megaohms when filled with an internal solution containing 140 mM CsCl, 10 mM EGTA, 10 mM HEPES, 3 mM MgATP, 0.6 mM NaGTP, and 2 mM CaCl2 (pH adjusted to 7.2 with KOH, ~315 mosM). Recordings were filtered at 2 kHz. Data were analyzed using pCLAMP9 (Axon Instruments) and GraphPad Prism software. Student' t test or one-way analysis of variance combined with a Newman-Keuls post-test were used to compare the different values, which were considered significant at p < 0.05. Results are presented as the means ± S.E., and n is the number of cells used.

Imaging of PKC Translocation—To study PKC translocation as a function of temperature, we generated a human PKCbeta1 construct with the C terminus fused to GFP using the pEGFP-N1 plasmid (Clontech). PKCbeta1-GFP was inserted into pcDNA4 (Invitrogen) and transfected into CHO-CaV3.2 cells. Two days after, cells were plated onto 12-mm glass coverslips. The following day, phorbol 12-myristate 13-acetate (PMA) treatments were performed (100 nM in the culture medium for 10 min at 37 °C or at room temperature), and cells were then fixed for 20 min in 4% paraformaldehyde. Digital images were acquired on a Leica microscope and further analyzed using Adobe Photoshop.

Luminometric Analysis of HA-tagged CaV3.2 Channels—tsA-201 cells were cultured in 24-well plates and transfected with a GFP-CaV3.2-HA construct (27). Two days after transfection, PMA treatments were performed (100 nM in the culture medium for 30 or 60 min at 37 °C), and cells were then rinsed with phosphate-buffered saline (PBS) and fixed for 5 min in 4% paraformaldehyde. After three PBS washes, half of the wells were permeabilized with 0.1% Triton X-100 for 5 min and rinsed three times with PBS. Cells were then incubated for 30 min in blocking solution (PBS plus 1% fetal bovine serum). The GFP-CaV3.2-HA protein was detected using a rat anti-HA monoclonal antibody (1:1000 dilution; clone 3F10, Roche Diagnostics) after incubation for 1 h at room temperature. After four washes with PBS plus 1% fetal bovine serum for 10 min, cells were incubated for 30 min with horseradish peroxidase-conjugated goat anti-rat secondary antibody (1:1000 dilution; Jackson ImmunoResearch Laboratories, West Grove, PA). Cells were rinsed four times with PBS for 10 min before addition of SuperSignal enzyme-linked immunosorbent assay Femto maximum sensitivity substrate (Pierce). Luminescence was measured using a VICTOR2 luminometer (PerkinElmer Life Sciences), and the protein amount in each well was then measured using the BCA assay (Pierce) to normalize the measurements. All data were normalized to the level of signal obtained for incubation with the control medium (1:10,000 Me2SO). Four independent sets of transfection experiments were performed under each condition, and the results are presented as the means ± S.E.


Figure 2
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  		FIGURE 2.
Temperature dependence of PMA effects on CaV3.3 T-currents in transiently transfected tsA-201 cells. Shown is the effect of 100 nM PMA or the control solution (10-min incubation) on CaV3.3 currents at several temperatures, i.e. at 22 (room temperature), 27, 30, 32, and 37 °C. Electrophysiological experiments were performed at room temperature during the following 50 min in the presence of PMA or the control solution. Currents were elicited by a –45-mV depolarization of 450-ms duration applied immediately after the whole-cell configuration from a holding potential of –80 mV. Increases in CaV3.3 currents as a function of temperature were fitted with a sigmoidal Hill equation, where T-current increase (Y) = Ymin + (Ymax–Ymin)/(1 + (T0.5/T)Hill slope), where T is temperature. The fit gave a T0.5 value of 30.54 °C and a Hill slope of 18.87.

 


Figure 3
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  		FIGURE 3.
PMA induces an increase in CaV2.3 currents in transiently transfected tsA-201 cells at 37 °C but not at room temperature (~22 °C). A–C, effect of 100 nM PMA or the control solution (Ctrl) on CaV2.3 currents when incubated for 10 min at 37 °C (A and B) or at room temperature (R.T.; C) before electrophysiological experiments, which were performed at room temperature during the following 50 min in the presence of PMA or the control solution. Currents were elicited by a +10-mV depolarization of 500-ms duration applied immediately after the whole-cell configuration from a holding potential of –80 mV. ***, p < 0.001.

 
In Vitro PKA Assays—tsA-201 cells transfected with HA-tagged CaV3.2 or CaV2.1 were lysed in Nonidet P-40 buffer (1% Nonidet P-40, 150 mM NaCl, and 10 mM Tris-HCl (pH 7.6) supplemented with a protease inhibitor mixture (Roche Diagnostics)) for 30 min on ice. Lysates were centrifuged at 10,000 x g for 20 min, precleared for 1 h with fetal calf serum-Sepharose beads, and incubated overnight with specific antibodies and protein A-Sepharose beads (Amersham Biosciences). Beads were washed three times with Nonidet P-40 buffer before PKA assays. In vitro PKA assays were performed on Sepharose beads for 30 min at 30 °C with 10 units of the PKA catalytic subunit (Sigma) in 40 µl of medium containing 50 mM Tris-HCl, 10 mM MgCl2, 10 mM HEPES, 10 mM dithiothreitol, 1 mM Na3VO4, 1 mM MgATP, 0.15 mM CaCl2, 500 µM okadaic acid, and 10 µCi of [{gamma}-32P]ATP (Amersham Biosciences). After three washes with cold Nonidet P-40 buffer, standard SDS-PAGE was performed, and phosphorylated proteins were analyzed using a PhosphorImager.

Chemical Reagents—All compounds (which were from Sigma, except Gö 6976, which was from BIOMOL International LP) were dissolved in water (1 mM), except PMA, 4{alpha}-PMA, and Gö 6976, which were dissolved in Me2SO (1 mM); aliquoted; and kept at –80 °C in the dark. Control experiments were carried out using the solvent alone.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
To investigate the effects of protein kinase activation in mammalian cells expressing low voltage-activated T-currents, various protein kinase activators (or the control solution) were incubated with cells at 37 °C or at room temperature (~22 °C) for 10 min before electrophysiological recordings, which were performed at room temperature in either the presence or absence of the activator (Fig. 1, A and B, insets). Under these conditions, the PKC activator PMA (100 nM) induced a strong increase in the CaV3.1 currents (~100%, p < 0.001, n > 70) when preincubated at 37 °C (Fig. 1A). Similar results were obtained for CaV3.2 and CaV3.3 channels because PMA induced an ~105% increase (p < 0.001, n > 90) in CaV3.2 currents and had a maximal effect on CaV3.3 currents (~145%, p < 0.001, n > 100) (Fig. 1C). Interestingly, we found that PMA had no significant effect when incubated at room temperature (up to 1 h of incubation; p > 0.05 and n > 30 for each CaV3 current) (Fig. 1, B–D). We then evaluated the PMA effects on CaV3.3 currents during incubation at several intermediate temperatures (27, 30, and 32 °C). We found that the PMA effects gradually developed at 30 and 32 °C (~60 and 100% increases, respectively; p < 0.05; n > 40) (Fig. 2), whereas no effect was observed at 27 °C (p > 0.05, n > 34) (Fig. 2). The PMA effects as a function of temperature can be described by a sigmoidal Hill equation (Fig. 2), which indicates that the temperature producing PMA half-effects is ~30.5 °C and the Hill slope ~18.9, indicating a strong temperature dependence of PMA effects. We also investigated the temperature dependence of the PMA effects on high voltage-activated CaV2.3 currents, which are increased by PMA at room temperature in Xenopus oocytes (26). As observed for CaV3 currents, we found that 100 nM PMA induced a strong increase in CaV2.3 currents in human embryonic kidney 293 cells when incubated at 37 °C (~150%, p < 0.001, n > 30) (Fig. 3, A and B), whereas PMA has no effect when incubated at room temperature (p > 0.05, n > 35) (Fig. 3C).


Figure 4
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  		FIGURE 4.
Translocation of PKCbeta1-GFP in mammalian CHO cells as a function of temperature. Shown are the effects of 100 nM PMA or the control solution (Ctrl) on the cellular localization of PKCbeta1-GFP when incubated for 10 min at 37 °C (A and B) or at room temperature (RT; C and D). Scale bars = 20µM.

 
It has been described that PKC activation is associated with a redistribution of its subcellular localization (28). We asked whether this phenomenon is altered in mammalian cells at a non-physiological temperature (Fig. 4). To this end, we generated a GFP-tagged PKCbeta1 construct (PKCbeta1-GFP; see "Materials and Methods") to visualize PMA-induced PKC translocation in transfected mammalian cells (Fig. 4, B and C). As described for PKC of the classical family (28), we found that 100 nM PMA induced PKC translocation to the plasma membrane in mammalian CHO cells when incubated for 10 min at 37 °C (Fig. 4B). However, PMA had no effect on PKC translocation when incubated at room temperature (Fig. 4D).


Figure 5
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  		FIGURE 5.
Involvement of PKC in PMA-induced increases in the three CaV3 currents. A–C, summary of the data obtained for CaV3.1, CaV3.2, and CaV3.3, respectively, after a 10-min incubation at 37 °C of several compounds that activate (PMA, 100 nM), inhibit (chelerythrine (Chele.), 1 µM; and Gö 6976, 1 µM), or are ineffective (4{alpha}-PMA, 100 nM) on PKC. PKC inhibitors were applied at least 4 h (chelerythrine) or 18 h (Gö 6976) prior to the 37 °C incubation protocols. Currents were elicited by a –45-mV depolarization (200-ms duration for CaV3.1 and CaV3.2 currents and 450-ms duration for CaV3.3 currents) applied immediately after the whole-cell configuration from a holding potential of –80 mV. Ctrl, control. ***, p < 0.001.

 


Figure 6
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  		FIGURE 6.
Effects of PMA on T-current properties. A, steady-state CaV3.1 current activation properties. CaV3.1 currents were elicited by incremental test pulse depolarizations (from –80 to +50 mV, 10-mV increments) of 200-ms duration applied every 5 s from a holding potential of –100 mV. B, steady-state CaV3.1 current inactivation properties. Currents were elicited by a –30-mV test pulse of 200-ms duration applied from holding potentials ranging from –110 to –50 mV (10-s duration, 5-mV increments). C, no significant effect of 100 nM PMA after a 10-min incubation at 37 °C on the current-voltage curves of CaV3.1 currents. Current-voltage curves were obtained from the experiments illustrated in A. D, no significant effect of 100 nM PMA after a 10-min incubation at 37 °C on the steady-state inactivation (h{infty}) curve of CaV3.1 currents as well as on the steady-state activation (m{infty}) curve. The steady-state inactivation (h{infty}) curve was obtained from experiments illustrated in B, whereas the steady-state activation (m{infty}) curve was deduced from the fit of the current-voltage curves presented in C. For clarity, the corresponding symbol of the m{infty} curve was omitted. E and F, no significant effect of 100 nM PMA on the steady-state activation (m{infty}) and inactivation (h{infty}) properties of CaV3.2 and CaV3.3 currents, respectively. The steady-state activation (m{infty}) and inactivation (h{infty}) curves of CaV3.2 and CaV3.3 currents were deduced from same experiments shown in A and B, except that the test pulses to elicit CaV3.3 currents were of 450-ms duration. Ctrl, control.

 
We next investigated whether PMA effects on CaV3 currents are specific to PKC activation (Fig. 5). To assess the involvement of PKC, we used 4{alpha}-PMA (100 nM), a PMA analog inactive on PKC, and chelerythrine or Gö 6976 (both at 1 µM), two selective inhibitors of PKC (Fig. 5). We found that 4{alpha}-PMA had no significant effect on the three CaV3 currents (p > 0.05 and n > 25 for each CaV3 current), whereas both chelerythrine and Gö 6976 suppressed PMA effects (p > 0.05 and n > 25 for each CaV3 current). In addition, we found no basal effect of chelerythrine on the three CaV3 currents (p > 0.05 and n > 60 for each CaV3 current) (Fig. 5, A–C).

To estimate whether PKC activation-induced increase in T-currents is associated with changes in macroscopic biophysical properties, we next performed steady-state activation and inactivation protocols (Fig. 6, A and B, insets). As shown for CaV3.1 (Fig. 6, A–D), PMA induced no significant change in steady-state activation and inactivation properties. For CaV3.1, current-voltage curves (Fig. 6C) indicated steady-state activation V0.5 values of –48.6 ± 1.4 mV (n = 10) under control conditions and –47.6 ± 1.3 mV (n = 13, p > 0.05) after PMA treatment, whereas slope values were 4.1 ± 0.4 mV (n = 10) under control conditions and 4.3 ± 0.4 mV (n = 13, p > 0.05) after PMA treatment. Similarly, steady-state inactivation properties were not significantly modified by PKC activation (Fig. 6D) because V0.5 values were –70.7 ± 0.7 mV (n = 10) under control conditions and –71.8 ± 0.6 mV (n = 11, p > 0.05) after PMA treatment, whereas slope values were 4.7 ± 0.2 mV (n = 10) under control conditions and 4.1 ± 0.2 mV (n = 11, p > 0.05) after PMA treatment. In the same way, neither the steady-state activation nor inactivation properties of the CaV3.2 (n > 10 under each condition, p > 0.05) (Fig. 6E) and CaV3.3 (n > 10 under each condition, p > 0.05) (Fig. 6F) currents were significantly affected by PMA treatment.

We also investigated whether the surface expression of CaV3 channels in mammalian tsA-201 cells is modulated by PKC activation (supplemental Fig. 1). To this end, we used a CaV3.2 channel construct containing an extracellular HA tag, which allowed its surface (non-permeabilized condition) and total expression (permeabilized condition) (supplemental Fig. 1A) to be measured by enzyme-linked immunosorbent assay/luminometry. We found that PMA treatment did not induce significant changes in both membrane expression (supplemental Fig. 1B) and total expression (supplemental Fig. 1C) of CaV3.2-HA channels after 30 min or 1 h of treatment at 37 °C. Also, protein kinase activation did not influence the ratio of membrane expression to total expression, which was ~20% under all conditions (supplemental Fig. 1D), further indicating that surface expression of CaV3.2-HA channels is not altered.


Figure 7
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  		FIGURE 7.
Bt2cAMP induces an increase in CaV3 T-currents in transiently transfected tsA-201 cells at 37 °C but not at room temperature. A and B, effect of 1 mM Bt2cAMP (dB-cAMP) on CaV3.1 currents when incubated for 10 min at 37 °C and at room temperature (RT), respectively, before electrophysiological experiments. C and D, summary of the data obtained for CaV3.1, CaV3.2, and CaV3.3 currents upon incubation of Bt2cAMP at 37 °C and at room temperature, respectively. Currents were elicited by a –45-mV depolarization (200-ms duration for CaV3.1 and CaV3.2 currents and 450-ms duration for CaV3.3 currents) applied immediately after the whole-cell configuration from a holding potential of –80 mV. Ctrl, control. ***, p < 0.001.

 
We next explored whether, as observed for PKC, cyclic nucleotide-dependent protein kinases modulate T-currents in mammalian cells at physiological temperatures. To investigate the effects of PKA, we used dibutyryl cAMP (Bt2cAMP), a membrane-permeant analog of cAMP (Fig. 7). We found that Bt2cAMP induced an increase in the three CaV3 T-currents when incubated at 37 °C (Fig. 7, A–C) but had no significant effect when incubated at room temperature (p > 0.05 and n > 30 for each CaV3 current) (Fig. 7, B–D). As observed above with PMA, Bt2cAMP produced a stronger effect on CaV3.3 currents (~140% increase, p < 0.001, n > 60) compared with CaV3.2 currents (~70% increase, p < 0.001, n > 80) and CaV3.1 currents (~55% increase, p < 0.001, n > 80) (Fig. 7C). Furthermore, we observed a similar temperature dependence using monobutyryl cAMP (the bioactive product of Bt2cAMP acting on PKA). Indeed, monobutyryl cAMP treatment increased CaV3.3 currents at 37 °C (~110% increase, p < 0.01, n > 35) but not at room temperature (p > 0.05, n > 40). These latter results demonstrate that the temperature dependence described here does not involve endogenous esterases or amidases, which convert Bt2cAMP into monobutyryl cAMP (for review, see Ref. 29). We next probed the involvement of PKA activation in the Bt2cAMP effects using the specific PKA inhibitor KT5720. KT5720 suppressed Bt2cAMP-induced increases in T-currents (p > 0.05 and n > 20 for each CaV3 current) (Fig. 8, A–C) but had no effect on basal T-currents (p > 0.05 and n > 20 for each CaV3 current). It should be noted that in previous experiments, we used the PKA inhibitor H-89, but we observed that although this compound abolished Bt2cAMP effects, it had strong direct inhibitory effects at micromolar concentrations on the three CaV3 currents and especially on CaV3.2 (data not shown). We further investigated whether PKA effects would involve a direct phosphorylation of CaV3 channels. To this end, we took advantage of a commercially available PKA catalytic subunit that is constitutively active and that allows in vitro kinase assays of immunoprecipitated HA-tagged CaV3.2 channels (Fig. 8D). We found that PKA induced phosphorylation of CaV3.2 channels, but not of CaV2.1 channels, which are insensitive to PKA (6, 8) and were used here as a negative control (Fig. 8D).


Figure 8
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  		FIGURE 8.
Involvement of PKA in Bt2cAMP-induced increases in the three CaV3 currents and direct phosphorylation of CaV3.2 channels by PKA. A–C, summary of the data obtained for CaV3.1, CaV3.2, and CaV3.3, respectively, after 10 min of incubation at 37 °C of Bt2cAMP (dB-cAMP; 1 mM) and/or KT5720 (a PKA inhibitor; 0.5 µM). KT5720 was applied at least 4 h at 1 µM prior to the 37 °C incubation protocols. Currents were elicited by a –45-mV depolarization (200-ms duration for CaV3.1 and CaV3.2 currents and 450-ms duration for CaV3.3 currents) applied immediately after the whole-cell configuration from a holding potential of –80 mV. Ctrl, control. D, effects of the constitutively active PKA catalytic subunit on immunoprecipitated HA-tagged CaV2.1 and CaV3.2 channels in in vitro phosphorylation assays. In vitro phosphorylation assays were performed at 30 °C using [{gamma}-32P]ATP, and 32P-labeled proteins were quantified using a PhosphorImager after standard SDS-PAGE. The presence of HA-tagged CaV2.1 and CaV3.2 channels was further confirmed by Western blotting (not shown). ***, p < 0.001.

 
In contrast with the results obtained for PKA activation, the activation at physiological temperature of PKG with dibutyryl cGMP (Bt2cGMP) had no significant effect on the three CaV3 currents (p > 0.05 and n > 40 for each CaV3 current) (supplemental Fig. 2, A and B). Furthermore, we treated cells with the specific PKG inhibitor KT5823 (supplemental Fig. 2B) and found that this compound had no significant effect on the three CaV3 current densities (p > 0.05 and n > 20 for each CaV3 current) (supplemental Fig. 2B).

Finally, we further investigated the effect of PKC, PKA, and PKG activation in a CHO cell line stably expressing CaV3.2 currents (Fig. 9). In these cells, we found that PMA and Bt2cAMP (but not Bt2cGMP) induced an increase in CaV3.2 currents when incubated at physiological temperature (Fig. 9A). As observed above in transiently transfected tsA-201 cells, PMA induced stronger effects on CaV3.2 currents (~125% increase) compared with Bt2cAMP (~50% increase, p < 0.001, and n > 30 under each condition) (Fig. 9B). However, PMA and Bt2cAMP had no significant effect when incubated at room temperature (up to 1 h of incubation; p > 0.05 and n > 24 under each condition) (Fig. 9C).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The main findings of our study are that recombinant T-channels are modulated by PKA and PKC (but not by PKG) in different mammalian cells and that these effects are temperature-dependent. Considering the essential role of T-channels in cardiac and neuronal pacemaking (10, 30–32) as well as in slow wave sleep (33, 34), absence epilepsy (35, 36), and pain perception (37–39), the investigation of their modulation at the molecular level is of crucial importance. Three T-channel subunits have been identified: CaV3.1, CaV3.2, and CaV3.3 (7, 10). Among these three subunits, only CaV3.2 is modulated in mammalian cells by beta{gamma}-subunits of G proteins and by Ca2+/calmodulin-dependent protein kinase II (11–15). Consequently, the molecular pathways implicated in CaV3.1 and CaV3.3 regulation remain largely unknown (9). In this study, using the PKA activator Bt2cAMP and the PKC activator PMA, we have demonstrated that PKA and PKC activation increases CaV3.1, CaV3.2, and CaV3.3 currents in mammalian tsA-201 cells. Both Bt2cAMP and PMA produced stronger effects on CaV3.3 currents (~150% increase) compared with CaV3.1 and CaV3.2 currents. In addition, CaV3.1 and CaV3.2 currents were more sensitive to PMA (~100% increase) compared with Bt2cAMP (~50% increase). The involvement of PKA in Bt2cAMP effects was further demonstrated with KT5720, a specific PKA inhibitor (40), which suppressed the Bt2cAMP-induced increase in T-currents. In the same way, PKC is implicated in PMA-induced T-current increase, as assessed with 4{alpha}-PMA, an analog of PMA inactive on PKC. Furthermore, the PMA effect is abolished by the PKC inhibitors chelerythrine and Gö 6976 (41, 42). This pharmacological profile of PKC inhibition suggests involvement of the classical PKC family (42), the members of which are recruited by Gq-coupled receptors. In contrast with PKA and PKC activation, we found that PKG activation with Bt2cGMP had no effect on the three CaV3 currents. This lack of effect of PKG activation could be explained by a high basal activity of this kinase in tsA-201 cells. However, basal PKG inhibition with the specific PKG inhibitor KT5823 (40) had no significant effect on the three CaV3 current densities, further indicating that PKG does not modulate T-currents. Because the results described above could be restricted to the cell type studied and/or to transient expression, we investigated the effect of PKC, PKA, and PKG activation in a CHO cell line stably expressing CaV3.2 currents. In these cells, we confirmed that PMA and Bt2cAMP (but not Bt2cGMP) induced an increase in CaV3.2 currents. As observed above in transiently transfected tsA-201 cells, PMA induced stronger effects on CaV3.2 currents compared with Bt2cAMP. Therefore, our results demonstrate that protein kinase modulation of T-currents does not depend on the mammalian cell type studied.


Figure 9
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  		FIGURE 9.
Effects of PMA, Bt2cAMP, and dibutyryl cGMP (Bt2cGMP) on CaV3.2 currents stably expressed in CHO cells. A, effects of 100 nM PMA, 1 mM Bt2cAMP and 1 mM Bt2cGMP after 10 min incubation at 37 °C on CaV3.2 currents stably expressed in CHO cells. B, summary of the data obtained for PMA, Bt2cAMP and Bt2cGMP after 10 min incubation at 37 °C. C, summary of the data obtained for PMA, Bt2cAMP, and Bt2cGMP after 10 min incubation at room temperature (~22 °C). Currents were elicited by a –45 mV depolarization of 200 ms duration applied immediately after the whole-cell configuration from a holding potential of –80 mV. **, p < 0.01; ***, p < 0.001.

 
Modulation of CaV3 currents by protein kinases has been observed in Xenopus oocytes but not in mammalian cells expressing CaV3 currents (11, 16–19). This absence of kinase modulation in mammalian cells could be explained by the presence or absence of specific protein kinase isoforms, protein kinase-interacting proteins, and/or T-channel auxiliary subunits. In addition, T-channel pore {alpha}1-subunits are also subject to alternative splicing, which influences their electrophysiological properties (43–45) and possibly their regulation. However, we have demonstrated in this study that the currents generated by the minimal isotype of each pore {alpha}1-subunit (including no additional exon) (46) are modulated by protein kinases when expressed alone (without any auxiliary subunit) in mammalian cells. We have also reported that T-currents are not under the control of basal activation of protein kinase in mammalian cells, as assessed with specific kinase inhibitors. Furthermore, we have highlighted here that the physiological temperature is crucial for protein kinase effects. Indeed, we have shown that both PKA and PKC effects occur near physiological temperature (37 °C) but not at room temperature (22–27 °C) in both tsA-201 and CHO cell lines. This temperature dependence is well described by a sigmoidal Hill function, which indicates that the temperature producing PMA half-effects on CaV3.3 currents is ~30.5 °C and the Hill slope ~18.9, indicating a strong temperature dependence of PMA effects. A similar temperature dependence was observed for the three CaV3 currents as well as for CaV2.3 currents (a high voltage-activated calcium channel), which was strongly increased by PKC at 37 °C in mammalian cells. In addition, we have provided evidence that PMA-induced PKC translocation to the plasma membrane (where T-channels display their activity) is altered in mammalian cells at non-physiological temperatures. It should be noted that contrary to mammalian cells, PKC translocation is fully preserved in Xenopus oocytes at room temperature (47). In this context, it appears that our results on CaV3 and CaV2.3 currents are reminiscent of those obtained for modulation of L-type CaV1.2 currents by PKA. Indeed, although the CaV1.2 subunit is directly phosphorylated by PKA in in vitro kinase assays at Ser1928 (8, 48), the phosphorylation and modulation of L-type CaV1.2 currents by PKA activators does not occur in human embryonic kidney 293 cells in the absence of AKAP (A protein kinase-anchoring protein) (8, 49, 50). In fact, AKAP binds and tethers PKA to particular sites at the plasma membrane where CaV1.2 channels reside, allowing their phosphorylation at room temperature in mammalian cells and the increase in the current (8, 49–51). Overall, although we cannot completely exclude that kinase activity is involved in the temperature dependence described here, our results strongly suggest that kinase localization (translocation) is impaired at non-physiological temperature. It is interesting to note that PMA effects on T-currents occur near 30 °C, a temperature that is permissive for export from the Golgi network (52, 53), which is implicated in the translocation of PKC (54). Indeed, incubation at 20 °C, which blocks exocytotic vesicle traffic from the Golgi network, abolishes the PMA-induced translocation of PKC to the plasma membrane in mammalian NIH 3T3 cells (54). Therefore, it is attractive to suggest that this phenomenon could be involved in the temperature dependence described in our study (as suggested by our PKC-GFP imaging).

We have also provided data demonstrating that protein kinase effects do not involve changes in T-current macroscopic biophysical properties that could explain the modulation. In addition, the surface expression of CaV3 channels is not altered upon kinase activation, as assessed by enzyme-linked immunosorbent assay/luminometry experiments. Furthermore, the in vitro kinase assays suggest that PKA could act directly on T-channels. Overall, our results are in good agreement with those obtained for the PKA modulation of L-type currents (22). Indeed, the CaV1.2 subunit is directly phosphorylated by PKA, whereas L-type current macroscopic properties are weakly affected by PKA activation. Furthermore, single channel recordings have highlighted an increase in the number of functional channels (resulting from an increase in the proportion of non-blank sweeps), which underlies PKA effects on L-type currents (see Ref. 22 for an insightful review). Therefore, it is tempting to suggest that this mechanism might be conserved for kinase action on VGCCs and that PKA and PKC activation transforms nonfunctional forms of T-channels to functional forms.

Xenopus oocytes and mammalian heterologous expression systems are currently used to study recombinant ion channels. Although these expression systems have provided a lot of important information, especially about electrophysiological ion channel properties, the results obtained in these two systems concerning ion channel modulation are often divergent (9). Our present findings indicate that at least for calcium channels, the physiological temperature is determinant for studying ion channel modulation by protein kinases in mammalian cells. In conclusion, our study clearly establishes the role of PKA and PKC in the modulation of CaV3 T-currents.


   FOOTNOTES
 
* This work was supported in part by CNRS; Agence Nationale de la Recherche Research Grant ANR-05-NEUR-031; and research grants from Agence de Recherche contre le Cancer-Institut contre le Cancer 2006, the Institut UPSA de la Douleur, and the Association Française contre les Myopathies. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement"in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. 1 and 2. Back

2 Supported by the Ligue Francaise contre l'Épilepsie. Back

3 Supported by the Ligue Régionale contre le Cancer (Languedoc-Roussillon). Back

4 Supported by a grant from Pfizer France. Back

1 To whom correspondence should be addressed: Dépt. de Physiologie, Inst. de Génomique Fonctionnelle, 141, Rue de la Cardonille, Montpellier 34094, France. Tel.: 33-499-619-939; Fax: 33-499-619-901; E-mail: jean.chemin{at}igf.cnrs.fr.

5 The abbreviations used are: VGCCs, voltage-gated Ca2+ channels; PKC, protein kinase C; PKA, protein kinase A; PKG, protein kinase G; CHO, Chinese hamster ovary; GFP, green fluorescent protein; HA, hemagglutinin; PMA, phorbol 12-myristate 13-acetate; PBS, phosphate-buffered saline; Bt2cAMP, dibutyryl cAMP; Bt2cGMP, dibutyryl cGMP. Back


   ACKNOWLEDGMENTS
 
We thank Drs. F. A. Rassendren, G. Vassort, D. Joubert and C. Quittau-Prevostel as well as all members of the Département de Physiologie for constructive comments on the manuscript. We are grateful to Dr. E. Bourinet and E. Kupfer for the development of the CHO-CaV3.2 cell line and Dr. T. Snutch for the gift of CaV2.3, {alpha}2{delta}1, and beta4 constructs. We are grateful to A. Lacan for help with PKC experiments.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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