A T-type channel-calmodulin complex triggers αCaMKII activation
© The Author(s). 2017
Received: 24 March 2017
Accepted: 28 July 2017
Published: 11 August 2017
Calmodulin (CaM) is an important signaling molecule that regulates a vast array of cellular functions by activating second messengers involved in cell function and plasticity. Low voltage-activated calcium channels of the Cav3 family have the important role of mediating low threshold calcium influx, but were not believed to interact with CaM. We find a constitutive association between CaM and the Cav3.1 channel at rest that is lost through an activity-dependent and Cav3.1 calcium-dependent CaM dissociation. Moreover, Cav3 calcium influx is sufficient to activate αCaMKII in the cytoplasm in a manner that depends on an intact Cav3.1 C-terminus needed to support the CaM interaction. Our findings thus establish that T-type channel calcium influx invokes a novel dynamic interaction between CaM and Cav3.1 channels to trigger a signaling cascade that leads to αCaMKII activation.
Calcium channels mediate a wide range of cellular functions that involve activation of calcium-dependent enzymes that can shape neuronal output . Members of the Cav3 calcium channel family are designed to regulate calcium entry in the subthreshold voltage range to control action potential generation [2–4] and transmitter release [5–8]. Dysregulated Cav3 channels contribute to pathologies such as epilepsy, chronic pain, and ataxia [1, 9–11]. It has been well established that high voltage-activated (HVA) calcium channels interact with the calcium sensing protein calmodulin (CaM) to regulate channel function [12–14]. Moreover, calcium entry through HVA calcium channels can trigger a second messenger cascade by which CaM activates Ca2+/CaM-dependent protein kinase II (CaMKII) that can modify cell activity and long-term plasticity [15–19]. In contrast, the Cav3 channel family is reported to lack CaM binding consensus motifs that would permit calcium-dependent interactions with CaM [20, 21] and there are no reports of Cav3 calcium influx triggering a signaling cascade for CaMKII activation. The available data then suggest that activation of CaM-CaMKII signaling by voltage-dependent calcium channels is restricted to HVA calcium channels that open in response to large membrane depolarizations.
We now report that under resting conditions, CaM constitutively associates with the C-terminus of Cav3.1, as revealed in both a heterologous expression system and rodent brain. In response to Cav3.1-mediated calcium influx the Cav3.1-CaM association is lost, accompanied by phosphorylation of αCaMKII in both a heterologous expression system and neurons in hippocampus and cerebellum. These results are important in providing the first evidence of a role for Cav3.1 channels in triggering αCaMKII as a second messenger involved in mediating a host of cellular functions.
Animal care and tissue dissection
Sprague-Dawley rats and C57/BL/6 mice were obtained from Charles River and maintained according to guidelines of the Canadian Council for Animal Care, and euthanized according to Standard Operating Procedures defined by the University of Calgary Animal Resource Center. All chemicals were obtained from Sigma (Oakville, ON, Canada) unless otherwise indicated. Animals at P19–40 were anaesthetized by isofluorane inhalation until unresponsive to ear pinch and the brain dissected out in the presence of ice cold sucrose-cutting solution composed of (in mM): 215 sucrose, 25 NaHCO3, 20 D-glucose, 2.5 KCl, 1.25 NaH2PO4 and 3 MgCl2 preoxygenated by carbogen (95% O2, 5% CO2) gas . For coimmunoprecipitations brain tissue was transferred to a lysis buffer medium comprised of (in mM): 150 NaCl, 50 Tris pH 7.5, 0.5% sodium-decholic acid, 1% NP-40, 2.5 EGTA, 0–3 CaCl2, and 1 tablet complete protease inhibitor cocktail (EDTA free) in 10 ml, and stored at 4 °C. A separate set of experiments prepared brain tissue to create homogenates in different buffered levels of calcium concentration according to Lai et al. . For tissue slice histochemistry sagittal cerebellar tissue slices of 150 μm thickness were prepared from P30-P40 male rats by Vibratome in ice-cold sucrose-cutting solution as above. For tissue destined for recordings, slices were incubated at 37 °C in artificial cerebrospinal fluid (aCSF) composed of (mM): 125 NaCl, 1.0 KCl, 1.5 CaCl2, 1.5 MgCl2, 25 NaHCO3, and 25 D-glucose bubbled with carbogen gas, with a subset exposed to aCSF containing 50 mM [K]o for 10 min prior to immediate removal and fixation for immunolabel processing. Calcium influx was restricted to Cav3 calcium channels throughout by cutting tissue slices and maintaining tissue in aCSF containing 30 μM Cd2+, 1 μM TTX, 10 μM DNQX and 25 μM or 100 μM DL-AP5.
Low-density cultures of hippocampal neurons were prepared from P0-P1 pups of C57/BL/6 mice initially anesthetized by placing on ice for 5–10 min. Dissected hippocampi were dissociated by papain treatment and trituration and plated on poly-l-lysine and laminin-coated glass coverslips in 24 well culture dishes at a density of 3500–5300 cells/cm2. After plating, cells were allowed to grow on coverslips for 10 days in medium composed of (in mM): free neurobasal medium (BME), 1% B-27 supplement, 5% FBS, 0.6% glucose, 1 Na-Pyruvate, 2 L-glutamine, 10 HEPES, 1% Pen-strep. All cell culture chemicals were obtained from Life Technologies (Burlington, ON, Canada).
Wild-type human Cav3.1b cDNA was donated by T. Snutch (Vancouver, BC, Canada) (GenBank accession number AF134986.1), HA-Cav3.1 and Kir2.1 cDNA from E. Bourinet (Institut de Génomique Fonctionnelle, Montpellier, FR), and CaM cDNAs by J. Adelman (Vollum Institute, OR, USA). GFP-Cav3.1 and GFP-αCaMKII were subcloned into a GFP mammalian vector (pCMV6-AN-GFP) (OriGene Technologies, Rockville, MD, USA) and mKate-CaM prepared from a pmKate2-N mammalian expression vector (Axxora, Farmingdale, NY, USA). CaMKIIN was amplified from a mouse hippocampal cDNA library (RIKEN) and cloned into the pcDNA3.1 vector (Addgene, Cambridge, MA, USA) . To create a deletion of the C-terminus, PCR with specific primers was performed on Cav3.1 outside of the C-terminus region, followed by sequencing and subcloning into the pCMV6-AN-GFP vector (OriGene Technologies, Rockville, MD, USA) or the pcDNA3.1 vector (Addgene, Cambridge, MA, USA) and sequenced for final confirmation. The GFP and mKate tags were attached to the N-terminal regions of target proteins. To create the Cav3.1 pore mutant, a single point mutation PCR was performed on E354K in the pore region with the primer sequences: 5′-ACAGGTCGTTGAGCCGC-3′, followed by sequencing and subcloning into the pcDNA3.1 vector (Addgene, Cambridge, MA, USA) and sequenced for final confirmation.
cDNA constructs were transiently transfected into tsA-201 cells using the calcium phosphate method . Coimmunoprecipitation experiments used 5 μg cDNA for each of Cav3.1, HA-Cav3.1, CaM, CaM1234, and GFP-Cav3.1 lacking the C-terminus (GFP-Cav3.1ΔCT). FRET studies used 1 μg cDNA of GFP-Cav3.1 and mKate-CaM, and live cell imaging experiments and immunocytochemistry used 1 μg Cav3.1 or Cav3.1ΔCT or Cav3.1 pore mutant, 1 μg CaM, and 0.7 μg GFP-αCaMKII. tsA-201 were also transfected with 0.3 μg Kir2.1 cDNA where indicated. Transfected cells were incubated at 37 °C (5% CO2) for 24 h and then transferred to a 30 °C incubator (5% CO2) for 48–72 h prior to imaging studies or incubated at 30 °C for 48 h before being lysed using a buffered calcium concentration buffer for coimmunoprecipitations.
tsA-201 cells transiently transfected with Cav3.1 or Cav3.1ΔCT and either CaM or CaM1234 were lysed in a buffer containing (in mM): 150 NaCl, 50 Tris, 2.5 EGTA, 0.5% sodium-decholic acid, 1% NP-40, 0–3 CaCl2, pH 7.5. The nominal level of free calcium in solutions was calculated using Maxchelator software (http://maxchelator.stanford.edu/CaEGTA-NIST.htm) to conduct coimmunoprecipitations in the presence of different levels of free calcium concentration ranging from 0 to 1 mM (see ). Brain tissue was prepared in cell lysis buffer using a hand held glass homogenizer. Lysates from tsA-201 cells were centrifuged at 13,000 rpm for 1 min at 4 °C and supernatants were transferred to new tubes. Solubilized proteins were incubated with 1 μg of rabbit anti-Cav3.1 antibody (1:1000) corresponding to amino acid residues of the I-II linker of rat CaV3.1 , or with anti-CaM antibody (1:1000; Millipore, Etobicoke, ON, Canada) and 30 μl of Protein G beads (Life Technologies, Burlington, ON, Canada) while rotating overnight at 4 °C. Coimmunoprecipitates were washed three times with phosphate buffered saline (PBS) in (mM): 137 NaCl, 2.7 KCl, 1 Na2HPO4, 1.8 KH2PO4, pH 7.4 reconstituted in an equal volume of 2X loading buffer (in mM): 100 Tris, 100 2-mercaptoethanol, 4% SDS, 0.02% bromophenol blue, 20% glycerol, pH 6.8 and incubated at 100 °C for 5 min. Eluted samples were loaded on 6–10% Tris-glycine gel and resolved using SDS-PAGE. Samples were transferred to 0.2 μm PVDF membrane (Millipore, Etobicoke, ON, Canada) and Western blot analysis performed using a mouse monoclonal anti-CaM antibody (1:1000; Millipore, Etobicoke, ON, Canada) or a mouse monoclonal anti-Hemagglutinin (HA) antibody (0.4 μg; Roche, Mississauga, ON, Canada). Secondary antibodies were conjugated to horseradish peroxidase (HRP; 1:5000; Molecular Probes, Eugene, OR, USA) and reacted with ECL solution (Life Technologies, Burlington, ON, Canada).
GST-pull down assay
To prepare the GST-C-terminus of Cav3.1, we transformed Cav3.1 C-terminus GST fusion protein plasmids into BL21 and induced the expression of recombinant proteins by 0.5 mM isopropyl-β-D-1-thiogalactopyranoside. GST fusion proteins were purified from bacteria using glutathione-Sepharose 4B beads (Life Technologies, Burlington, ON, Canada) according to the protocol recommended by the manufacturer. The GST tag was then cleaved using Precision Protease (Life Technologies, Burlington, ON, Canada). Complete cleavage of the fusion protein was verified using Western blotting. The recombinant Cav3.1 C-terminus peptides were then applied to CaM sepharose beads and incubated overnight. The CaM sepharose beads were washed three times with PBS, reconstituted in equal volume of loading buffer (100 mM Tris, 4% SDS, 0.02% bromophenol blue, 20% glycerol, 100 mM 2-mercaptoethanol, pH 6.8) and incubated at 100 °C for 5 min. Eluted samples were loaded on a 6–10% Tris-glycine gel and resolved using SDS-PAGE. Samples were transferred to 0.2 μm PDVF membrane (Millipore, Etobicoke, ON, Canada) and western blot analysis was performed using a goat anti-Cav3.1 antibody targeted to the C-terminus region of the channel (1:1000; Santa Cruz, Dallas, TX, USA). The secondary antibody used was a goat antibody conjugated to HRP (1:5000; Molecular Probes, Eugene, OR, USA) and reacted with ECL solution (Life Technologies, Burlington, ON, Canada).
Cerebellar tissue slices lysates of 150 μm thickness were incubated for 10 min in any of three conditions: low [K]o (1 mM), high [K]o (50 mM), or high [K]o with mibefradil (1 μM) and Ni2+ (300 μM). Homogenates were then made in the same solutions using a hand held glass homogenizer. Eluted lysates were loaded on 6–10% Tris-glycine gel and resolved using SDS-PAGE. Samples were transferred to 0.2 μm PVDF membrane (Millipore, Etobicoke, ON, Canada) and Western blot analysis performed using a monoclonal mouse anti-αCAMKII (1:1000; Santa Cruz, Dallas, TX, USA) or a polyclonal rabbit anti-p-αCAMKII (Thr286) (1:1000; Santa Cruz, Dallas, TX, USA). The secondary antibodies used were the appropriate mouse or rabbit antibodies conjugated to HRP (1:5000; Molecular Probes, Eugene, OR, USA) and reacted with ECL solution (Life Technologies, Burlington, ON, Canada).
Live cell fluorescence spectral confocal imaging
Cultured tsA-201 cells were transiently transfected with GFP-Cav3.1 and mKate-CaM constructs for FRET imaging. Cells were seeded onto poly-L-lysine coated 35 mm plates. Cells were then incubated for 24 h at 37 °C and 5% CO2, washed and replaced with colorless imaging medium (in mM): 148 NaCl, 3 KCl, 10 HEPES, 0 or 3 CaCl2, 8 glucose, 1 MgCl2, pH 7.3 at 25 °C. Cultured cells were examined with a Nikon Eclipse C1si spectral confocal laser-scanning microscope with a 40×/1.3NA oil immersion objective. For FRET imaging GFP-Cav3.1 was excited at 457 nm and mKate-CaM at 561 nm. Emission spectra of GFP and mKate were recorded between 400 nm to 750 nm. The FRET signal was measured every 10 s for 450 s.
ΔR(t) is the % change of mKate:GFP intensity ratio as a function of time (positive values mean an increase in mKate emission relative to GFP, implying greater FRET and therefore closer physical proximity of the two fluorescent proteins).
FmKate(t) and FGFP(t) are the mean fluorescence intensities of the two fluorophores as a function of time
FmKate(0) and FGFP(0) are the mean fluorescence intensities of the two fluorophores at time 0
For tests using GFP-αCaMKII expression tsA-201 cells were placed in colorless imaging medium comprised of (in mM): 148 NaCl, 1 or 50 KCl, 10 HEPES, 3 CaCl2, 8 Glucose, 1 MgCl2, pH 7.4 at 25 °C. For tests using GFP-αCaMKII transfection of hippocampal cultures, cells were transferred to aCSF containing 30 μM Cd2+, 1 μM TTX, 10 μM DNQX and 25 μM or 100 μM DL-AP5. The GFP-αCaMKII was excited at 457 nm and fluorescence signal in the cytoplasmic and nuclear compartment of cells was measured (ImageTrak). Images were recorded every 5 or 10 s for 250 or 400 s in control (1 mM [K]o) and test media (50 mM [K]o), and in the case of Fig. 4a, b, returned to aCSF containing low [K]o. The aggregation was calculated as the change in pixel variance of GFP fluorescence (ΔSD) presented as a percentage change from control variance (SD0G) to quantify the degree of GFP-αCaMKII aggregation (ΔSD/SD0G (%)) (Figs. 4 and 6; Additional file 5: Figure S5). Alternatively, GFP-αCaMKII aggregation was calculated as the mean fluorescence intensity change (ΔF) presented as a change from control mean (F0) (Figs. 5 and 7; Additional file 8: Figure S8).
Tests for phosphorylation of GFP-αCaMKII and phosphorylation of αCaMKII were conducted on cultured tsA-201 cells transiently transfected with cDNA, mouse hippocampal cell cultures, and rat cerebellar tissue slices maintained as above and exposed to control or test conditions (1 mM or 50 mM [K]o). Cultured cells and tissues were then quickly washed in PBS and fixed by exposure to 4% paraformaldehyde (pH 7.4) at room temperature for 1 h and overnight at 4 °C. The cells or tissue were transferred to a working solution of 3% normal donkey or horse serum (Jackson Immuno-Research, West Grove, PA, USA), 0.2% TWEEN and 2% dimethylsulphoxide in phosphate buffer (PB). Primary antibodies were reacted for 48 h at 4 °C and washed in working solution 3 X 15 min and secondary antibodies for 4 h at room temperature. Primary antibodies corresponded to a monoclonal mouse anti-p-αCAMKII (Thr286) (1:500; Life Technologies, Burlington, ON, Canada). Secondary antibodies (1:1000) were the appropriate AlexaFluor-488 or −594 conjugated rabbit IgGs (Molecular Probes, Eugene, OR, USA). After washing in PB, sections were mounted on gel-coated slides, coverslipped with anti-fade medium and stored at −20 °C. Controls consisted of omitting the primary antibodies.
A single compartment model of Cav3 T-type calcium channels was modified from data originally presented in Engbers et al. . The model had three conductances including Cav3, HCN and a leak current. Cav3 current was based on recordings from rat cerebellar Purkinje cells, and assigned the parameters of Va −30 mV, ka −4.6, λact 3 msec; Vh: −68 mV, kh: 6.8, λinact 12 msec; ECa = 40 mV; single channel conductance 9 pS. Parameters for IH were Vh −80 mV, k 3, λact 200 ms; EH = −20 mV. Eleak = −77 mV. Calcium diffusion was modeled using 10 hemispherical compartments with radii of 20–200 nm (20 nm increments) using a calcium diffusion coefficient of 220 μm2/msec. Simulations were run with custom made scripts in Matlab r2016a .
Statistical analysis was performed using Igor software (Wavemetrics, Lake Oswego, OR, USA) or IBM SPSS Statistics 20 Software (Armonk, NY, USA). The homogeneity of variances was assessed with Levene’s and Bartlett’s tests. Statistical significance was determined using a paired or unpaired Student’s t-test, one-way analysis of variance (ANOVA), or with the non-parametric multiple comparison Dunn-Holland-Wolfe and Mann-Whitney Wilcoxon tests as appropriate. Statistical significance was set at p < 0.05 with * p < 0.05, ** p < 0.01, ns, not significant. The average values are presented either as mean ± SEM or SD as indicated.
Cav3.1 coimmunoprecipitates with CaM in a calcium-dependent manner
It is known that at resting levels of calcium Cav1.2 channels are preassociated with ApoCaM and that an increase in the internal concentration of calcium leads to an increase in CaM binding affinity for the channel and a structural rearrangement of the CaM channel complex [13, 28–30]. To test the relative calcium dependence of the association between Cav3.1 and CaM, we immunoprecipitated Cav3.1 in the presence of specific levels of buffered calcium . In this way we could test for potential constitutive binding of CaM in nominally zero calcium, near resting levels of calcium in neurons (100 nM), and in progressively higher but physiological levels of calcium . CaM coimmunoprecipitated with Cav3.1 at 0 and 100 nM levels of calcium from lysates of either rat brain or tsA-201 cells expressing Cav3.1 and CaM (Fig. 1e, f) (Additional file 1: Figure S1). However, the Cav3.1-CaM coimmunoprecipitation was lost in the presence of 5 μM or greater calcium concentrations in both rat brain and tsA-201 cells (Fig. 1e, f; Additional file 1: Figure S1). To test if the loss of a Cav3.1-CaM association reflects a calcium-dependent interaction with CaM we coexpressed in tsA-201 cells Cav3.1 and the calcium binding deficient CaM mutant CaM1234, and found that CaM1234 coimmunoprecipitated with Cav3.1 for all concentrations of calcium tested (Fig. 1g).
These results indicate that the Cav3.1-CaM association occurs preferentially in low or resting levels of calcium, suggesting that Cav3.1 channels are constitutively associated with CaM. In contrast, there is a calcium-dependent dissociation of CaM from the Cav3.1 channel, revealing a process that is distinctly different from that reported between HVA calcium channels and CaM.
Dynamics of Cav3.1 and CaM interactions revealed by FRET
Given that coimmunoprecipitation experiments indicated a calcium-dependent loss of the Cav3.1-CaM band on western blots (Fig. 1e, f), we used FRET to further test the calcium dependence of the Cav3.1-CaM association in live cells (measured in all cases from defined ROIs pooled from 3 to 6 separate experiments). Under normal conditions, tsA-201 cells display a depolarized membrane potential that leads to a voltage-dependent inactivation of Cav3.1 calcium channels. To allow these cells to rest at a more hyperpolarized potential we coexpressed cDNA for the inward rectifying potassium channel Kir2.1 . Coexpressing Kir2.1 cDNA (0.3 μg) with GFP-Cav3.1 and mKate-CaM promoted a resting potential of −57.2 mV ± 2.8 mV in 1 mM [K]o, and a depolarization to −24.1 mV ± 5.8 mV (n = 3) in the presence of 50 mM [K]o. Modeling studies of the degree of calcium accumulation near the mouth of a Cav3 calcium channel pore indicate peak values as high as 50 μM . We repeated these studies for Cav3 current parameters recorded from cerebellar Purkinje cells and previously modeled with respect to activation of BK potassium channels . Here the model reports an increase in predicted [Ca]i to >35 μM at 20 nm distance, and 7 μM at 40 nm distance from the channel pore (Additional file 2: Figure S2) (model parameters as described previously) . This is important in validating the potential for Cav3 channels to provide the calcium necessary to promote CaM dissociation given that the coimmunoprecipitation between Cav3.1 and CaM was lost at calcium concentrations of 5 μM or more (Fig. 1e, f). Using this protocol we used the degree to which 50 mM [K]o evoked a Cav3-mediated calcium influx that could promote a loss of FRET between GFP-Cav3.1 and mKate-CaM.
Cav3.1 channel-mediated calcium influx activates αCaMKII
CaM is known to activate CaMKII to act as a second messenger involved in a wide range of functions that include gene transcription to synaptic plasticity [15, 16, 19]. Given indications that calcium influx through Cav3 channels reduced the association between Cav3.1-CaM detected at rest, we explored the potential for CaM to activate CaMKII in response to a depolarizing stimulus.
Our ability to visualize GFP-αCaMKII aggregate formation during live cell imaging allowed us to test calcium-dependent events that lead to αCaMKII activation. One potential source for voltage-gated calcium entry that triggers αCaMKII activation is that of Cav1.x L-type calcium channels . Although HVA calcium channels were not expressed in these experiments we conducted several controls to ensure that calcium influx was restricted to Cav3.1 channels in tsA-201 cells. First, cells expressing GFP-αCaMKII without Cav3.1 coexpression did not exhibit GFP-αCaMKII aggregation (Fig. 4e, f). Second, expressing a Cav3.1 pore mutant together with GFP-αCaMKII and CaM in tsA-201 cells fully blocked the high [K]o-evoked GFP-αCaMKII aggregation (Additional file 5: Figure S5c, d), despite the ability for the Cav3.1 pore mutant to conduct calcium (Additional file 3: Figure S3). We also confirmed that calcium influx was restricted to Cav3.1 channels by blocking L-type calcium channels with 30 μM Cd2+ (Additional files 5 and 6: Figures S5e-h and S6), and by finding that the high [K]o-induced aggregation of GFP-αCaMKII was blocked by 1 μM mibefradil and 300 μM Ni2+ (Additional file 5: Figure S5g, h). High [K]o-evoked GFP-αCaMKII aggregation was further blocked by 0.1 mM BAPTA-AM (Additional file 5: Figure S5i, j), indicating a requirement for an increase in [Ca]i subsequent to Cav3.1 channel activation. Together these data strongly suggest that all voltage-gated calcium influx that lead to αCaMKII activation in these experiments was conducted by Cav3.1 channels.
The dependence of GFP-αCaMKII aggregation upon calcium interactions with CaM was established by a loss of aggregate formation in high [K]o when CaM expression was substituted with the EF hand mutant CaM1234 (Fig. 4g, h). The activation of GFP-αCaMKII is known to require Ca2+/CaM to bind to the autoregulatory domain of αCaMKII to promote autophosphorylation and self-association of the αCaMKII holoenzymes into aggregates [17, 18]. αCaMKII activity and aggregation can also be inhibited through expression of CaMKIIN, a peptide that binds to the catalytic pocket of αCaMKII [24, 34]. To test if the depolarization-induced formation of GFP-αCaMKII aggregates depended on αCaMKII phosphorylation, we expressed GFP-αCaMKII and CaMKIIN in tsA-201 cells coexpressing Cav3.1 and CaM. These tests revealed that CaMKIIN blocked formation of GFP-αCaMKII aggregates following exposure to high [K]o (Fig. 4i, j). The outcome of these tests establish that a membrane depolarization that triggers Cav3 channel-mediated calcium influx activates αCaMKII through a calcium-dependent interaction with CaM and activation of αCaMKII.
Cav3.1 calcium activation of αCaMKII depends on an intact Cav3.1 C-terminus
Cav3 calcium influx triggers αCaMKII phosphorylation in neurons
To complement the immunolabeling tests we further examined pαCaMKII protein density in cerebellar tissue slices exposed to low [K]o or high [K]o in the presence or absence of mibefradil/Ni2+-containing medium. Tissue was lysed in a medium containing the same final solution and prepared for Western blot analysis. In each case a band for pαCaMKII was detected, but the ratio of band densities (pαCaMKII/αCaMKII) was significantly greater for tissue exposed to high [K]o (n = 3; p < 0.05), a difference that was not present in tissues preexposed to mibefradil/Ni2+ (Fig. 7h).
CaM is known to establish an association with HVA calcium channels to provide feedback regulation of channel activity or downstream activation of CaMKII. The current study is important in showing that the Cav3.1 calcium channel exhibits a novel calcium-dependent interaction with CaM that also triggers CaM-dependent activation and phosphorylation of αCaMKII. Moreover, Cav3-mediated interactions can be detected in both tsA-201 cells as well as hippocampal cerebellar neurons.
CaM association with Cav3.1 channels
The complexity and wealth of interactions that define CaM binding and modulation of HVA calcium channels have emerged over many years of work. An HVA calcium channel (e.g. Cav1.x) generally forms a pre-association with calcium free ApoCam at resting levels of internal calcium concentration, and a calcium-dependent increase in CaM binding affinity [13, 28–30]. The assays conducted here reveal that Cav3.1 channels exhibit a constitutive association with ApoCam at resting levels of calcium. The occurrence of FRET between Cav3.1 and CaM at resting levels of internal calcium confirms a close protein-protein interaction given that FRET requires two molecules to be positioned within 10–100 Å distance. CaM binding to HVA calcium channels involves at least an IQ domain on the calcium channel C-terminus, as well as an NSCaTE domain on the N-terminus [13, 35–40]. The amino acid sequence of the Cav3.1 C-terminus does not appear to present a clear IQ domain as defined for HVA channels, nor does it exhibit a canonical CaM binding motif. Our work establishes that the Cav3.1 C-terminus is required to detect a coimmunoprecipitation between CaM and Cav3.1, with GST pull down assays supporting a direct interaction between CaM and the C-terminal region. The precise amino acid residues on the C-terminus responsible for the Cav3.1-CaM interaction remain to be determined.
The Cav3.1-CaM association is further distinguished from CaM-HVA calcium channel interactions in that membrane depolarization promotes a loss of FRET between Cav3.1 and CaM. We note that a loss of FRET could reflect a repositioning of CaM on the Cav3.1 channel, however, the loss of coIP between Cav3.1 and CaM at [Ca] greater than 1 μM indicates a physical dissociation. Moreover, Cav3.1 calcium influx proves to be sufficient to promote Cav3.1-CaM dissociation, as established by preventing the depolarization-induced loss of FRET using Cav3 calcium channel blockers or by expressing a non calcium-conducting Cav3.1 channel pore mutant. These findings are important in revealing a dynamic interaction between Cav3.1 and CaM that is essentially opposite to that of HVA calcium channels.
Calcium influx can activate multiple forms of CaMKII that translocate to perinuclear compartments [15, 18, 41–47], with the current work addressing αCaMKII. Activation of αCaMKII has been traced to calcium influx via specific HVA calcium channels in different cell types [15, 17, 18, 24, 44]. The current study is the first to reveal an interaction between Cav3.1 calcium influx and subsequent activation of αCaMKII. An important question is the extent to which αCaMKII is activated specifically by CaM molecules that are preassociated with Cav3.1 under resting conditions and dissociate upon channel opening, as opposed to CaM activated in the cytoplasm subsequent to a Cav3.1-mediated increase in [Ca]i. A role for CaM that dissociates from the Cav3.1 channel was supported by our finding that expression of a Cav3.1 mutant lacking the C-terminus (removing the CaM binding site) blocked depolarization-induced GFP-αCaMKII aggregation despite an equivalent elevation of [Ca]i as found with intact Cav3.1 channels (Fig. 5). Others have shown that pH can augment calcium-dependent clustering of GFP-αCaMKII  so it is possible that this could contribute to our effects. But the current study establishes that calcium influx through the Cav3 channel class can initiate the steps that lead to an aggregation of GFP-αCaMKII that is fully reversible.
In neurons, synaptic stimulation induces long term changes in synaptic efficacy that involves CaMKII activation and translocation to perinuclear or synaptic regions [17, 46, 48, 49]. In this regard we found that exposure to high [K]o invoked a substantial increase in the number of GFP-αCaMKII aggregates in the cytoplasm of hippocampal neuronal cultures, and of pαCaMKII labeling in processes of cultured hippocampal neurons and in cerebellar Purkinje cell bodies. Moreover, all these effects were blocked by exposure to Cav3 channel blockers. These data verify that Cav3 channel-triggered activation of αCaMKII occurs in neuronal circuits known to exhibit long-term plasticity of synaptic function, including Cav3-mediated forms of LTP in cerebellar neurons [50, 51].
Thus, the sequence of events identified for Cav3.1-CaM-αCaMKII interactions differs from that of HVA calcium channels, revealing separate pathways to activate αCaMKII. A Cav3-initiated process is also distinct in that Cav3.1 channels can be activated over a larger voltage range than HVA calcium channels. Cav3.1-mediated activation αCaMKII may then be prominent in certain cell types or brain regions where these channels regulate cell excitability at even subthreshold potentials. However, we note that αCaMKII activation by way of Cav3.1 or HVA channel calcium influx is not mutually exclusive. Rather, we interpret these results as evidence that Cav3 calcium influx is sufficient to activate αCaMKII through a novel interaction with CaM, providing a means to complement the role for HVA calcium channels in triggering αCaMKII activation.
We gratefully acknowledge M. Kruskic, L. Chen, S. Hameed, J. Hameed, S. Srinivasan, W. Chen, and F. Hiess for expert technical assistance, and R. Rehak for early experimental work. This manuscript is dedicated to the memory of Dr. Shahid Hameed who left a long lasting impact on the Zamponi, Turner, and Stys labs.
This work was supported by grants from the Canadian Institutes of Health Research (R.W.T., G.W.Z., P.K.S.) and National Science and Engineering Research Council (G.W.Z.). Trainee support was provided by an Alberta Innovates - Health Solutions (AI-HS) studentship (A.P.R., B.A.S) and Postdoctoral Fellowship (F-X. Z.), and a Queen EII Scholarship (A.P.R.). R.W.T., P.K.S. and G.W.Z. are AI-HS Scientists and P.K.S. and G.W.Z. hold Canada Research Chairs.
Availability of data and materials
The datasets used and/or analysed during the current study available from the corresponding author on reasonable request.
Protein biochemical and immunocytochemical tests were conducted by HA and APR, imaging by HA and IM, tissue slice experiments by APR and HA, and modeling by JDTE. cDNA constructs were prepared by HA, FZ, and BAS, PKS consulted on imaging experiments, provided reagents, and wrote the ImageTrak software, HA, IM, RWT, GWZ wrote the manuscript. RWT and GWZ supervised the study. All authors read and approved the final manuscript.
The animal procedures used in this study were reviewed and approved by the University of Calgary Animal Care Committee according to the guidelines of the Canadian Council of Animal Care (ACC14–0207).
Consent for publication
All authors have read the current work and consent for its publication.
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Simms BA, Zamponi GW. Neuronal voltage-gated calcium channels: structure, function, and dysfunction. Neuron. 2014;82(1):24–45.View ArticlePubMedGoogle Scholar
- Lambert RC, Bessaih T, Crunelli V, Leresche N. The many faces of T-type calcium channels. Pflugers Arch. 2014;466(3):415–23.View ArticlePubMedGoogle Scholar
- Cribbs L. T-type calcium channel expression and function in the diseased heart. Channels (Austin). 2010;4(6):447–52.View ArticleGoogle Scholar
- Mesirca P, Torrente AG, Mangoni ME. Functional role of voltage gated Ca(2+) channels in heart automaticity. Front Physiol. 2015;6:19.View ArticlePubMedPubMed CentralGoogle Scholar
- Weiss N, Hameed S, Fernandez-Fernandez JM, Fablet K, Karmazinova M, Poillot C, Proft J, Chen L, Bidaud I, Monteil A, et al. A Ca(v)3.2/syntaxin-1A signaling complex controls T-type channel activity and low-threshold exocytosis. J Biol Chem. 2012;287(4):2810–8.View ArticlePubMedGoogle Scholar
- Jacus MO, Uebele VN, Renger JJ, Todorovic SM. Presynaptic Cav3.2 channels regulate excitatory neurotransmission in nociceptive dorsal horn neurons. J Neurosci. 2012;32(27):9374–82.View ArticlePubMedPubMed CentralGoogle Scholar
- Garcia-Caballero A, Gadotti VM, Stemkowski P, Weiss N, Souza IA, Hodgkinson V, Bladen C, Chen L, Hamid J, Pizzoccaro A, et al. The deubiquitinating enzyme USP5 modulates neuropathic and inflammatory pain by enhancing Cav3.2 channel activity. Neuron. 2014;83(5):1144–58.View ArticlePubMedGoogle Scholar
- Carabelli V, Marcantoni A, Comunanza V, de Luca A, Diaz J, Borges R, Carbone E. Chronic hypoxia up-regulates alpha1H T-type channels and low-threshold catecholamine secretion in rat chromaffin cells. J Physiol. 2007;584(Pt 1):149–65.View ArticlePubMedPubMed CentralGoogle Scholar
- Coutelier M, Blesneac I, Monteil A, Monin ML, Ando K, Mundwiller E, Brusco A, Le Ber I, Anheim M, Castrioto A, et al. A recurrent mutation in CACNA1G alters Cav3.1 T-type Calcium-Channel conduction and causes autosomal-dominant cerebellar ataxia. Am J Hum Genet. 2015;97(5):726–37.View ArticlePubMedPubMed CentralGoogle Scholar
- Morino H, Matsuda Y, Muguruma K, Miyamoto R, Ohsawa R, Ohtake T, Otobe R, Watanabe M, Maruyama H, Hashimoto K, et al. A mutation in the low voltage-gated calcium channel CACNA1G alters the physiological properties of the channel, causing spinocerebellar ataxia. Mol Brain. 2015;8:89.View ArticlePubMedPubMed CentralGoogle Scholar
- Zamponi GW. Targeting voltage-gated calcium channels in neurological and psychiatric diseases. Nat Rev Drug Discov. 2016;15(1):19–34.View ArticlePubMedGoogle Scholar
- Ben-Johny M, Yue DT. Calmodulin regulation (calmodulation) of voltage-gated calcium channels. J Gen Physiol. 2014;143(6):679–92.View ArticlePubMedPubMed CentralGoogle Scholar
- Pitt GS, Zuhlke RD, Hudmon A, Schulman H, Reuter H, Tsien RW. Molecular basis of calmodulin tethering and Ca2+−dependent inactivation of L-type Ca2+ channels. J Biol Chem. 2001;276(33):30794–802.View ArticlePubMedGoogle Scholar
- Hudmon A, Schulman H, Kim J, Maltez JM, Tsien RW, Pitt GS. CaMKII tethers to L-type Ca2+ channels, establishing a local and dedicated integrator of Ca2+ signals for facilitation. J Cell Biol. 2005;171(3):537–47.View ArticlePubMedPubMed CentralGoogle Scholar
- Wheeler DG, Barrett CF, Groth RD, Safa P, Tsien RW. CaMKII locally encodes L-type channel activity to signal to nuclear CREB in excitation-transcription coupling. J Cell Biol. 2008;183(5):849–63.View ArticlePubMedPubMed CentralGoogle Scholar
- Lisman J, Yasuda R, Raghavachari S. Mechanisms of CaMKII action in long-term potentiation. Nat Rev Neurosci. 2012;13(3):169–82.PubMedPubMed CentralGoogle Scholar
- Hudmon A, Lebel E, Roy H, Sik A, Schulman H, Waxham MN, De Koninck P. A mechanism for Ca2+/calmodulin-dependent protein kinase II clustering at synaptic and nonsynaptic sites based on self-association. J Neurosci. 2005;25(30):6971–83.View ArticlePubMedGoogle Scholar
- Hudmon A, Schulman H. Neuronal CA2+/calmodulin-dependent protein kinase II: the role of structure and autoregulation in cellular function. Annu Rev Biochem. 2002;71:473–510.View ArticlePubMedGoogle Scholar
- Catterall WA, Leal K, Nanou E. Calcium channels and short-term synaptic plasticity. J Biol Chem. 2013;288(15):10742–9.View ArticlePubMedPubMed CentralGoogle Scholar
- Lee JH, Daud AN, Cribbs LL, Lacerda AE, Pereverzev A, Klockner U, Schneider T, Perez-Reyes E. Cloning and expression of a novel member of the low voltage-activated T-type calcium channel family. J Neurosci. 1999;19(6):1912–21.PubMedGoogle Scholar
- Zamponi GW. Calmodulin lobotomized: novel insights into calcium regulation of voltage-gated calcium channels. Neuron. 2003;39(6):879–81.View ArticlePubMedGoogle Scholar
- McKay BE, McRory JE, Molineux ML, Hamid J, Snutch TP, Zamponi GW, Turner RW. Ca(V)3 T-type calcium channel isoforms differentially distribute to somatic and dendritic compartments in rat central neurons. Eur J Neurosci. 2006;24(9):2581–94.View ArticlePubMedGoogle Scholar
- Lai MM, Hong JJ, Ruggiero AM, Burnett PE, Slepnev VI, De Camilli P, Snyder SH. The calcineurin-dynamin 1 complex as a calcium sensor for synaptic vesicle endocytosis. J Biol Chem. 1999;274(37):25963–6.View ArticlePubMedGoogle Scholar
- Flynn R, Labrie-Dion E, Bernier N, Colicos MA, De Koninck P, Zamponi GW. Activity-dependent subcellular cotrafficking of the small GTPase Rem2 and Ca2+/CaM-dependent protein kinase IIalpha. PLoS One. 2012;7(7):e41185.View ArticlePubMedPubMed CentralGoogle Scholar
- Khosravani H, Altier C, Simms B, Hamming KS, Snutch TP, Mezeyova J, McRory JE, Zamponi GW. Gating effects of mutations in the Cav3.2 T-type calcium channel associated with childhood absence epilepsy. J Biol Chem. 2004;279(11):9681–4.View ArticlePubMedGoogle Scholar
- Christensen PC, Welch NC, Brideau C, Stys PK. Functional ionotropic glutamate receptors on peripheral axons and myelin. Muscle Nerve. 2016;54(3):451–9.View ArticlePubMedGoogle Scholar
- Engbers JD, Zamponi GW, Turner RW. Modeling interactions between voltage-gated Ca (2+) channels and KCa1.1 channels. Channels (Austin). 2013;7(6):524–9.View ArticleGoogle Scholar
- Ben Johny M, Yang PS, Bazzazi H, Yue DT. Dynamic switching of calmodulin interactions underlies Ca2+ regulation of CaV1.3 channels. Nat Commun. 2013;4:1717.View ArticlePubMedPubMed CentralGoogle Scholar
- Erickson MG, Alseikhan BA, Peterson BZ, Yue DT. Preassociation of calmodulin with voltage-gated Ca(2+) channels revealed by FRET in single living cells. Neuron. 2001;31(6):973–85.View ArticlePubMedGoogle Scholar
- Erickson MG, Liang H, Mori MX, Yue DT. FRET two-hybrid mapping reveals function and location of L-type Ca2+ channel CaM preassociation. Neuron. 2003;39(1):97–107.View ArticlePubMedGoogle Scholar
- Augustine GJ, Santamaria F, Tanaka K. Local calcium signaling in neurons. Neuron. 2003;40(2):331–46.View ArticlePubMedGoogle Scholar
- Choi J, Park JH, Kwon OY, Kim S, Chung JH, Lim DS, Kim KS, Rhim H, Han YS. T-type calcium channel trigger p21ras signaling pathway to ERK in Cav3.1-expressed HEK293 cells. Brain Res. 2005;1054(1):22–9.View ArticlePubMedGoogle Scholar
- Lacinova L, Kurejova M, Klugbauer N, Hofmann F. Gating of the expressed T-type Cav3.1 calcium channels is modulated by Ca2+. Acta Physiol (Oxf). 2006;186(4):249–60.View ArticleGoogle Scholar
- Vest RS, Davies KD, O'Leary H, Port JD, Bayer KU. Dual mechanism of a natural CaMKII inhibitor. Mol Biol Cell. 2007;18(12):5024–33.View ArticlePubMedPubMed CentralGoogle Scholar
- Pate P, Mochca-Morales J, Wu Y, Zhang JZ, Rodney GG, Serysheva II, Williams BY, Anderson ME, Hamilton SL. Determinants for calmodulin binding on voltage-dependent Ca2+ channels. J Biol Chem. 2000;275(50):39786–92.View ArticlePubMedGoogle Scholar
- Tang W, Halling DB, Black DJ, Pate P, Zhang JZ, Pedersen S, Altschuld RA, Hamilton SL. Apocalmodulin and Ca2+ calmodulin-binding sites on the CaV1.2 channel. Biophys J. 2003;85(3):1538–47.View ArticlePubMedPubMed CentralGoogle Scholar
- Dick IE, Tadross MR, Liang H, Tay LH, Yang W, Yue DT. A modular switch for spatial Ca2+ selectivity in the calmodulin regulation of CaV channels. Nature. 2008;451(7180):830–4.View ArticlePubMedPubMed CentralGoogle Scholar
- Asmara H, Minobe E, Saud ZA, Kameyama M. Interactions of calmodulin with the multiple binding sites of Cav1.2 Ca2+ channels. J Pharmacol Sci. 2010;112(4):397–404.View ArticlePubMedGoogle Scholar
- Kim J, Ghosh S, Nunziato DA, Pitt GS. Identification of the components controlling inactivation of voltage-gated Ca2+ channels. Neuron. 2004;41(5):745–54.View ArticlePubMedGoogle Scholar
- Simms BA, Souza IA, Zamponi GW. A novel calmodulin site in the Cav1.2 N-terminus regulates calcium-dependent inactivation. Pflugers Arch. 2014;466(9):1793–803.View ArticlePubMedGoogle Scholar
- Cohen SM, Li B, Tsien RW, Ma H. Evolutionary and functional perspectives on signaling from neuronal surface to nucleus. Biochem Biophys Res Commun. 2015;460(1):88–99.View ArticlePubMedPubMed CentralGoogle Scholar
- Ma H, Groth RD, Cohen SM, Emery JF, Li B, Hoedt E, Zhang G, Neubert TA, Tsien RW. gammaCaMKII shuttles Ca(2)(+)/CaM to the nucleus to trigger CREB phosphorylation and gene expression. Cell. 2014;159(2):281–94.View ArticlePubMedPubMed CentralGoogle Scholar
- Deisseroth K, Heist EK, Tsien RW. Translocation of calmodulin to the nucleus supports CREB phosphorylation in hippocampal neurons. Nature. 1998;392(6672):198–202.View ArticlePubMedGoogle Scholar
- Li B, Tadross MR, Tsien RW. Sequential ionic and conformational signaling by calcium channels drives neuronal gene expression. Science. 2016;351(6275):863–7.View ArticlePubMedPubMed CentralGoogle Scholar
- Wheeler DG, Groth RD, Ma H, Barrett CF, Owen SF, Safa P, Tsien RW. Ca(V)1 and Ca(V)2 channels engage distinct modes of Ca(2+) signaling to control CREB-dependent gene expression. Cell. 2012;149(5):1112–24.View ArticlePubMedPubMed CentralGoogle Scholar
- Deisseroth K, Bito H, Tsien RW. Signaling from synapse to nucleus: postsynaptic CREB phosphorylation during multiple forms of hippocampal synaptic plasticity. Neuron. 1996;16(1):89–101.View ArticlePubMedGoogle Scholar
- Bading H, Ginty DD, Greenberg ME. Regulation of gene expression in hippocampal neurons by distinct calcium signaling pathways. Science. 1993;260(5105):181–6.View ArticlePubMedGoogle Scholar
- Marsden KC, Shemesh A, Bayer KU, Carroll RC. Selective translocation of Ca2+/calmodulin protein kinase IIalpha (CaMKIIalpha) to inhibitory synapses. Proc Natl Acad Sci U S A. 2010;107(47):20559–64.View ArticlePubMedPubMed CentralGoogle Scholar
- Fink CC, Meyer T. Molecular mechanisms of CaMKII activation in neuronal plasticity. Curr Opin Neurobiol. 2002;12(3):293–9.View ArticlePubMedGoogle Scholar
- Pugh JR, Raman IM. Mechanisms of potentiation of mossy fiber EPSCs in the cerebellar nuclei by coincident synaptic excitation and inhibition. J Neurosci. 2008;28(42):10549–60.View ArticlePubMedPubMed CentralGoogle Scholar
- Ly R, Bouvier G, Schonewille M, Arabo A, Rondi-Reig L, Lena C, Casado M, De Zeeuw CI, Feltz A. T-type channel blockade impairs long-term potentiation at the parallel fiber-Purkinje cell synapse and cerebellar learning. Proc Natl Acad Sci U S A. 2013;110(50):20302–7.View ArticlePubMedPubMed CentralGoogle Scholar