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Table of Contents
Abstract
Introduction
Results And Discussion
Architecture Of The Ca V 2.3 Complex
Functional Heterogeneity Of The Voltage-Sensing Domains
Molecular Mechanism Of Closed-State Inactivation
Modulation Of Open-State Inactivation
Methods
Expression And Protein Purification Of The Human Ca V 2.3 Complex
Cryo-EM Sample Preparation And Data Collection
Cryo-EM Data Processing
Model Building
Whole-Cell Voltage-Clamp Recordings Of Ca V 2.3 Channels In HEK 293-T Cells
Electrophysiological Data Analysis
Supporting Information
Author Contributions
Data Availability
Conflict Of Interest
Competing Interest Statement
Competing Interest Statement
Abstract
High-voltage-activated R-type Ca V 2.3 channel plays pivotal roles in many physiological activities and is implicated in epilepsy, convulsions, and other neurodevelopmental impairments. Here, we determine the high-resolution cryo-electron microscopy (cryo-EM) structure of human Ca V 2.3 in complex with the α2δ1 and β1 subunits. The VSD II is stabilized in the resting state. Electrophysiological experiments elucidate that the conformational change of VSD II in response to variation in membrane potential is not required for channel activation, whereas the other VSDs are essential for channel opening. The intracellular gate is blocked by the W-helix. A pre-W-helix adjacent to the W-helix can significantly regulate closed-state inactivation (CSI) by modulating the association and dissociation of the W-helix with the gate. Electrostatic interactions formed between the negatively charged domain on S6 II , which is exclusively conserved in the Ca V 2 family, and nearby regions at the alpha-interacting domain (AID) and S4-S5 II helix are identified. Further functional analyses indicate that these interactions are critical for the open-state inactivation (OSI) of Ca V 2 channels.
Introduction
Voltage-gated calcium (Ca V ) channels mediate calcium influx to cells in response to changes in membrane potential 1 - 3 . Their cellular roles have been emphasized for decades in a variety of studies and include hormone secretion 4 , 5 , neurotransmitter release 6 , 7 , and muscle contraction 8 , 9 . Ca V channel members are categorized into the Ca V 1, Ca V 2, and Ca V 3 subfamilies based on sequence identity or alternatively classified into T-, L-, P/Q-, N-, and R-types according to their pharmacological and biophysical profiles 1 . The so-called pharmacoresistant (R-type) Ca V 2.3 is widely expressed in the brain and enriched in the hippocampus, cerebral cortex, amygdala, and corpus striatum 10 - 12 . Electrophysiological investigations revealed that currents mediated by Ca V 2.3 are resistant to common Ca V blockers or gating modifiers such as nifedipine, nimodipine, ω-Aga-IVA, etc 13 . Ca V 2.3 channels exhibit cumulative inactivation in response to brief and repetitive depolarizations, a process known as preferential closed-state inactivation (CSI) 14 . Furthermore, Ca V 2.3 is involved in a broad spectrum of neuronal activities 10 , 12 , 15 . Previous studies have reported that Ca V 2.3 participates in multiple physiological processes in the central nervous system, such as inducing long-term potentiation (LTP) and post-tetanic potentiation in mossy fiber synapses 16 , modulating the burst firing mode of action potentials 17 , 18 , and regulating synaptic strength in hippocampal CA1 pyramidal neurons 19 . In recent years, increasing evidence has revealed that dysfunction of Ca V 2.3 is linked to epilepsy 20 , 21 , convulsions 22 , 23 , and neurodevelopmental impairments 24 , suggesting that Ca V 2.3 is a pivotal player in the pathogenesis of a series of neurological disorders. The molecular basis of Ca V channels has been investigated extensively over the past several decades, including structural studies of L-type Ca V 1.1 25 , 26 , N-type Ca V 2.2 27 , 28 , and T-type Ca V 3.1 29 and Ca V 3.3 30 in the apo form or distinct modulator-bound states. These structures provide substantial insights into the architecture, subunit assembly, and modulator actions of the Ca V channels. However, the gating mechanism of Ca V channels is still far from fully understood. For instance, in the Ca V 2.2 structure, the VSD II is trapped in a resting state by a PIP 2 molecule at a membrane potential of ∼0 mV 27 , 28 . The functional roles of the VSD II trapped in the resting state by PIP 2 remain unknown. A considerable number of pathogenic mutations have been identified in the VSDs of neuronal Ca V 2 channels, demonstrating that VSD dysfunctions contribute to the genesis of spinocerebellar ataxia (SCA), episodic ataxia (EA), and familial hemiplegic migraine (FHM) 31 . Moreover, the Ca V 2.2 and Ca V 2.3 channels inactivate preferentially from the intermediate closed state along the activation pathway, which is important in controlling the short-term dynamics of synaptic efficacy 14 , 32 . In our previous study, we elucidated that residue W768 on the W-helix located within the DII-III linker serves as a key determinant of the CSI of the Ca V 2.2 channel 28 . However, Ca V 2.3 is characterized by a more prominent preferential CSI than Ca V 2.2 14 . It is also interesting to explore the modulation mechanism of CSI in Ca V 2.3. Furthermore, the high-voltage-activated (HVA) Ca V 1 and Ca V 2 channels harbor a conserved α-helix connecting Domain I and Domain II (alpha-interaction domain, or AID). Previous studies have indicated that AID might contribute to the open-state inactivation (OSI) of the HVA Ca V channels 33 , 34 . However, the inactivation properties of Ca V 1 and Ca V 2 channels are dramatically different 33 . Mechanistic insight into the inactivation processes of the HVA Ca V channels will help us to fully uncover the physiological role of Ca V channels and facilitate the development of therapeutic solutions for Ca V -related diseases. In this study, we expressed and purified human Ca V 2.3 in complex with auxiliary subunits α2δ1 and β1 and unveiled the high-resolution structure of this protein complex. Further mutagenesis and electrophysiological experiments were performed. Our results provide insights into the pharmacological resistance properties of Ca V 2.3, the asynchronous functional roles of the VSDs, the mechanism by which the pre-W-helix regulates the CSI, and the OSI process modulated by the negatively charged domain on S6 II (S6 II NCD ).
Results and Discussion
Architecture of the Ca V 2.3 complex
To gain structural insights into the Ca V 2.3 complex, we expressed and purified full-length wild-type human Ca V 2.3 α1E subunit (CACNA1E), α2δ1 (CACNA2D1) and β1 (CACB1) using a HEK 293-F expression system. The Ca V 2.3-α2δ1-β1 complex was solubilized using n-Dodecyl-βD-maltoside (DDM) and purified using a strep-tactin affinity column, followed by further purification by size-exclusion chromatography (SEC) in a running buffer containing glycol-diosgenin (GDN) to remove protein aggregates (Supplementary Figure 1a, see Method section for details). The peak fractions were subsequently collected and concentrated for cryo-EM sample preparation (Supplementary Figure 1b). A total of 2,096 micrographs were collected. Data processing of the dataset gave rise to a 3.1-Å cryo-EM map, which is rich in high-resolution structural features, including densities for side chains, lipid molecules and glycosylations ( Figure 1 , Supplementary Figure 2, and Table S1). The Ca V 2.3 complex exhibited a conventional shape that resembles that of the Ca V 2.2 and Ca V 1.1 complexes ( Figure 1 ). The complex is composed of the transmembrane α1E subunit, extracellular α2δ1 subunit, and intracellular β1 subunit ( Figure 1a–1b ). The α1E subunit is a pseudotetrameric pore-forming subunit and can be divided into Domain I (DI) to IV (DIV). Each domain of the α1E subunit is composed of 6 transmembrane helices (S1–S6), comprising the voltage-sensing domain (S1–S4) and the pore domain (S5–S6). The P1 and P2 helices located between the S5 and S6 helices formed the selectivity filter ( Figure 1c ). Similar to other Ca V channels, Ca V 2.3 harbors four extracellular loops (ECLs) that is also positioned between S5 and S6 helices in the pore domain ( Figure 1a–1c ). The ECL I and ECL II are critical for the association between the α1 subunit and the α2δ1 subunit ( Figure 1a–1b ). The S6 helices from the four domains converge on the cytoplasmic side and form the intracellular gate of the channel. In our structure, the intracellular gate was determined in its closed state, in line with the observations from other voltage-gated channels ( Figure 1c ). Moreover, the closed gate of Ca V 2.3 is further stabilized by the W-helix from the DII-DIII linker, which is consistent with a previous study on Ca V 2.2 and indicates that Ca V 2.3 also adopts the CSI mechanism 28 . Most Ca V channels serve as pharmaceutical targets of a variety of small-molecule drugs or peptide toxins 13 . However, previous studies indicate that Ca V 2.3 is resistant to many Ca V modulators, such as nimodipine (L-type), omega-Aga-IVA (P/Q-type), and omega-CTx-GVIA (N-type) 13 . To clarify the structural basis underlying the pharmacoresistance of Ca V 2.3, we compared the structures of Ca V 2.3 with Ca V 1.1 or Ca V 2.2 in their ligand-bound states ( Figure 1d–1e ). In the structure of nifedipine-bound Ca V 1.1, the nifedipine molecule was located within the DIII-DIV fenestration and stabilized by surrounding residues. However, two critical residues are substituted in Ca V 2.3, namely, Y1296 and F1708. The bulky sidechains of these two residues in Ca V 2.3 occupy the DIII-DIV fenestration site and thus hinder the binding of nifedipine ( Figure 1d ). Meanwhile, a previous study reported that the Q1010 of Ca V 1.2 is important for sensitivity to dihydropyridine (DHP) and the Q1010M mutant had a decreased sensitivity to DHP molecules 35 . The equivalent position in Ca V 2.3 is occupied by M1300, thus also contributing to the pharmacoresistance of Ca V 2.3 to DHP molecules. Structural comparison of Ca V 2.3 and ziconotide-bound Ca V 2.2 demonstrated that the ECL I loop of Ca V 2.3 adopts a different conformation, and residues D263 and P264 are placed close to the central axis, giving rise to clashes between the ziconotide and the ECL I of Ca V 2.3 ( Figure 1e ). Moreover, other residues on P-loops and ECLs that are involved in ziconotide binding are also not conserved in Ca V 2.3 (Supplementary Figure 3), rendering Ca V 2.3 insensitive to the ziconotide. Gain-of-function mutations and polymorphisms of Ca V 2.3 channels have already been implicated in the pathological process of developmental and epileptic encephalopathy. Thirteen pathogenic mutations have been identified in Ca V 2.3 24 . Our high-resolution structure provides a structural template to map all of these pathogenic mutations, which are distributed throughout the complex structure ( Figure 1f ). Eight of thirteen mutations are located around the intracellular gate, such as I603L, F698S, and I701V, and result in a hyperpolarizing shift in the half-activation voltage 24 ( Figure 1f ).
Functional heterogeneity of the voltage-sensing domains
The voltage-dependent gating characteristics of voltage-gated channels are conferred by their voltage-sensing domains (VSDs). The VSDs of Ca V are conserved helix bundles consisting of S1, S2, S3 and S4 helices (Supplementary Figure 4a). S4 was found to be a positively charged 3 10 helix, harboring approximately five or six arginines or lysines as gating charges lining one side of the helix at intervals of three residues (Supplementary Figure 4a). The positively charged S4 helices move vertically toward the intracellular or extracellular side of the cell in response to the hyperpolarization or depolarization of the membrane potential. The conformational change of VSD is coupled to the pore domain by a short amphipathic helix S4-S5, which connects the S4 helix of VSD to the S5 helix from the pore domain, thus regulating the transition of the intracellular gate between the open and closed states. Although the four VSDs of Ca V channels are considerably similar in terms of sequence and overall structure, they contribute differentially to the opening of pore 36 . Superimposition of the structures of Ca V 2.3 and Ca V 2.2 revealed that they are comparable overall (r.m.s.d. = 1.46 Å for 2,222 Cα atom pairs). The pore domain was fairly superimposable between Ca V 2.3 and Ca V 2.2, including the S5 and S6 helices and extracellular loops (ECLs) I, III and III (Supplementary Figure 4b). VSD I , VSD III , and VSD IV in the activated state and VSD II in the resting state were also determined in both Ca V 2.3 and Ca V 2.2 (Supplementary Figure 4a–4b). However, a structural discrepancy was visualized at the ECL IV between the two structures. The ECL IV of Ca V 2.3 extends from the pore domain and lies above the S1-S2 III linker, whereas the ECL IV of Ca V 2.2 is much shorter and wraps around the pore domain before touching the extracellular side of VSD III ( Figure 2a ). Four residues on ECL IV of Ca V 2.3, namely, P1680, D1681, T1682, and T1683, are involved in the interactions with the residues on the S1-S2 III linker, especially V1176, L1177, T1178, and N1179, which consequently stitches the voltage sensing domain to the pore domain at the extracellular side ( Figure 2a ). To explore the functional roles of this interaction, we substituted 1680 PDTT 1683 on ECL IV with four glycines (Ca V 2.3 4G ) to disrupt the contacts between ECL IV and S1-S2 III loop. Electrophysiological studies indicated that the voltage dependency of the activation curve of Ca V 2.3 4G displayed a ∼5-mV positive shift (P < 0.0001, two-tailed unpaired t -test) compared to that of wild-type Ca V 2.3 ( Figure 2b and Supplementary Figure 5). We thus speculate that the interactions between ECL IV and the S1-S2 III loop may stabilize the VSD III in a certain conformation relative to the pore domain that requires less electrical energy to activate the channel, reminiscent of the cholesterol regulation on Ca V channels, potentially by stabilizing interactions between the extracellular end of the S1-S2 helix hairpin and the pore domain 37 , 38 . The VSD II s of Ca V 2.3 and Ca V 2.2 were determined in the resting (S4 down) state, while most other VSDs in the voltage-gated channels, such as Ca V 1.1, Ca V 3.1, and Na V s, were determined in the activated (S4 up) state (Supplementary Figure 4c–4d). This finding suggests that VSD II plays a unique role in the gating mechanism among Ca V 2 members. Structural analyses of Ca V 2.2 have suggested that the VSD II is trapped in the resting state by a PIP 2 molecule 27 , 28 . The cryo-EM map of Ca V 2.3 is of high quality around S4-S5 II (Supplementary Figure 2e), and we identified a density lying above the S4-S5 II helix that serves as a wedge to hinder the upward movement of the S4 helix. However, the density in Ca V 2.3 appears to be a single strip-shaped density and does not look like a PIP 2 molecule. To shed light on the functional roles of VSD II in the gating mechanism of Ca V 2.3, we constructed a gating charge neutralization mutant on the VSD II (VSD II4Q , R572Q/R575Q/R578Q/K581Q). Interestingly, the neutralization mutation of the VSD II (VSD II4Q ) exhibits ∼9-mV left shifts in the voltage dependency of both activation and steady-state inactivation compared to the wild-type ( Figure 2b and Supplementary Figure 5a–5b). Consequently, the current density-voltage curve of the VSD II 4Q mutant was also left shifted ( Figure 2c ). Moreover, we tested whether the VSD II 4Q mutant has a distinct CSI profile. The cumulative inactivation of this mutant in response to action potential (AP) trains is markedly enhanced (Supplementary Figure 5c and 5e). These results suggested that the conformation of VSD II influences the gating of Ca V 2.3. However, its OSI kinetics remain unaltered (Supplementary Figure 5d). In contrast, the gating charge-neutralized mutation in the VSD I , VSD III , and VSD IV resulted in failure to mediate inward current ( Figure 2d ), in line with previous results showing that the VSD I , VSD III , and VSD IV are important for gating of the closely-related Ca V 2.2 39 - 41 , while the VSD II is not necessary for channel activation by sensing the depolarization of membrane potential; instead, the VSD II is crucial to modulating channel properties, such as CSI and voltage dependency of channel activation and inactivation.
Molecular mechanism of closed-state inactivation
Preferential closed-state inactivation is a featured kinetic characteristic of neuronal voltage-gated calcium channels 14 , 28 . During the state-transition pathway in the activation of Ca V channels, CSI occurs preferentially in a specific pre-open closed state and a voltage-dependent manner. CSI can be visualized by the cumulative inactivation in response to action potential (AP) trains, as reported in previous studies, which demonstrated that the peak current triggered by each AP shrank sequentially, suggesting that a substantial amount of the channels turn inactivated after the repolarization of an AP. CSI plays an important role in the orchestrated modulation mechanism of Ca V channels and is of vital importance for the precise regulation of physiological processes such as neurotransmitter release and synapse plasticity. CSI is detected in all neuronal Ca V 2 channels at distinct levels 14 . The R-type Ca V 2.3 displayed a more prominent CSI than the N-type Ca V 2.2, and both showed far more prominent CSI than the P/Q-type Ca V 2.1 14 . Previous structural investigations on the N-type Ca V 2.2 channel have revealed that a conserved W-helix is the structural determinant of CSI, and W768 from the W-helix on the DII-DIII linker functions as a lid blocking the pore and stabilizes the intracellular gate in its closed states by hydrophobic interactions. However, the molecular mechanisms underlying the CSI of Ca V s are still not fully understood, as the conserved W-helix is unable to explain the disparity of CSI mechanisms among the Ca V 2.1, Ca V 2.2, and Ca V 2.3 channels. The W-helix determined in Ca V 2.2 is also conserved and well resolved in Ca V 2.3 ( Figure 3a–3b ). The W-helix in Ca V 2.3 ( 772 RHHMSVWEQRTSQLRKH 788 ) is a positively charged short helix and positioned underneath the intracellular gate, with W778 inserting into the gate and forming extensive interactions with residues from surrounding gating helices ( Figure 3a–3b ). These structural observations are consistent with the W-helix of Ca V 2.2. First, we designed the Ca V 2.3 W/Q (W778Q) to disrupt interactions between the W-helix and intracellular gate ( Figure 3c–3g and Supplementary Figure 6). It turns out that the Ca V 2.3 W/Q exhibited a ∼8-mV positive shift on the steady-state inactivation curve ( Figure 3d ) and an alleviated cumulative inactivation in response to AP trains ( Figure 3e and 3g ) without affecting the voltage dependence of channel activation ( Figure 3c ), consistent with the observations in Ca V 2.2 28 , suggesting that the W778 is important for CSI process of Ca V 2.3 channel. Moreover, we speculate that the positively charged residues on the W-helix could putatively respond to the membrane potential change and may be important for initiation of the CSI process. To evaluate our speculations, we constructed the Ca V 2.3 RK/A mutant by substituting the R781, R786 and K787 with alanine (R781A/R786A/K787A). The activation curve of the Ca V 2.3 RK/A mutant remains unaltered ( Figure 3c ). However, the CSI of Ca V 2.3 RK/A mutant was significantly suppressed, exhibiting a ∼9 mV right shift on the steady-state inactivation curve ( Figure 3d ), an alleviated cumulative inactivation to AP trains ( Figure 3e and 3g ), and an accelerated recovery rate from CSI ( Figure 3f and Supplementary Figure 6b). These alterations on the CSI profile suggested that the positively charged R781, R786 and K787 are critical for the CSI mechanism of Ca V 2.3. Intriguingly, sequence alignment among the neuronal Ca V s revealed that a peptide segment (pre-W-helix) that resembled the W-helix is located adjacent to the W-helix of Ca V 2.3 ( Figure 4a ). The sequence of pre-W-helix ( 753 RHHMSMWEPRSSHLRER 769 ) is nearly identical to that of the W-helix (∼65% identity), including the conserved tryptophan plug (W759) and positively charged residues (R762, R767, and R769) ( Figure 4a ). However, we noticed that a non-conserved proline (P761) was in the middle of the pre-W-helix, which may undermine the stability of both the pre-W-helix itself and its interactions with the gate. Moreover, in our cryo-EM map of the Ca V 2.3 complex, the helical density beneath the gate perfectly fits the atomic model of the W-helix, enabling us to unambiguously determine that the W-helix, instead of the pre-W-helix, exists in our structure (Supplementary Figure 2e). Considering that the pre-W-helix is located immediately before the W-helix and that they share high sequence identity, we speculate that the pre-W-helix participates in the CSI event of Ca V 2.3. To evaluate the contribution of the pre-W-helix to the CSI of Ca V 2.3, we constructed two mutants by deleting the W-helix (Δ w-helix ) and pre-W-helix (Δ pre-w-helix ) ( Figure 4b–4e , 4k , and Supplementary Figure 6). The Δ w-helix exhibited a ∼4-mV positive shift on the steady-state inactivation curve ( Figure 4c ) and alleviated cumulative inactivation in response to AP trains ( Figure 4d and 4k ) without affecting the voltage dependence of channel activation ( Figure 4b ), indicating that the W-helix plays pivotal roles in the CSI of Ca V 2.3. However, compared with the CSI of Ca V 2.2, which was almost abolished by deleting the W-helix, a substantial portion of the CSI in the Δ w-helix mutant remained unaltered ( Figure 4d and 4k ), suggesting that the CSI modulation of Ca V 2.3 is distinct from that of Ca V 2.2 and that other elements may also contribute to the CSI of Ca V 2.3. We also used a two-pulse protocol to assess the recovery rate from CSI, i.e., the release process of CSI (Supplementary Figure 6a). Consistent with the electrophysiological results above, an accelerated recovery rate from CSI was observed in the Δ w-helix compared to the WT ( Figure 4e and Supplementary Figure 6b). Strikingly, a negative-shift of ∼8 mV was detected on the inactivation curve of the Δ pre-w-helix mutant ( Figure 4c ), and its cumulative inactivation to AP trains was surprisingly enhanced ( Figure 4d and 4k ), demonstrating that the development process of CSI in the Δ pre-w-helix was significantly boosted. Considering that the pre-W and W helices are close to each other and share high sequence identity, we speculate that the pre-W-helix may serve as a competitive negative regulator to interfere with the binding of the intracellular gate to the W-helix ( Figure 4f ). In the absence of the pre-W-helix, the CSI is consequently enhanced ( Figure 4c–4d , and 4k ). Paradoxically, compared with the WT, the recovery rate from CSI of the Δ pre-w-helix mutant was substantially accelerated ( Figure 4e and Supplementary Figure 6b). This suggests that once CSI occurs, the pre-W-helix appears to stabilize the W-helix for interaction with the gate during membrane potential repolarization, thereby slowing the recovery of Ca V 2.3 from the CSI. We also designed a mutant by deleting both the pre-W-helix and the W-helix (Δ pre-w /Δ w-helix ). This mutant displayed a ∼5-mV positive shift on the inactivation curve ( Figure 4c ), reduced cumulative inactivation to AP trains ( Figure 4d and 4k ), and an accelerated recovery rate from CSI ( Figure 4e and Supplementary Figure 6b). These effects of this double deletion construct are essentially identical to those of the Δ w-helix , demonstrating that the modulatory role of the pre-W-helix on CSI is largely dependent on the W-helix, probably by regulating the binding or dissociation of the W-helix with the intracellular gate. To further investigate the regulatory mechanism on the CSI of Ca V 2.3, we constructed two mutants by neutralizing the arginines (Ca V 2.3 preR/Q , R753Q/R762Q/R767Q/R769Q) or substituting the tryptophan on the pre-W-helix with glutamine (Ca V 2.3 preW/Q , W759Q) ( Figure 4g–4k and Supplementary Figure 6). Impressively, the mutants Ca V 2.3 preR/Q and Ca V 2.3 preW/Q exhibit similar gating kinetics and voltage dependence of channel activation and inactivation, but significantly different from that of the WT Ca V 2.3 channel ( Figure 4g–4k , Supplementary Figure 5a–5b and Supplementary Figure 6b). In particular, they displayed a ∼7-mV negative shift on inactivation curve ( Figure 4h ), enhanced cumulative inactivation to AP trains ( Figure 4i and 4k ), and an accelerated recovery rate from CSI ( Figure 4j and Supplementary Figure 6b), highly identical to the CSI profile of the Δ pre-w-helix . The kinetic characteristics of these mutants indicated that the positively charged residues and W759 are important for the regulatory effect of the pre-W-helix. Our data also show that the development and release of CSI are two independent processes. In particular, the Δ pre-w-helix , Ca V 2.3 preR/Q and Ca V 2.3 preW/Q exhibited enhanced cumulative inactivation during AP trains ( Figure 4d , 4i and 4k ) but a faster recovery rate from CSI ( Figure 4e , 4j and Supplementary Figure 6b) at a membrane potential of –100 mV, suggesting that the channel may have distinct conformational states along the activation pathway. The W-helix may bind preferentially to the channel in the intermediate closed state(s) near the open state, thus leading to the largest inactivation before the channel opens ( Figure 4f ). At strongly hyperpolarized potentials (∼100 mV), the channel is probably stabilized in a different closed state, exhibiting a lower affinity to the W-helix, causing the W-helix to detach from the gate and the channel to recover from the CSI ( Figure 4f ). Taken together, the pre-W-helix in Ca V 2.3 plays significant roles in regulating the association or dissociation of the W-helix with the intracellular gate and thus exerts a regulatory effect on the development and release of the CSI. Interestingly, on the genomic DNA of Ca V 2.3, the pre-W-helix is encoded by exon 19 (residues R748–R769) ( Figure 4a ). Early studies have reported that alternative splicing events might occur at this site, as exon 19 is spliced out in the mature mRNA encoding Ca V 2.3e, an isoform of Ca V 2.3 that displays decreased Ca 2+ sensitivity in its calcium-dependent modulation mechanisms 42 . Gene expression profiling has revealed that Ca V 2.3e is enriched in endocrine tissues, including the kidney and pancreas, and the majority of Ca V 2.3 in the brain contains the pre-W-helix 43 . This implies that the kinetic variability of Ca V 2.3 mediated by the pre-W-helix is an important regulatory mechanism to fine-tune the properties of the Ca V 2.3 channel to adapt to the distinct physiological needs of neuronal and endocrinal excitable cells 42 .
Modulation of open-state inactivation
Upon the opening of the pore, Ca V channels undergo an inactivation process called open-state inactivation (OSI), which describes the mechanism by which the intracellular gate shifts swiftly from the open state into the inactivation state 33 , 44 . This process is an intrinsic mechanism that precisely regulates calcium influx into cells during depolarization. Neuronal (P/Q-, N- and R-type) Ca V 2 channels bear a stronger OSI and mediate a rapidly inactivated current, while cardiac (L-type) Ca V channels display a much weaker OSI, mediating a long-lasting current 45 . Dysfunction of OSI, i.e., the gain-of-function of neuronal Ca V s, is linked to a series of neurological disorders, including trigeminal neuralgia 46 , myoclonus-dystonia-like syndrome 47 , and epileptic encephalopathies 21 , 24 . Although the structures of L-type Ca V 1.1 25 and N-type Ca V 2.2 27 , 28 complexes have been elucidated at high resolution, the structural basis for their distinct OSI properties remains elusive. Intriguingly, when comparing the EM density maps of the Ca V 1.1, Ca V 2.2, and Ca V 2.3 complexes, we observed that AID displayed a blurred density in Ca V 1.1 but was clearly resolved in Ca V 2.2 and Ca V 2.3 (Supplementary Figure 7). Previous studies suggested that the AID helix is able to regulate the OSI in Ca V channels 33 , 34 . In the structure of Ca V 2.3, the S6 II helix is much longer than that of Ca V 1.1 and extends into the cytosol. Interestingly, we identified a negatively charged domain 715 DEQEEEE 721 in the intracellular juxtamembrane region of the S6 II helix (S6 IINCD ) ( Figure 5a–5b ). Taking a closer look at the structure, this negatively charged region forms electrostatic interactions with the positively charged R590 on S4-S5 II , as well as R371 and R378 on the AID ( Figure 5b ). Strikingly, the S6 IINCD is conserved among P/Q-, N- and R-type channels ( Figure 5c ). The equivalent segment of L-type Ca V channels contains fewer negatively charged residues, interspersed with some positively charged residues, indicating that the electrostatic interactions between S6 IINCD and AID are not present in the L-type Ca V channels, which is consistent with structural observations that the AID of Ca V 1.1 has high motility ( Figure 5c and Supplementary Figure 7). Additionally, the positively charged R590 and R378 are conserved only in Ca V 2 channels. R378 is further reverted to a negatively charged glutamate in the Ca V 1 subfamily, reflecting the coevolutionary linkages within this interaction site ( Figure 5c ). We speculate that the charge interactions centered on S6 II NCD is critical to regulating the OSI of Ca V 2 channels. To validate our hypothesis, we mutated the key residues to disrupt electrostatic interactions cross-linking the S6 II , S4-S5 and AID helices, including replacing 715 DEQEEEE 721 with 715 NNQNNNN 721 (Ca V 2.3 NQ ), R590Q, R378Q/E, and R371Q/E ( Figure 5d and Supplementary Figure 8). We employed Ba 2+ as the charge carrier in our whole-cell patch clamp analysis to exclude the effects of calcium-dependent inactivation (CDI) of the Ca V 2.3 channels. The Ca V 2.3 NQ mutant mediates a current that decays much more slowly than the wild-type Ca V 2.3 during a 200-ms test pulse (Supplementary Figure 8a–8b), suggesting that OSI is remarkably decreased. To quantify the OSI of the Ca V 2.3 mutants, we employed the R200 value as an indicator, which is calculated by the mean current density at the end of the 200-ms test pulse divided by the peak amplitude ( Figure 5d and Supplementary Figure 8c). Specifically, the mean value of R200 increased from 0.17±0.01 in the wild-type Ca V 2.3 to 0.31±0.03 in Ca V 2.3 NQ under the test pulse holding at 10 mV ( Figure 5d and Supplementary Figure 8c). Moreover, R590E and R590Q also exhibited remarkably suppressed OSI, with increased R200 values of 0.35±0.04 and 0.27±0.03, respectively (10-mV test pulse) ( Figure 5d and Supplementary Figure 8c), suggesting that the R590-S6 IINCD interaction plays an essential role in the development of OSI in Ca V 2 channels. In contrast, mutants on the AID side show complicated effects on OSI kinetics ( Figure 5d and Supplementary Figure 8c). In particular, R378Q displayed an enhanced OSI, with a decreased R200 value of 0.11±0.02 (10-mV test pulse). R371E, which is located in the adjacent region of R378, exhibited an enhanced OSI as well, displaying an R200 value of 0.09±0.01 (10-mV test pulses). Nevertheless, OSI of the R371Q and R378E mutants do not show significant difference with that of the WT Ca V 2.3 channel ( Figure 5d and Supplementary Figure 8c). Moreover, the time course of channel inactivation could be well fitted by a single exponential. Conclusions drawn using time constants are nearly identical to those using R200 values (Supplementary Figure 8d), further supporting that S6 IINCD , R371 (AID), R378 (AID), and R590 (S4-S5 II ) play important roles in OSI modulation. However, the interaction between the AID and S6 IINCD could go beyond the current structures of Ca V channels and is worth investigation in future studies.
Methods
Expression and protein purification of the human Ca V 2.3 complex
Full-length Ca V 2.3 α1E (CACNA1E, UniProt ID: Q15878 isoform 1, or Ca V 2.3d), α2δ1 (CACNA2D1, UniProt ID: P54289), and β1 (CACB1, UniProt ID: Q02641) were amplified from a human cDNA library and subcloned into pEG BacMam vectors. To detect the expression and assembly levels of the Ca V 2.3 complex, a superfolder GFP, an mCherry, and an mKalama tag were fused to the C-terminal, N-terminal, and N-terminal regions of the Ca V 2.3 α1, α2δ1, and β1 subunits, respectively. Twin-Strep tags were tandemly inserted into the C-terminal and N-terminal of the α1 and α2δ1 subunits, respectively. The Bac-to-Bac baculovirus system (Invitrogen, USA) was used to conduct protein expression in HEK 293-F cells (Gibco, USA) following the manufacturer’s protocol. The bacmids were prepared using DH10Bac competent cells, and P1 viruses were generated from Sf 9 cells (Invitrogen, USA) after bacmid transfection. P2 viruses (1%, v/v) were used to infect HEK 293-F cells supplemented with 1% (v/v) fetal bovine serum. The cells were cultured at 37°C and 5% CO 2 for 12 hours before the addition of 10 mM sodium butyrate to the medium. The cells were cultured at 30°C and 5% CO 2 for another 48 h before harvest. No authentication was performed for the HEK 293-F or the Sf 9 cell line. No Mycoplasma contamination was observed. The cell pellets were resuspended at 4°C using Buffer W containing 20 mM HEPES pH 7.5, 150 mM NaCl, 5 mM β-mercaptoethanol (β-ME), 2 μg/mL aprotinin, 1.4 μg/mL leupeptin, and 0.5 μg/mL pepstatin A by a Dounce homogenizer, followed by centrifugation at 36,900 rpm for 1 h to collect the membrane. The membrane was resuspended again using Buffer W and solubilized by the addition of 1% (w/v) n-dodecyl-β-D-maltoside (DDM) (Anatrace, USA), 0.15% (w/v) cholesteryl hemisuccinate (CHS) (Anatrace, USA), 2 mM adenosine triphosphate (ATP) and 5 mM MgCl 2 on a rotating mixer at 4°C for 2 h. Addition of ATP and MgCl 2 is to remove associated heat shock proteins. The insoluble debris of cells was removed by another centrifugation at 36,900 rpm for 1 h. The supernatant was passed through a 0.22 μm filter (Millipore, USA) before being loaded into a 6 mL Strep-Tactin Superflow high capacity resin (IBA Lifesciences, USA). The resin was washed using 6 column volumes of Buffer W1 (20 mM HEPES pH 7.5, 150 mM NaCl, 5 mM β-ME, 0.03% (w/v) glycol-diosgenin (GDN) (Anatrace, USA), 2 mM ATP, and 5 mM MgCl 2 ). The purified Ca V 2.3 complex was eluted using 15 mL elution buffer (20 mM HEPES pH 7.5, 150 mM NaCl, 5 mM β-ME, 0.03% (w/v) GDN, and 5 mM d-Desthiobiotin (Sigma-Aldrich, USA)) and concentrated to 1 mL using a 100 kDa MWCO Amicon (Millipore, USA). The concentrated protein sample was subjected to further size-exclusion chromatography (SEC) by a Superose 6 Increase 10/300 GL gel filtration column (GE Healthcare, USA) using a flow rate of 0.3 mL/min and a running buffer containing 20 mM HEPES pH 7.5, 1.5 mM NaCl, 5 mM β-ME, and 0.01% (w/v) GDN (Anatrace, USA). The monodispersed peak fraction within 12–13.5 mL was pooled and concentrated to 5 mg/mL before preparing the cryo-EM grids.
Cryo-EM sample preparation and data collection
Holey carbon grids (Au R1.2/1.3 300 mesh) (Quantifoil Micro Tools, Germany) were glow-discharged using H 2 and O 2 for 60 s before being loaded with 2.5 μL purified Ca V 2.3 complex. The grids were automatically blotted for 4 s at 4°C and 100% humidity and flash-frozen in liquid ethane using a Vitrobot Mark IV (Thermo Fisher Scientific, USA). Cryo-EM data were collected using a 300 kV Titan Krios G2 (Thermo Fisher Scientific, USA) equipped with a K2 Summit direct electron detector (Gatan, USA) and a GIF Quantum LS energy filter (Gatan, USA). The dose rate was set to ∼9.2 e − /(pixel*s), and the energy filter slit width was set to 20 eV. A total exposure time of 6.72 s was dose-fractioned into 32 frames. A nominal magnification of ×130,000 was used, resulting in a calibrated super-resolution pixel size of 0.52 Å on images. SerialEM 48 was used to automatically acquire the movie stacks. The nominal defocus range was set from –1.2 μm to –2.2 μm.
Cryo-EM data processing
Motion correction was performed on 2,096 movie stacks using MotionCor2 49 with 5 × 5 patches, generating dose-weighted micrographs. The parameters of the contrast transfer function (CTF) were estimated using Gctf 50 . Particles were initially picked using the blob picker in cryoSPARC 51 , followed by 2D classifications to produce 2D templates and ab initio reconstruction to generate an initial reference map. Another round of particle picking was conducted using Template Picker in cryoSPARC, generating a dataset of 787,518 particles that were used for further processing. All data processing steps were performed in RELION-3.1 52 unless otherwise specified. A round of multi-reference 3D classification was conducted against one good and 4 biased references, generating 5 classes. Class 5 (75.2%), which was calculated using the good reference, displayed a classical shape of Ca V complexes featuring a transmembrane subunit and two soluble subunits residing on both sides of the micelle. Particles from class 5 were re-extracted and subjected to another round of 3D classification, resulting in 6 classes. Classes 1, 4, and 5 (43.5%) displayed well-resolved structural features, including continuous transmembrane helices of the α1E subunit and secondary structures within the α2δ1 and β1 subunits. To improve the map quality, Bayesian polish and CTF refinement were then conducted. The following 3D auto refinement generated a 3.1-Å map. The particle dataset was then imported back to cryoSPARC, where the final map was generated by Non-uniform (NU) refinement, which was reported at 3.1 Å according to the golden-standard Fourier shell correlation (GSFSC) criterion.
Model building
The cryo-EM map of Ca V 2.3 was reported at near-atomic resolution, which enabled us to reliably build and refine the model. The structure of the Ca V 2.2-α2δ1-β1 complex (PDB ID: 7VFS) 28 was selected as the starting model because of the high sequence identity and was docked into the map of Ca V 2.3 complexes using UCSF Chimera 53 . Side chains of the a1 subunit were manually mutated according to the sequence alignment between Ca V 2.3 and Ca V 2.2 and adjusted according to the EM density using Coot 54 . Side chains of α2δ1 were also manually adjusted according to the EM density. β1 was initially fit into the EM maps as a rigid body and manually refined against a low-resolution map of the Ca V 2.3 complex in Coot due to local structural heterogeneity. The manually adjusted models were then automatically refined against the cryo-EM maps using the integrated Real Space Refinement program within the PHENIX software package 55 . Model stereochemistry was also evaluated using the Comprehensive validation (cryo-EM) tool in PHENIX. All the figures were prepared using Open-Source PyMOL (Schrödinger, USA), UCSF Chimera 53 , or UCSF ChimeraX 56 .
Whole-cell voltage-clamp recordings of Ca V 2.3 channels in HEK 293-T cells
HEK 293-T cells were cultured with Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, USA) supplemented with 15% (v/v) fetal bovine serum (FBS) (PAN-Biotech, Germany) at 37°C with 5% CO 2 . The cells were grown in culture dishes (d = 3.5 cm) (Thermo Fisher Scientific, USA) for 24 h and then transiently transfected with 2 μg of control or mutant plasmid expressing the human R-type Ca V 2.3 calcium channel complex (Ca V 2.3 α1E, β1, α2δ1) using 1.2 μg of Lipofectamine 2000 Reagent (Thermo Fisher Scientific, USA). Patch-clamp experiments were performed 12 to 24 hours post-transfection at room temperature (21∼25°C) as described previously. Briefly, cells were placed on a glass chamber containing 105 mM NaCl, 10 mM BaCl 2 , 10 mM HEPES, 10 mM D-glucose, 30 mM TEA-Cl, 1 mM MgCl 2 , and 5 mM CsCl (pH = 7.3 with NaOH and an osmolarity of ∼310 mosmol -1 ). Whole-cell voltage-clamp recordings were made from isolated, GFP-positive cells using 1.5∼2.5 MΩfire-polished pipettes (Sutter Instrument, USA) filled with standard internal solution containing 135 mM K-gluconate, 10 mM HEPES, 5 mM EGTA, 2 mM MgCl 2 , 5 mM NaCl, and 4 mM Mg-ATP (pH = 7.2 with CsOH and an osmolarity of ∼295 mosmol -1 ). Whole-cell currents were recorded using an EPC-10 amplifier (HEKA Electronik, Germany) at a 20 kHz sample rate and were low-pass filtered at 5 kHz. The series resistance was 2∼4.5 MΩ and was compensated 80∼90%. The data were acquired by the PatchMaster program (HEKA Electronik, Germany). To obtain activation curves of Ca V 2.3 channels, cells were held at –100 mV, and then a series of 200-ms voltage steps from –60 mV to +50 mV in 5-mV increments were applied. The steady-state inactivation properties of Ca V 2.3 channels were assessed with 10-s holding voltages ranging from –100 mV to –15 mV (5-mV increments) followed by a 135-ms test pulse at +10 mV. To assess the time-dependent recovery from CSI, cells were depolarized to –40 mV (pre-pulse) for 1500 ms to allow Ca V 2.3 channels to enter CSI, and recovery hyperpolarization steps to –100 mV were applied for the indicated period (4 ms – 2,048 ms), followed by a 35-ms test pulse at +10 mV. To assess the cumulative inactivation of Ca V 2.3 channels in response to AP trains, the cells were held at –100 mV, and then the physiologically relevant AP trains was applied. The AP trains used to stimulate the HEK 293-T cells were recorded from a mouse hippocampal CA1 pyramidal neuron after current injection in the whole-cell current-clamp mode 57 . The spike pattern contained 13 action potentials in 2 s (mean frequency = 6.5 Hz). The percentage inactivation of Ca V 2.3 channels was calculated from the first spike eliciting maximal current to the other spikes in the AP trains. To analyze the extent of OSI, the ratio of remaining currents at 200 ms post-depolarization and the peak currents was calculated.
Electrophysiological data analysis
All data are reported as the mean ± SEM. Data analyses were performed using Origin 2019b (OriginLab, USA) and Prism 9 (GraphPad, USA). Steady-state activation curves were generated using a Boltzmann equation. where G is the conductance, calculated by G = I/(V-V rev ), where I is the current at the test potential and V rev is the reversal potential; G max is the maximal conductance of the Ca V 2.3 channel during the test pulse; V is the test potential; V 0.5 is the half-maximal activation potential; and k is the slope factor. Steady-state activation and inactivation curves were generated using a Boltzmann equation. where I is the current at the indicated test pulse; I max is the maximal current of Ca V 2.3 activation during the test pulse; V is the test potential; V 0.5 is the half-maximal inactivation potential; and k is the slope factor. Recovery curves from CSI were calculated from the results of 7 to 9 independent experiments where a series of recovery traces from inactivation time points were acquired. The data were fit using a single exponential of the following equation. where I is the current at the indicated intervals; I max is the current at 2048 ms; y 0 is the remaining current at –40 mV for 1500 ms; t is the indicated hyperpolarization time; and τ is the time constant of recovery from CSI. Statistical significance ( p < 0.05) was determined using unpaired Student’s t tests or one-way ANOVA with Tukey’s post hoc test.
Supporting information
Author contributions
Y.Z. conceived the project and supervised the research. Y.G. and Y.Q. carried out the molecular cloning experiments. Y.G. expressed and purified protein samples. Y.D. prepared samples for cryo-EM study. Y.G. and B.Z. carried out cryo-EM data collection. Y.G. processed the cryo-EM data. Y.G. built and refined the atomic model. Y.Z., Y.G., X.C.Z., and Y.W. analyzed the structure. Y.Z., Z.H., Y.G., S.X., and J.S. designed the electrophysiological experiments. S.X., X.C., H.X., C.P., and S.L. conducted the whole-cell voltage patch-clamp analysis. Y.G. wrote the original draft of the manuscript and prepared the figures. Y.Z., Y.G. and X.C.Z. edited the manuscript with input from all authors.
Data availability
The cryo-EM density map of the Ca V 2.3-α2δ1-β1 complex has been deposited in the Electron Microscopy Data Bank (EMDB) under the accession code EMD-33285. The coordinate for the Ca V 2.3 complex has been deposited in the Protein Data Bank (PDB) under the PDB ID 7XLQ.
Conflict of interest
All authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Competing Interest Statement
Competing Interest Statement
The authors have declared no competing interest.
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