Regulating quantal size of neurotransmitter release through a GPCR voltage sensor

Significance

The amount of neurotransmitter release triggered by Ca²⁺ from a presynaptic single vesicle (quantal size, QS) is fundamental and determines the strength of synaptic transmission. The present study provides compelling evidence that the QS of catecholamine is increased by membrane depolarization per se (bypassing Ca²⁺) via the voltage-sensitive ATP-GPCR/P2Y₁₂ in sympathetic chromaffin cells. D76-P2Y₁₂ and D127-P2Y₁₂ are revealed as the voltage-sensing sites by introducing two GPCR reporting systems. We also establish the physiology relevance of voltage dependence of catecholamine QS, offering a signaling pathway—Vm → P2Y₁₂(D76/D127) → Giβγ → QS → myocyte contractility—for diverse functions in the nervous system and other systems containing GPCRs.

Abstract

Current models emphasize that membrane voltage (Vm) depolarization-induced Ca²⁺ influx triggers the fusion of vesicles to the plasma membrane. In sympathetic adrenal chromaffin cells, activation of a variety of G protein coupled receptors (GPCRs) can inhibit quantal size (QS) through the direct interaction of G protein Giβγ subunits with exocytosis fusion proteins. Here we report that, independently from Ca²⁺, Vm (action potential) per se regulates the amount of catecholamine released from each vesicle, the QS. The Vm regulation of QS was through ATP-activated GPCR-P2Y₁₂ receptors. D76 and D127 in P2Y₁₂ were the voltage-sensing sites. Finally, we revealed the relevance of the Vm dependence of QS for tuning autoinhibition and target cell functions. Together, membrane voltage per se increases the quantal size of dense-core vesicle release of catecholamine via Vm → P2Y₁₂(D76/D127) → Giβγ → QS → myocyte contractility, offering a universal Vm-GPCR signaling pathway for its functions in the nervous system and other systems containing GPCRs.

According to the classical Ca²⁺ hypothesis of presynaptic transmission and neuroendocrine secretion, a presynaptic action potential activates Ca²⁺ influx which triggers Ca²⁺-dependent quantal (all or none) vesicular release of neurotransmitter/hormone (1). In 1996, we reported the example of Ca²⁺-dependent subquantal release, in which only part of the vesicular content of a native transmitter (catecholamine) is released during a transient fusion event (fusion pore flickers, or kiss and run), leading to a smaller quantal size (QS) in sympathetic adrenal chromaffin cells (ACCs) (2). Subquantal kiss-and-run release was subsequently confirmed in ACCs (3, 4), other endocrine cells (5, 6), glia (7, 8), and neurons (9–11). In 2005, the regulator of subquantal kiss-and-run release was found: G protein-coupled receptor (GPCR)-dependent Giβγ reduces QS (7). In addition, endogenous dynamin-1 limits expansion of the vesicle fusion pore and maintains all Ca²⁺-induced exocytotic release via the kiss-and-run/subquantal release mode under physiological conditions in ACCs (12). More recently, the mechanism underlying subquantal catecholamine release has been revealed to consist of the joint kiss-and-run fusion pore and matrix binding (13).

The molecular machine that gates the fusion pore is known to involve the SNARE complex and other regulators (14, 15), including synaptotagmin (16), dynamin (12, 17–19), microdomain Ca²⁺ (7, 11, 20), and GPCR-dependent Giβγ (12, 19, 21–23). Giβγ is downstream of Gi-GPCR activation because only Gi is sufficiently abundant to allow Giβγ to function (24). There are many Gi-GPCRs, including those activated by the native transmitters acetylcholine (25, 26), somatostatin, and ATP (or ADP). ATP inhibition of QS (termed AIQS here) is induced via the GPCR-Gi-βγ pathway in ACCs (19). Its physiological relevance lies in the fact that ATP release (13) evoked from a neighboring cell directly leads to AIQS (19). AIQS is downstream of the ligand-GPCR-Gi-βγ signaling pathway, which is initiated by a native ligand that regulates the GPCR.

ATP is the ligand of two families of purinergic receptors, P2Xs and P2Ys. P2Xs are ion channels and P2Ys are GPCRs, including Gq (P2Y_{1, 2, 4, 6, 11}) and Gi types (P2Y_{12, 13, 14}) (27, 28). Among the P2Ys, P2Y₁ exists in most tissues, including epithelial and endothelial cells, platelets, and immune cells (27, 29), and P2Y₁₂ is strongly expressed on platelets, where it plays fundamental roles in their activation and aggregation (28, 30, 31), as well as in microglia (32, 33), smooth muscle cells (34), and chromaffin cells (35, 36). P2Y₁₂ is a major target for drugs against cardiovascular diseases (thrombosis, stroke, and myocardial infarction) and currently, the top-selling drugs targeting P2Y₁₂ (clopidogrel and cangrelor) are used for antithrombotic therapy (37–39). However, the subtype(s) of P2Ys that mediate AIQS remains unknown in ACCs.

To estimate the effects of GPCR-Giβγ on single-vesicle fusion, we used highly sensitive electrochemical microcarbon fiber electrodes (CFEs, 7-µm diameter) to measure the vesicle contents released from single fusion pores in ACCs (2, 12, 13, 19). For real-time imaging of vesicle fusion events, total internal reflection fluorescence (TIRF) microscope imaging in neuropeptide Y (NPY)-pHluorin transfected ACCs allows the distinction of kiss-and-run from full-fusion-like single-vesicle fusion modes (12, 13, 40). To determine the sites responsible for specific functions in a GPCR, reconstitution of the GPCR in a reporting system and its functional assay are needed (38, 41–43).

In the present study, by using CFEs to measure QS and/or the fusion mode of single-vesicle release events, TIRF live imaging to confirm single-vesicle fusion modes, and two complementary GPCR reporting systems (a high-throughput assay of Gi-α-IP3-[Ca²⁺]_i for screening sites and a precision assay of Gi-βγ-GIRK current for validation) to determine the voltage-sensing sites in the P2Y-GPCR, we discovered that P2Y₁₂ mediates AIQS, which is disinhibited by depolarization (membrane voltage [Vm]) via two voltage-sensing sites (D76 and D127) in native neuroendocrine chromaffin cells.

Results

Depolarization Per Se Relieves the Intrinsic AIQS.

We have previously demonstrated that subquantal catecholamine release occurs in ACCs (2), and the QS is regulated by GPCR-Gi-βγ following elevation of the intracellular Ca²⁺ concentration [Ca²⁺]_i by caffeine (20 mM) (19). When caffeine was applied for 10 s, a burst of amperometric spikes—representing quantal catecholamine release from single large dense-core vesicles—was evoked in ACCs (Fig. 1A). Indeed, ATP inhibited the QS (Fig. 1A, see also refs. 13, 19), or the AIQS phenotype was produced by the smaller QS (Fig. 1A, see also ref. 19). Surprisingly, when a cell was depolarized by high KCl (70 mM), the AIQS phenotype was abolished (Fig. 1A). The effect of Vm on GPCR-Gi-βγ-based AIQS was fully reversible (Fig. 1A) and independent of [Ca²⁺]_i (Fig. 1B). The statistics of this Vm-dependent AIQS are shown in Fig. 1 D and E and SI Appendix, Fig. S1. ACCs were bathed in 0 Ca²⁺ solution containing 1 mM EGTA during caffeine stimulation to ensure no Ca²⁺ influx during depolarization. [Ca²⁺]_i measurements using Fura-2 showed that the caffeine-induced [Ca²⁺]_i elevation was independent of depolarization (Fig. 1 B and F). The depolarization from −70 mV to −20 mV by 70 mM KCl (but not caffeine or ATP) was confirmed by current-clamp recordings (Fig. 1 C and F). Note that ATP was required for the Vm-dependent AIQS phenotype and in the absence of ATP, depolarization had no effect on the QS (SI Appendix, Figs. S1 and S14). Furthermore, the QS did not change with repetitive caffeine stimulation (SI Appendix, Fig. S2).

Fig. 1.

ATP inhibition of quantal size (AIQS) of vesicular catecholamine release is attenuated by membrane depolarization voltage (Vm). (A) Basic characteristics of voltage-dependent AIQS. (Upper) cartoon of Vm-AIQS with steps 1 through 3. (Lower) ATP (100 µM) inhibits the QS of amperometric spikes induced by caffeine (Caf; 20 mM for 10 s) and this is reversed by 70 mM KCl (70K)-induced depolarization (dashed boxes show averaged quantal events). Each cell was exposed four times to caffeine-containing solutions. The reversible Vm-AIQS contains four steps (1, 2, 3, and 2′): (step 1) Ca²⁺ is released from the ER store by caffeine and triggers vesicular catecholamine release events recorded as spikes in amperometric current (I_amp) recordings; (step 2) ATP activates P2Y receptors and reduces QS; (step 3) membrane depolarization (by 70K) removes the inhibitory effect of ATP; and finally (step 2′) AIQS recovers after removing depolarization. (B) Intracellular Ca²⁺ [Ca²⁺]_i (F₃₄₀/F₃₈₀) elevation measured by Fura2/AM, corresponding to A. (C) Membrane potential recorded by current clamp, corresponding to A. Caffeine with or without ATP does not change the membrane potential, while 70 mM KCl depolarizes it by ∼50 mV. (D) Cumulative distribution of QS corresponding to the four steps (1, 2, 3, and 2′) of Fig. 1A (198 spikes for group 1, 157 spikes for group 2, 193 spikes for group 3, and 166 spikes for group 2′. The four groups are matched measurements from n = 15 cells. For groups 2 vs. 3, ***P < 0.001, K-S test). (E) Quantitative analysis of single-vesicle release events (quanta) from cells of Fig. 1D. Analyzed spikes meet the 5-SD noise threshold criterion. ATP significantly reduces QS, while depolarization by 70 mM KCl completely removes the inhibitory effect of ATP. When the 70 mM KCl is washed out, QS is inhibited by ATP again (the four groups are matched measures from n = 15 cells, paired Friedman test, post hoc Dunn’s multiple comparisons test). (F, Left) Depolarization does not affect the caffeine-induced [Ca²⁺]_i elevation in Ca²⁺-free bath (n = 14 cells, Friedman test, post hoc Dunn’s multiple comparisons test). (F, Right) KCl (70 mM) depolarizes Vm by ∼50 mV (from –70 mV to –20 mV, n = 5 cells, paired Student’s t test). (G) Confirmation of voltage-dependent AIQS by another orthogonal assay of TIRFM imaging. Statistics of TIRF imaging data (SI Appendix, Fig. S4) confirm that the probability ratio [FFL ratio = N(FFL)/N(FFL + KAR), where N(FFL) is the number of full-fusion-like (FFL) events and N(FFL + KAR) is the total number of FFL and kiss-and-run (KAR) events]. Similar to the QS index from CFE recordings in E (multiple treatments were applied to the same cells), the corresponding index (FFL ratio) from TIRFM imaging is also increased by depolarization (70K) (n = 9 cells, Friedman test, post hoc Dunn’s multiple comparisons test, further details in SI Appendix, Fig. S4 and Materials and Methods). A–C show different cells with the same time scale. Data are presented as the mean ± SEM. (E–G) *P < 0.05, **P < 0.01, ***P < 0.001; NS, not significant.

To ensure that the relief of the inhibitory effect of ATP is due to membrane depolarization per se, we repeated the experiments shown in Fig. 1, but replacing 70 mM KCl depolarization by whole-cell voltage clamp. Similar to 70 mM KCl depolarization (Fig. 1), whole-cell depolarization from −70 mV to 0 mV abolished the AIQS (SI Appendix, Fig. S3). Thus, the AIQS phenotype is produced by membrane depolarization per se.

In addition to CFE recordings, single-vesicle exocytosis can be imaged live by TIRF, which measures the dynamics of single large dense-core vesicle release of pHluorin-tagged NPY in neurons (40, 44) and ACCs (13, 45). The Ca²⁺-dependent full-fusion-like mode, corresponding to full quantal release, and the kiss-and-run mode, corresponding to subquantal release (13) were identified by TIRF imaging (SI Appendix, Fig. S4 A and B and Movies S1–S4, see also refs. 13, 40, 45). Consistent with the CFE recordings, TIRF imaging revealed that ATP shifted the fusion mode from full-fusion-like to kiss-and-run (full-fusion-like ratio changed from 68 to 33%), while this AIQS-like effect was largely abolished by depolarization (full-fusion-like ratio changed from 33 to 62%, Fig. 1G and SI Appendix, Fig. S4).

P2Y₁₂ Is the ATP Receptor for Voltage-Dependent AIQS.

Since AIQS was present in ACCs (Fig. 1, see also ref. 19), we aimed to identify the associated receptor. Because AIQS is probably produced by an unknown subtype of P2Y receptor coupled to Gi in ACCs (19), we first identified candidates in pharmacological experiments. When ACCs were treated with ATP (100 μM), 70 mM KCl depolarization disinhibited AIQS and increased QS (Fig. 2A). In contrast, suramin (100 μM), a broad-spectrum antagonist of P2 purinoceptors (46, 47), fully removed the effect of depolarization on AIQS (Fig. 2B). However, PPADS (50 μM), a broad-spectrum antagonist of P2X purinoceptors (48, 49), did not block the depolarization effect (Fig. 2C). Thus, depolarization relieves AIQS via Gi-P2Y(s). Strikingly, ARC66096 (10 μM, a specific antagonist of P2Y₁₂) (50, 51) blocked the effect of depolarization on AIQS (Fig. 2D). These pharmacological experiments suggested that P2Y₁₂ is a candidate for the depolarization-induced relief of AIQS.

Fig. 2.

P2Y₁₂ is the ATP receptor for voltage-dependent AIQS—pharmacological evidence. (A) AIQS is regulated by voltage. Statistically, normalized QS (QS normalized by averaged QS in control bath, black) is significantly increased by 70 mM KCl (70K) depolarization (red) in ACCs treated with 100 µM ATP (n = 17 cells, **P < 0.01, paired Student’s t test). Inset, averaged quantal spikes without (black solid line) and with (red dashed line) 70 mM KCl depolarization. (B) Suramin (100 µM, a broad-spectrum P2XR and P2YR blocker) removes the effect of 70 mM KCl depolarization on AIQS (n = 13 cells, P = 0.49, paired Student’s t test). (C) PPADS (50 µM, a P2XR-specific blocker) does not remove the effect of 70 mM KCl depolarization on AIQS (n = 8 cells, *P < 0.05, paired Student’s t test). (D) ARC66096 (10 µM, a P2Y₁₂ blocker) removes the 70 mM KCl-depolarization effect on AIQS (n = 19 cells, P = 0.74, paired Student’s t test). Data are presented as the mean ± SEM. *P < 0.05, **P < 0.01; NS, not significant.

To confirm the pharmacological findings, we first performed reverse transcriptional PCR (RT-PCR) of rat adrenal medulla and found four Gq-coupled receptors (P2Y₁, P2Y₂, P2Y₄, and P2Y₆) and three Gi-coupled receptors (P2Y₁₂, P2Y₁₃, and P2Y₁₄) (SI Appendix, Fig. S5A and Table S2). Second, following the pharmacological results (Fig. 2), we designed and validated two knockdown (KD) shRNAs targeting P2Y₁₂ in rat ACCs (SI Appendix, Fig. S5 C–F) and found that P2Y₁₂ was the major receptor mediating AIQS. Western blot analysis of HEK293A cells and immunofluorescent staining of native chromaffin cells revealed that P2Y₁₂ was knocked down (∼80% reduction) by the shRNAs and could be fully rescued (Fig. 3 A and B). Depolarization disinhibited the AIQS in control scrambled (Fig. 3C and SI Appendix, Fig. S6A), but not P2Y₁₂-KD (shRNA 1 or 2) ACCs (Fig. 3 D and E and SI Appendix, Fig. S6 B and C), while overexpressing shRNA-resistant P2Y₁₂ fully recovered the Vm dependence of AIQS (Fig. 3F and SI Appendix, Fig. S6D). Thus, P2Y₁₂ is essential for voltage-dependent AIQS in rat ACCs. Finally, using P2Y₁₂-knockout (KO) mice (SI Appendix, Fig. S5B), the depolarization-sensitive AIQS was present only in wild-type (WT) but not P2Y₁₂-KO cells (Fig. 3G). Taken together, P2Y₁₂ is not only the ATP receptor for AIQS, but is also responsible for its Vm dependence.

Fig. 3.

P2Y₁₂ is the ATP receptor for voltage-dependent AIQS—genetic evidence. (A) Western blots and statistics of P2Y₁₂ expression in HEK293A cells transfected with scrambled, P2Y₁₂-KD shRNA1, P2Y₁₂-KD shRNA2, and P2Y₁₂-rescue plasmids (five independent experiments, one-way ANOVA, post hoc Tukey’s multiple comparisons test). (B) Immunostaining and statistics of P2Y₁₂ expression (scrambled, 25 cells; shRNA1, 26 cells; shRNA2, 10 cells; rescue, 17 cells, one-way ANOVA, post hoc Tukey’s multiple comparisons test). (Scale bar, 5 μm.) (C, Left) Typical CFE I_amp traces of ATP (100 µM)-induced AIQS from scrambled cells with or without 70 mM KCl (70K) depolarization. (C, Right) Statistics of the effect of depolarization on QS in 26 scrambled cells (**P < 0.01, Wilcoxon test). Inset, averaged quantal spikes without (black solid line) and with (red dashed line) 70 mM KCl depolarization (Vm). (D and E) Statistics of averaged quantal events and AIQS in 21 P2Y₁₂-KD-shRNA1 cells (D, P = 0.64, Wilcoxon-test) and 19 P2Y₁₂-KD shRNA2 cells (E, P = 0.59, Wilcoxon test) with or without 70 mM KCl depolarization (Vm). (F) Statistics of AIQS in 30 P2Y₁₂-rescued cells (**P < 0.01, Wilcoxon test) with or without 70 mM KCl depolarization (Vm). (G, Upper Left) 70 mM KCl depolarization-induced AIQS is intact in a WT cell. (G, Upper Right) Cumulative distribution curve of QS (n = 14 cells, four groups [1, 2, 3, and 2′] of quantal events [167, 141, 140, and 125]). For groups 2 vs. 3, ***P < 0.001, K-S test and statistics of Vm-AIQS in 14 WT cells (Friedman test, post hoc Dunn’s multiple comparisons test). (G, Lower Left) 70 mM KCl depolarization-induced AIQS is eliminated in a P2Y₁₂-KO cell. (G, Lower Right) Cumulative distribution curve of QS (n = 12 cells; groups 1, 2, 3, and 2′ of 136, 120, 118, and 101 events). For groups 2 vs. 3, NS, not significant, K-S test and statistics of Vm-AIQS in 12 P2Y₁₂-KO cells (Friedman test, post hoc Dunn’s multiple comparisons test). Data are presented as the mean ± SEM. (C–G) **P < 0.01, ***P < 0.001; NS, not significant.

Depolarization Regulates P2Y₁₂-Gi Signals via the Voltage-Sensing Sites D76 and D127 in Reconstituted Systems.

Based on the finding that P2Y₁₂ mediated the Vm dependence of AIQS in ACCs (Figs. 1–3 and SI Appendix, Figs. S1–S6), we proposed that P2Y₁₂ activity is voltage dependent and P2Y₁₂ contains voltage-sensing sites. To test this hypothesis at the molecular level, we designed a P2Y₁₂ reporting system via the P2Y₁₂-Gi-α-IP3-[Ca²⁺]_i signaling pathway (Fig. 4A), which is a high-throughput assay for multiple cells per experiment with [Ca²⁺]_i imaging (Fig. 4B) to assess the voltage-dependence of P2Y₁₂ in reporting cells (Fig. 4C). The Ca²⁺ indicator GCaMP3, P2Y₁₂, and the Gαi3q chimera were co-overexpressed in HeLa cells and the Gαi3q chimera results in coupling with Gαi-coupled receptors but signaling through the Gαq-mediated PLC-IP3-[Ca²⁺]_i mobilization pathway (42, 52, 53) (Fig. 4 A and B and SI Appendix, Fig. S7 A, B, and E and see also Materials and Methods). Membrane depolarization by 70 mM KCl (SI Appendix, Fig. S7H) inhibited the increased [Ca²⁺]_i induced by 2MesADP (P2Y₁₂ agonist) (27) in the P2Y₁₂-reconstituted reporting cells (Fig. 4 C and E and Movies S5–S7), confirming that Vm modulates P2Y₁₂ function: depolarization decreases P2Y₁₂ activation as measured by the Gαi pathway. Using this high-throughput imaging assay, we screened for voltage-sensing sites in P2Y₁₂ and found two amino acid sites: D76 and D127 (Fig. 4 C–E and SI Appendix, Fig. S10A) (54). Either D76N or D127N mutation abolished the Vm dependence of P2Y₁₂ (Fig. 4E and SI Appendix, Fig. S8), indicating that these are the voltage-sensing sites of P2Y₁₂.

Fig. 4.

Dissecting the voltage-sensing sites of P2Y₁₂ by two reconstituted GPCR-signaling assays. (A) Cartoon of the P2Y₁₂-Gi-α-IP3-[Ca²⁺]_i assay. HeLa cells were transfected with the plasmids P2Y₁₂, GCaMP3 (cytosol), and Gαi3q. The P2Y₁₂-specific agonist 2MesADP (10 µM) activates the P2Y₁₂-Gαi3q-PLC-IP3 pathway to induce Ca²⁺ release from the ER (SI Appendix, Fig. S7). (B) Confocal image of multiple HeLa cells transfected with P2Y₁₂, Gαi3q, and GCaMP3 plasmids. (C) [Ca²⁺]_i (ΔF/F₀) triggered by 2MesADP with or without depolarization. Overlapped gray [Ca²⁺]_i traces from many cells in the same dish; black/red traces show averaged [Ca²⁺]_i. Depolarization (70 mM KCl, 70K) reversibly decreases the averaged amplitude of ΔF/F₀ spikes. (D) P2Y₁₂ purinergic receptor model based on the crystal structure (PDB accession no. 4PXZ) (54). Arrows indicate the positions of the two candidate Vm-sensing residues D76 and D127. (E) Statistics of evoked [Ca²⁺]_i signals. Compared with the WT (C), Vm depolarization reduced [Ca²⁺]_i (ΔF/F₀) of P2Y₁₂-WT, but not D76N or D127N mutation. For WT, n = 108 cells; for the D76N mutation, n = 45 cells; for the D127N mutation, n = 76 cells (SI Appendix, Fig. S8). Friedman test, post hoc Dunn’s multiple comparisons test. (F) Cartoon of the P2Y₁₂-Gi-βγ-GIRK current assay (SI Appendix, Fig. S9). (G) Whole-cell recordings of GIRK current (I_GIRK) induced by 140 mM KCl (140 K) in HEK293A cells coexpressing GIRK1/4 and P2Y₁₂ (∆I₁₀ and ∆I₁₀₀ were I_GIRK induced by 10 and 100 µM ADP). Note that I_GIRK is larger at 100 μM than at 10 μM ADP (see also SI Appendix, Fig. S9E). (H) I_GIRK evoked by 10 µM or 100 µM ADP measured at Vm = −40 mV (Left) or −100 mV (Right). The I_GIRK ratio (∆I₁₀/∆I₁₀₀) was defined as γ(Vm) = ∆I₁₀/∆I₁₀₀. Statistics are shown in J (WT). (I) The γ(Vm)-Vm curve showing that γ(Vm) is voltage dependent. (J) Statistics of γ(Vm). For WT-P2Y₁₂, γ(Vm) is potentiated at −100 mV versus −40 mV. At −40 mV, γ(Vm) = 0.55; at −100 mV, γ(Vm) = 0.79, (n = 29 cells, ***P < 0.001, paired Student’s t test). With the D76N or D127N mutation, the Vm-dependence of γ(Vm) is abolished (n = 15 cells for D76N; n = 18 cells for D127N, see also SI Appendix, Fig. S10, paired Student’s t test). (K) Summary of the effects of P2Y₁₂ mutations on the ratio γ(Vm) (SI Appendix, Fig. S10). Data are presented as the median with interquartile range (E) or the mean ± SEM (J). *P < 0.05, **P < 0.01, ***P < 0.001; NS, not significant.

Next, to further analyze and quantify the voltage-sensitive sites of P2Y₁₂, we designed another reporting system for P2Y₁₂ activation based on the P2Y₁₂-Gi-βγ-GIRK signaling pathway. In this precision assay, we used patch clamp to evaluate the voltage dependence of P2Y₁₂ (Fig. 4 F–I and SI Appendix, Fig. S9, see also refs. 31, 41, 43). P2Y₁₂ and GIRK1/4 channels together were coexpressed in HEK293A cells. Following ADP stimulation, the activation of P2Y₁₂ was detected by the whole-cell GIRK current I_GIRK (Fig. 4 F and G and SI Appendix, Fig. S9). To assess the voltage dependence of P2Y₁₂, we calculated the ratio of I_GIRK at the paired ADP doses 10 μM (middle dose) and 100 μM (saturation dose): γ(Vm) = ∆I₁₀/∆I₁₀₀ = I_GIRK (10 μM)/I_GIRK (100 μM), where Vm is the holding potential (SI Appendix, Fig. S11B). Importantly, γ(Vm) was larger at −100 mV than at −40 mV, indicating stronger P2Y₁₂ activation and greater ADP-P2Y₁₂ binding affinity with hyperpolarization (Fig. 4 H–J). To determine whether the negatively charged D76 and D127 were critical for the Vm dependence, in addition to D76N and D127N (negative-to-neutral charge), we made D76E and D127E mutations (negative-to-negative charge). In contrast to native P2Y₁₂ (Fig. 4 H–J), the mutation D76N or D127N, but not D76E or D127E, abolished the voltage dependence of γ(Vm) (Fig. 4 J and K and SI Appendix, Fig. S10 B–F). These results from the precision assay confirmed that not only the sites but also the negative charges (either D or E) of both D76 and D127 were essential for the Vm dependence of P2Y₁₂. Taken together, D76 and D127 are the voltage-sensing sites of P2Y₁₂.

Validation of P2Y₁₂ Vm Sensors for AIQS in Chromaffin Cells.

Next, we examined the effects of the mutations D76N and D127N on the Vm dependence of AIQS (Vm-AIQS) in native ACC cells. First, comparing WT and KD cells (P2Y₁₂-shRNA-KD, Fig. 3 A–F), the rescue by shRNA1-resistant WT restored both AIQS and Vm-AIQS in ACCs (Fig. 5A). However, replacing the shRNA1-resistant WT by either shRNA1-resistant P2Y₁₂-D76N or -D127N, the Vm-AIQS effect (but not AIQS itself) was abolished (Fig. 5 B–D). Thus, both D76 and D127 are voltage-sensing sites of P2Y₁₂ responsible for the Vm-AIQS effect (Fig. 5E).

Fig. 5.

The P2Y₁₂-D76 and -D127 sites are voltage sensors in chromaffin cells. (A, Left) The depolarization dependence of AIQS is rescued by P2Y₁₂*, which is a functional P2Y₁₂ resistant to shRNA1. (A, Right) Statistics of QS (n = 11 cells, Friedman test, post hoc Dunn’s multiple comparisons test). Dashed boxes show averaged quantal events. (B and C, Left) The Vm dependence of AIQS cannot be rescued by either D76N-P2Y₁₂ (B) or D127N-P2Y₁₂ (C). (B and C, Right) Statistics of QS (B, n = 17 cells, one-way ANOVA, post hoc Tukey’s multiple comparisons test; C, n = 7 cells, Friedman test, post hoc Dunn’s multiple comparisons test). (D) Summary of the P2Y₁₂ Vm dependence of AIQS. (E) Model of voltage-sensitive P2Y₁₂ regulation of quantal release in native ACCs. ATP is released from chromaffin vesicles and activates autoinhibitory P2Y₁₂, which inhibits quantal catecholamine release. Meanwhile, depolarization deactivates P2Y₁₂ via the D76 and D127 sites, and thus relieves the inhibitory effect of ATP. Data are the mean ± SEM. (A–C) *P < 0.05, **P < 0.01; NS, not significant.

Physiological Relevance of the Vm Dependence of AIQS.

Next, we investigated whether the Vm dependence of AIQS exists in adrenal slice ACCs (Fig. 6A). CFEs were used to record the amperometric current (I_amp) representing catecholamine overflow from all nearby quantal release from slice ACCs bathed in Ca²⁺-free solution containing 10 mM EGTA after stimulation (Fig. 6 A and B). ATP reversibly inhibited the caffeine-induced I_amp (Fig. 6B). Importantly, corresponding to the Vm-AIQS in cultured ACCs (Fig. 1A), the Vm dependence of the ATP-inhibited I_amp induced by caffeine was preserved in adrenal slices, as 70 mM KCl depolarization disinhibited this I_amp (Fig. 6B). That this voltage-dependent phenomenon is independent of Ca²⁺ channels was demonstrated by using the Ca²⁺ channel blocker CdCl₂ (SI Appendix, Fig. S12)._. Similar to the Vm-dependent AIQS in cultured ACCs (Fig. 3G), the Vm dependence of ATP-inhibited I_amp in adrenal slices (Vm-AIQS in situ) was also abolished by P2Y₁₂-KO (Fig. 6C). Although depolarization in these slice experiments was by 70 mM KCl, the Vm-AIQS in situ likely remains with physiological stimulation because, when depolarization by 70 mM KCl (Figs. 1–3) was replaced by whole-cell depolarization with either action potentials (SI Appendix, Fig. S13) at physiological frequencies (55, 56) or Vm pulses (SI Appendix, Fig. S3), the Vm dependence of AIQS remained in cultured ACCs.

Fig. 6.

Potential relevance of voltage-dependent GPCR-P2Y₁₂ mediation of catecholamine release. (A) Image of a freshly prepared mouse adrenal slice 150 µm thick. (A, Inset) Enlargement showing a recording CFE (Φ 7 µm) in the slice with visible individual ACCs. (B, Left) In an adrenal slice bathed in 0 Ca²⁺ (containing 10 mM EGTA) and ATP (100 µM), the ATP-inhibited catecholamine overflow (I_amp, amperometric current) evoked by caffeine (20 mM for 40 s) is fully removed by 70 mM KCl (70K) depolarization in WT ACCs. (B, Right) Statistics of WT (n = 9 slices from four mice, Friedman test, post hoc Dunn’s multiple comparisons test). (C) As in B, except the Vm dependence of I_amp is abolished by P2Y₁₂-KO. (C, Right) Statistics of KO (n = 9 slices from four mice, one-way ANOVA, post hoc Tukey’s multiple comparisons test). (D, Upper) Cartoon of the protocol for the catecholamine fluid collection from adrenal slices treated with 0 Ca + ATP + caffeine (control fluid) or 0Ca + 70K + ATP + caffeine (70K fluid). The fluids were puffed onto myocytes through the patch pipette (10 to 12 µm in diameter) to test their effects. (D, Lower) Setup to record excitation–contraction coupling in a myocyte. (E) Diagram of how catecholamine fluid was used to potentiate myocyte contractility (see F for details). (F) Quantification of ΔL2/ΔL1, where ΔL1 and ΔL2 represent the contraction length of myocytes before and after application of depolarization fluid (0Ca + 70K + ATP + caffeine) versus control fluid (0Ca + ATP + caffeine) (n = 20 myocytes for control fluid, and n = 22 myocytes for depolarization fluid, unpaired Student’s t test). (G) Model of the Vm-AIQS signal pathway to a target cell. The Vm dependence of QS implies a broad physiological relevance by providing the entire Vm-P2Y₁₂ pathway in vivo. Data are presented as the mean ± SEM. (B, C, and F) *P < 0.05; NS, not significant.

To provide proof of concept for the physiological impact of the Vm-AIQS effect via catecholamine release from adrenal slices, we determined whether the caffeine-induced release could affect cardiac myocytes, the major peripheral targets of sympathetic ACCs (57, 58). We collected the fluid (30 µL/gland) from adrenal slices (two slices per gland) incubated in either control (0 mM Ca²⁺ + 100 µM ATP + 20 mM caffeine) or depolarization (control + 70 mM KCl) solution. Compared to the control solution, the depolarization solution should contain more catecholamine produced by the Vm-AIQS effect. These stock solutions were diluted 10 times into working solutions (2 mM Ca²⁺ and 2 mM caffeine) and applied to beating myocytes (Fig. 6D). In contrast to the control solution, the depolarization solution increased the contractility of myocytes (Fig. 6 D–F), implying potential regulation of cardiac excitation–contraction coupling by Vm-AIQS.

Finally, in addition to the ligand-GPCR pair of ATP-P2Y₁₂, somatostatin-GPCR is also known to inhibit QS in ACCs (13, 19). We found that, like ATP-P2Y₁₂, somatostatin-SSTR (somatostatin receptor) (59) had a similar phenotype of Vm dependence (Figs. 1–6 and SI Appendix, Fig. S15), indicating that the Vm dependence of GPCR-Gi signaling is likely a general phenomenon in native mammalian cells. In principle, the presence of two types of Gi-GPCRs (P2Y₁₂ and SSTR) allows the cell to be regulated by both neurotransmitters (ATP and somatostatin) arising from other vesicles or cells. On the other hand, a single depolarization can shut down the Vm-dependent autoinhibition of both Gi-GPCRs.

Discussion

Neurotransmitter release from presynaptic cells induced by action potentials is fundamental, and neurotransmission depends on QS. In the present work we found that, after Ca²⁺ as the coregulator to trigger vesicle fusion, depolarization—in addition to activating voltage-gated channels—per se acts as coregulator of GPCR to determine the QS. The P2Y₁₂-mediated Vm dependence of QS has broad physiological relevance, providing the entire pathway [Vm → P2Y₁₂ (D76/D127) → Giβγ → QS → secretion] the ability to modulate the hormone level in circulating plasma for targeted cells throughout the body (including cardiac myocytes, Fig. 6G).

The major finding of the present work was that Vm depolarization relieved AIQS (ATP inhibition of QS) via ATP-dependent GPCR (P2Y)-Giβγ in rodent ACCs. This was supported by the findings that in cultured ACCs: 1) depolarization per se relieved the AIQS, and increased the QS of caffeine-induced secretion by ∼200% (Fig. 1 A–F and SI Appendix, Figs. S1–S3), a characteristic termed Vm-AIQS. 2) TIRF imaging confirmed that depolarization shifted the vesicle fusion mode from kiss-and-run to full fusion and increased the proportion of full-fusion events from 33 to 62% (Fig. 1G and SI Appendix, Fig. S4 and Movies S1–S4), which confirmed Vm-AIQS. 3) Vm-AIQS also persisted in the more physiological adrenal slice (Fig. 6A–C). 4) Like ATP-P2Y, another native ligand-GPCR pair, somatostatin and its GPCR, possessed a Vm-AIQS-like phenotype with somatostatin replacing ATP (SI Appendix, Fig. S15), indicating that Vm-GPCR-QS signaling extends beyond GPCR-P2Y₁₂, and could be a modulatory mechanism shared by many GPCRs and cell types.

Regarding the molecular identity and mechanisms of Vm-AIQS, we found that P2Y₁₂ and its voltage-sensing sites are responsible because RT-PCR showed that P2Y₁₂ was present in rat and mouse ACCs (SI Appendix, Fig. S5 A and B); using pharmacological methods, Vm-AIQS was blocked by suramin (an antagonist of P2Ys and P2Xs) and ARC66096 (an antagonist of P2Y₁₂) (Fig. 2 B and D), but not by PPADS (an antagonist of P2Xs) (Fig. 2C); using genetic approaches, the phenotype was largely blocked either by P2Y₁₂-KO (Fig. 3G) or KD by two P2Y₁₂-shRNAs (Fig. 3 A–E), and was rescued by the corresponding shRNA-resistant P2Y₁₂ (Fig. 3F and SI Appendix, Fig. S6); using assays of P2Y₁₂ function in two complementary reconstitution systems (Fig. 4 and SI Appendix, Figs. S7 and S9), P2Y₁₂-D76 and -D127 were identified as the Vm-sensing sites, because mutagenesis of either D76N or D127N abolished the Vm dependence of P2Y₁₂ while their basic P2Y₁₂-Gi signals remained (Fig. 4 and SI Appendix, Figs. S8 and S10); and overexpressing either D76N or D127N in ACCs silenced by P2Y₁₂-shRNA#1 abolished the Vm dependence, while the control WT-P2Y₁₂ did not (Fig. 5). Taken together, D76 and D127 are the Vm-sensing sites in P2Y₁₂ responsible for Vm-AIQS.

Regarding physiological relevance, the Vm-AIQS occurred in the quantal release evoked by increasing cytosolic Ca²⁺ through either the caffeine-sensitive endoplasmic reticulum (ER) store (Fig. 1A) or by whole-cell dialysis with high Ca²⁺ (SI Appendix, Fig. S13). In principle, the quantal release evoked by Ca²⁺ influx through ion channels (19) or other internal Ca²⁺ stores (60, 61) would also be regulated by the Vm-dependent GPCRs. The Vm-AIQS was present not only in cultured ACCs (Figs. 1–3, and 5 and SI Appendix, Figs. S1, S3, S4, S6, and S13), but also in fresh adrenal slices (Fig. 6 A–C). In WT but not P2Y₁₂-KO slices, AIQS was preserved as ATP-inhibited I_amp (the sum of release from all nearby cells), which is proportional to the total catecholamine release from the adrenal medulla into the blood. Thus, the Vm dependence of AI-QS (or AI-I_amp) could substantially regulate the catecholamine level in the circulation and thus all of its peripheral targets (57, 58), including myocyte contractility (Fig. 6 D–F). Excitation–contraction coupling was enhanced by the fluid from adrenal slices exposed to 70 mM KCl, which depolarizes ACCs, disinhibits P2Y₁₂, and increases the catecholamine level in solution. This provides proof of concept, that the effect of Vm-AIQS can impact the heart and other peripheral functions through regulating the blood catecholamine level.

The discovery of the Vm sensors D76 and D127 was made possible by using two complementary GPCR reporting systems: a high-throughput [Ca²⁺]_i-imaging assay (Gαi3q) and a high-precision patch-clamp assay (Gβγ-GIRK) (Fig. 4 and SI Appendix, Figs. S7–S11). Our GPCR-Gαi3q-[Ca²⁺]_i assay used HeLa cells coexpressing P2Y₁₂, the Gαi3q chimera, and GCaMP3, which reported the function of P2Y₁₂ by cytosolic [Ca²⁺]_i using fluorescence imaging for screening Vm-P2Y₁₂ sites (Fig. 4 A–E and SI Appendix, Figs. S7 and S8). This assay is superior to previous assays (42, 52, 53) for its higher throughput using real-time imaging. Our high-precision patch-clamp assay consisted of HEK293A cells coexpressing P2Y₁₂ and GIRK1/4 ion channels, which reported P2Y₁₂ function as GIRK currents for Vm-GPCR analysis (Fig. 4 F–K and SI Appendix, Figs. S9–S11, see also refs. 41, 43). In principle, these two complementary assays can be used to study the Vm dependence and/or other functions of P2Y₁₂ (and other GPCRs, with slight modifications).

In contrast to the present findings of Vm-dependent QS in rodent neuroendocrine chromaffin cells, previously Parnas and Parnas have found the Vm-sensing sites responsible for M2-mediated GIRK currents in oocytes and provided pharmacological evidence supporting a Vm-dependent M2 effect on peripheral excitatory postsynaptic currents (26). Mahaut-Smith and colleagues found that Vm-dependent binding between ADP and P2Y₁ modulates [Ca²⁺]_i in nonexcitable blood megakaryocytes, with pharmacological evidence supporting Vm-dependent P2Y₁ (62). These earlier reports, however, lacked crucial evidence of the Vm dependence of GPCRs (M2 or P2Y₁) by genetic knockdown and rescue in native cells. In the present study, we demonstrated not only the phenotypes of the Vm dependence of catecholamine release via P2Y₁₂-AIQS (Figs. 1–3) and the molecular mechanisms (Fig. 4), but also crucial validation using genetic KO, KD, and rescue (Figs. 3 and 5). Particularly, we not only identified the Vm-sensing sites of P2Y₁₂ but also confirmed them in native ACCs (Figs. 2–5). These provide an example of physiological Vm per se having a direct physiological impact on quantal vesicle release in both cultured (Figs. 1–3 and 5 and SI Appendix, Figs. S1, S3, S4, S6, and S13) and slice ACCs (Fig. 6 A–C). The sodium ion (Na⁺) is known to allosterically modulate GPCR activation by binding the highly conserved TM2 aspartate residue in family A GPCRs (63–65). The present work demonstrated that the aspartate residues of TM2 (D76) and TM3 (D127) are the voltage sensors of P2Y₁₂ and regulate exocytosis in native cells and slices (Figs. 4–6), establishing the essential physiological relevance of GPCR voltage dependence, probably by influencing the interaction between the Na⁺ pocket of D76 and the DRY motif of D127 (66).

Since the structure and function of GPCRs are well conserved in all tissues, in addition to P2Y₁₂ regulating catecholamine release, future work is needed to determine: 1) the Vm dependence of other phenotypes (i.e., event number) by P2Y₁₂, or other GPCRs and physiological functions, including released vesicle cargos of synaptic neurotransmitters, neurotrophins (nerve growth factor, insulin), inflammatory factors (NF-κB, substance P), and other hormones (5-HT, dopamine); and 2) whether Vm-GPCR-QS also exists in the central nervous system, other nonneuronal systems, and other animals including humans.

The present work establishes a signal pathway linking excitation-quantal size (QS), Vm → GPCR-P2Y₁₂ → Giβγ → fusion pore → QS (Fig. 6G), which coexists with the canonical pathway linking excitation-quantal release, Vm → Ca²⁺ influx → quantal release in sympathetic chromaffin cells. As Vm and GPCRs are present in all neuronal and nonneuronal cells, the regulation of GPCR and its downstream signals (i.e., vesicle release) by Vm could affect physiological/pathological functions beyond adrenal chromaffin cells and sympathetic nervous system.

Materials and Methods

Animals and Chemicals.

The P2Y₁₂-KO mice on a C57BL/6 background were gifts from Junling Liu, Shanghai Jiaotong University, Shanghai, China, and maintained in the Animal Center of Peking University. Sprague-Dawley rats (adult, 150 g) were from Beijing Vital River Laboratory Animal Technology Co., Ltd. All procedures and animal care were approved by the Institutional Animal Care and Use Committee of Peking University (Beijing, China) and the Association for Assessment and Accreditation of Laboratory Animal Care. Adult mice (C57BL/6 strain, 1 to 4 mo old, both sexes) were used for all experiments. Mice and rats were housed under a 12-h light/dark cycle with food and water. The details of all animals and chemicals are in SI Appendix, Table S1.

Plasmids.

The full-length rat P2Y₁₂ receptor (P2Y₁₂, NM_022800) was subcloned into pIRES2-EGFP (Clontech) or p3XFLAG-CMV (Sigma). The P2Y₁₂ site mutations D76N, D127N, D76E, and D127E were produced by PCR using a QuikChange II Site-Directed Mutagenesis Kit (Agilent Technologies). The nucleotide target sequences GCA GTA AAT CGA ACT TCA TCA (P2Y₁₂-shRNA1) and GCT TCG TTC CCT TCC ACT TTG (P2Y₁₂-shRNA2) were chosen to silence the expression of P2Y₁₂. A random sequence (TTC TCC GAA CGT GTC ACG T) that was predicted to target no genes in human, rat, and mouse cells served as a negative control (scrambled) (Guangzhou RiboBio Co., Ltd). Annealed double-stranded oligonucleotides encoding the target sequences were inserted into the vector pRNAT-H1.1-RFP/GFP to generate plasmids expressing shRNAs against P2Y₁₂. An RNAi (P2Y₁₂-shRNA1)-resistant form of rat P2Y₁₂ for the rescue experiments was generated by introducing the following silent mutations: GTA GCA AGT CAA ATT TTA. All constructs were verified by DNA sequencing. The bidirectional expression vector pBI-CMV1 (a kind gift from D. E. Logothetis, Virginia Commonwealth University, Richmond, VA) was used to simultaneously and constitutively express GIRK1 and GIRK4. The Gαi3q was a kind gift from Xiao Yu, Shandong University, Shandong, China. The GCaMP3 was from Addgene Co. The NPY-pHluorin plasmid was constructed from NPY-Venus (a kind gift from Nikita Gamper, University of Leeds, Leeds, UK).

Cell Culture and Transfection.

Rat adrenal chromaffin cells (ACCs) were prepared as described previously (19, 67) and mouse ACCs were prepared similarly with minor modifications. Briefly, the adrenal glands were isolated from anesthetized animals (10% chloral hydrate), cut into pieces and, after removing the cortex, incubated in papain solution for 40 min at 37 °C. The pieces were then triturated gently through a 200-μL pipette tip. After centrifugation, cells were quickly plated on coverslips precoated with 0.1% poly-l-lysine, incubated at 37 °C under 5% CO₂, and used within 24 to 96 h. ACCs were transfected for genetic manipulations using a 10-μL Neon electroporation system (Invitrogen, MPK1096) according to the manufacturer’s instructions (68).

HEK293A cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and transfected with VigoFect (Vigorous Biotechnology Beijing Co.) using the following plasmids (per 3.5-cm diameter dish): For gene-silencing experiments, shRNAs (2 μg) and their target P2Y₁₂ (1 μg) were delivered into cells and Western blotting analysis was performed 4 d later. To measure GIRK currents, rat P2Y₁₂ (1 μg) and a bicistronic plasmid expressing the GIRK1 and GIRK4 subunits (1 μg) were transfected and after 24 h of expression, the HEK293A cells were placed on sterile, poly-l-lysine–coated glass coverslips and the GIRK current was recorded.

HeLa cells were cultured in DMEM supplemented with 10% FBS and transfected with P2Y₁₂, Gαi3q, and GCaMP3 (1 μg each) for 36 h using Lipo2000 (Invitrogen) before Ca²⁺-imaging experiments. The coexpression rate of the three plasmids in cells was ∼85%.

Adrenal Slice Preparation.

Adrenal slices were prepared as described previously (13, 69). Briefly, adrenal glands were removed from adult mice (2 to 4 mo old) and immediately placed in ice-cold cutting solution containing (in mM): 125 NaCl, 2.5 KCl, 2 CaCl₂, 1 MgCl₂, 1.25 NaH₂PO₄, 26 NaHCO₃, and 10 glucose (pH 7.4, saturated with 95% O₂ and 5% CO₂). Then, a single gland without fat was glued with 3% agarose to the stage of a vibratome (Leica VT 1200S) and cut into 150-µm slices. The slices were incubated for at least 30 min at 37 °C in cutting solution and then kept at room temperature before recordings. Slices were used within 12 h after cutting.

Electrophysiology.

An EPC10/2 amplifier with Patchmaster software (HEKA Elektronik) was used to obtain whole-cell patch-clamp recordings as described previously (40). The normal extracellular buffer was composed of (in mM): 145 NaCl, 2 CaCl₂, 2.8 KCl, 1 MgCl₂, 10 Hepes (pH 7.4). For the Ca²⁺-free solution (0Ca), 2 mM Ca²⁺ was replaced by 1 mM EGTA for cultured chromaffin cells or 10 mM EGTA for adrenal slices. Patch pipettes were filled with intracellular solution containing (in mM): 100 K⁺-aspartate, 40 KCl, 5 NaCl, 7 MgCl₂, 10 EGTA, 0.025 GTP, 5 Na⁺-ATP, 20 Hepes (pH 7.2). For GIRK current recordings, a high-K⁺ buffer was used as the extracellular solution (as above, but containing 140 mM K⁺ and 2.4 mM Na⁺) (43). GIRK currents were measured in the whole-cell configuration as inward currents (holding potential: −40 mV, −70 mV, or –100 mV) (43). For pure Ca²⁺ current recording, 2 mM CaCl₂ was increased to 10 mM CaCl₂ and 1 μM tetrodotoxin was added to the extracellular solution. The intracellular pipette solution contained (in mM) 153 CsCl, 1 MgCl₂, 10 H-Hepes, and 4 Mg-ATP, pH 7.2. During experiments, patched cells were continuously superfused with extracellular buffer or agonist-containing solution. Igor software (WaveMetrics) was used for all offline data analyses. All experiments were performed at room temperature unless otherwise indicated.

Electrochemistry.

Highly sensitive, low-noise, 7-μm carbon fiber electrodes (ProCFE, Dagan) were used for the electrochemical monitoring of quantal release of catecholamines from single ACCs as described previously (2, 19). Quantal events were analyzed as described previously (2, 19). Stimulation solutions (caffeine or other drugs) were delivered by a perfusion system (Yibo) with a fast exchange time (<100 ms).

TIRF Imaging.

TIRF images were captured on an inverted microscope with a 100× TIRF objective lens (numerical aperture, 1.45; Olympus IX-81) at an exposure time of 50 ms using an Andor electron-multiplying charge-coupled device with Andor iQ software. The temperature was kept at ∼35 °C throughout all TIRF experiments using a laboratory-made heater. Kiss-and-run and full-fusion-like events were defined as follows: in kiss-and-run events, the fluorescence signal was restricted to the center of the release site; in full-fusion-like events, the fluorescence signal diffused to the surroundings of the release site (SI Appendix, Fig. S4B and see also refs. 8, 13, 40). Exocytotic events were stimulated by puffing 20 mM caffeine and analyzed using ImageJ (NIH) as described previously (13).

Ca²⁺ Imaging.

For Ca²⁺ imaging using Fura-2, isolated chromaffin cells were incubated in a bath solution containing 5 μM Fura-2/AM (Molecular Probes) for 15 min at 37 °C. The [Ca²⁺]_i was measured by dual-wavelength ratiometric fluorometry. The Fura-2 was excited by light alternating between 340 nm and 380 nm using a monochromatic system (TILL Photonics), and the emission fluorescence was measured using a cooled charge-coupled device. The [Ca²⁺]_i was calculated from the ratio of F₃₄₀ to F₃₈₀, and the sampling frequency was 1 Hz, triggered by the HEKA amplifier. The Ca²⁺ fluorescence signals were analyzed using Igor software (Wavemetrix) (19).

For Ca²⁺-imaging using GCaMP3, HeLa cells (4th to 12th generation after recovery from freezing) were transfected with three plasmids: P2Y₁₂, Gαi3q, and GCaMP3 (1 μg each) for ∼36 h using Lipo2000. The P2Y₁₂-specific agonist 2MesADP (10 µM) was puffed onto the cells to trigger [Ca²⁺]_i oscillations (typically one to three spikes, SI Appendix, Fig. S7E). All of the cells were from three to six batches; 10 to 20 cells per batch for each condition (control or mutations) were imaged for statistics. Considering that the peak of the [Ca²⁺]_i oscillation represents the strength of P2Y₁₂-Gαi3q signaling, the amplitude of [Ca²⁺]_i in a cell was defined as the maximum peak during the 90-s stimulation (SI Appendix, Fig. S7E). [Ca²⁺]_i was measured by ΔF/F₀, where F₀ is basal fluorescence before 2MesADP stimulation and ΔF is the fluorescence change during a [Ca²⁺]_i spike. The [Ca²⁺]_i signals were captured on an inverted confocal microscope (Zeiss 710) using a 488-nm laser and the light emitted from GCaMP3 was recorded at 500 to 540 nm. Time series videos (512 × 512 pixels) were acquired at 1 Hz under a 40× oil objective lens (Zeiss). Data were analyzed with ImageJ.

Excitation–Contraction Coupling in Myocytes.

Myocytes were isolated from adult C57 mice (1.5 to 2 mo old, weight ∼25 g) (70–72). A bipolar electrode (laboratory made) was used to evoke contractions; a brief, 50-V, 5-ms pulse (Nihon Kohden, Electronic Stimulator SEN-3201, Isolator SS-102J) was applied for pacemaking. The contractions evoked before and after drug treatments were imaged under an inverted IX-81 Olympus microscope, and analyzed with ImageJ.

Western Blot.

Samples were lysed with lysate buffer containing 20 mM Hepes at pH 7.4, 100 mM KCl, 2 mM EDTA, 1% Nonidet P-40, 1 mM phenylmethanesulfonyl fluoride, and 2% proteinase inhibitor (539134, Calbiochem). The homogenate was centrifuged at 15,000 × g for 30 min and the supernatant was collected and boiled in sodium dodecyl sulfate–polyacrylamide gel electrophoresis buffer. Proteins were electrophoresed and transferred to nitrocellulose filter membranes. Each membrane was blocked by incubation for 1 h with PBS containing 0.1% Tween-20 (vol/vol), and 5% nonfat dried milk (wt/vol). After washing with 0.1% Tween-20 containing PBS (PBST), the blots were incubated with primary antibodies at 4 °C overnight in PBST containing 2% bovine serum albumin (BSA). Secondary antibodies were then applied at room temperature and left for 1 h. Blots were scanned with an Odyssey infrared imaging system (LI-COR Biosciences) and quantified with ImageJ. The primary antibodies were as follows: rabbit anti-P2Y₁₂ (Anaspec, AS-55043A), mouse anti-flag (F1804, Sigma), and mouse anti–β-actin (A5316, Sigma); the secondary antibodies were IRDye 800CW goat anti-rabbit IgG (LIC-926-32211, LI-COR Biosciences), and IRDye 680CW goat anti-mouse IgG (LIC-926-32220, LI-COR Biosciences).

Immunofluorescence.

Cells were prefixed in 4% paraformaldehyde for 15 min, washed three times with PBS, then permeabilized with 0.3% Triton in PBS for 3 min. After the cells were incubated with 2% BSA for 1 h, they were incubated with the primary antibody rabbit anti-P2Y₁₂ (Anaspec, AS-55043A) overnight at 4 °C. Then they were washed with 2% BSA in PBS, and incubated with the secondary antibody (Alexa Fluor 594 goat anti-rabbit IgG, A11037, Invitrogen). After that, the cells were mounted on coverslips immersed in 50% glycerol. Fluorescence images were acquired on a confocal microscope (Zeiss 710) and analyzed with ImageJ.

Reverse Transcription PCR.

Total RNA was extracted using the TRIzol reagent (Invitrogen) according to the manufacturer’s instructions and mRNA was reverse transcribed with the Transgen kit (AU311). The forward and reverse oligonucleotide primers we have used are listed in SI Appendix, Table S2.

Statistics.

All experiments were replicated at least three times. Data are shown as the mean ± SEM or the median with interquartile range. “n” represents the number of independent experiments as reported in the figure legends. All data were tested for normality prior to choosing a proper statistical test. If the data passed the normality test, the paired Student’s t test was applied for comparison between two matched groups and one-way ANOVA followed by Tukey’s multiple comparisons test was applied when multiple groups were compared with one variable. If the data did not pass normality, the Wilcoxon matched-pairs signed rank test (Wilcoxon test) was applied for comparison between two matched groups and the Friedman test followed by Dunn’s multiple comparisons test was used for multiple matched groups. The Kolmogorov–Smirnov (K-S) test was applied for cumulative distribution comparison between two groups. All tests were conducted using Prism V7.0 (GraphPad Software, Inc.) and SPSS 20.0 (Statistical Package for the Social Sciences). Statistical tests were two-tailed and the level of significance was set at P < 0.05 (*P < 0.05, **P < 0.01, ***P < 0.001).

Data Availability.

All study data are included in the article and supporting information.