Single Calcium Channel Nanodomains Drive Presynaptic Calcium Entry at Lamprey Reticulospinal Presynaptic Terminals

Dissociated axons retain functional terminals

Precise measures of evoked presynaptic Ca²⁺ influx requires direct recordings from the release face membrane of individual presynaptic terminals (Fig. 1A). Such recordings impeded in vivo at more conventional central synapses with single AZs because of the close apposition between the presynapse and postsynapse have, thus far, been possible only at acutely isolated specialized calyceal-type terminals (Stanley, 1991, 1993; Sheng et al., 2012). Using a similar approach to accessing the presynaptic VGCCs, we developed an acute dissociation of the lamprey spinal cord (Ramachandran and Alford, 2014), by applying a combination of enzymatic digestion of spinal cord tissue and acute mechanical dissociation, enabling isolation of reticulospinal axons with functional presynaptic terminals without any postsynaptic processes (Fig. 1B). This permits direct recordings from single AZs. Presynaptic terminals maintained evoked synaptic vesicle cycling via exo-endocytosis visualized by FM1-43 staining of recycling vesicle clusters during 30 mm KCl depolarization (Fig. 1C; Betz and Bewick, 1992; Photowala et al., 2005) enabling targeted recordings of Ca²⁺ currents from fluorescently identified single terminals (Fig. 1C,D). Excess FM dye was washed off with Advasep-7 (Kay et al., 1999). Notably, we recorded only from presynaptic terminals with simple AZs (diameter 0.4–2.8 µm, mean 1.6 ± 0.2 µm), with single release sites and univesicular release (Gustafsson et al., 2002; Photowala et al., 2006; Schwartz et al., 2007), allowing for correlations of Ca²⁺ measures presented in this work to the fusion requirements of a single synaptic vesicle. Dissociated axons retained structural integrity, verified by patching the axons in whole-cell patch clamp configuration and gently pressure loading the axons with the fluorescent dye Alexa Fluor 488 hydrazide. Loaded axons were monitored over 30 min for dye leakage, demonstrating uniform loading throughout the axon with no decrease in fluorescence over time (Fig. 2A). Dissociated axons retained physiological membrane potentials (−58 ± 1.6 mV, n = 17 axons) and the capacity to fire APs (n = 7 axons; Fig. 2B). They also retained evoked compensatory endocytosis shown by labeling of synaptic vesicle clusters with FM1-43 following stimulation with high K⁺ containing solution (Fig. 2C) and could be similarly labeled with an antibody against Synaptotagmin which demonstrated punctate localization (Fig. 2D).

• Download figure
• Open in new tab
• Download powerpoint

Figure 1.

Direct recording of VGCC currents at the release face membrane of individual AZs. Aa, Schematic of the lamprey spinal cord showing reticulospinal axon/spinal neuron synapses demonstrating paired recording. Ab, Image depicting the architecture of the reticulospinal synapse. Presynaptic axon is colored pink, presynaptic terminals are in green (from intracellular phalloidin labeling of actin surrounding vesicle clusters in the axon). Postsynaptic projections (gray) contact presynaptic terminals. Ac, Magnified image of black inset box in Ab. B, Giant axons were acutely dissociated. Representative image showing the dissociated reticulospinal axon preparation. Solid black circle indicates dissociated spinal cord end from which reticulospinal axons emerge out. Arrowheads (black) indicate reticulospinal axons with no apposing postsynaptic processes. C, Representative image of cell-attached patch clamp recording at a single presynaptic AZ of a dissociated reticulospinal axon labeled with FM1-43 (green puncta). Dashed white lines outline the pipette position. White arrowhead indicates nonpresynaptic terminal location. D, Schematic showing placement of a cell-attached electrode over the AZ identified by FM1-43 labeling of recycling vesicles. These are present in a defined cluster immediately adjacent to the AZ colocalized with actin labeled with phalloidin (Ab, Ac). The vesicle cluster margins extend beyond the AZ indicating that the patch electrode covers the entire structure.

Immunohistochemical characterization of multiple VGCC subtypes at reticulospinal AZs

We validated the retention of VGCC localization, after dissociation, at single presynaptic terminals using antibodies specifically labeling three different Ca²⁺ channel subtypes, N-type (CaV2.2), P/Q-type (CaV2.1), and R-type (CaV2.3), previously implicated in synaptic transmission from lamprey reticulospinal axon (Krieger et al., 1999; Büschges et al., 2000; Photowala et al., 2005). For each of the VGCC subtypes tested, we were able to observe distinct punctate labeling by VGCC antibodies along the axons at discrete locations (Fig. 2E–G), consistent with a restricted localization of VGCCs along the axonal membrane surface, similar to the distribution of synapses on these axons in situ. We verified that these punctate distributions localized to presynaptic terminals by labeling presynaptic AZs in the fixed axons, post VGCC antibody labeling, with Alexa Fluor 633-conjugated phalloidin (Fig. 2E–G; Photowala et al., 2005; Bleckert et al., 2012). Specificity of the antibody labeling was verified by preincubating each of the primary antibodies, before application, with the corresponding control antigen. No antibody labeling was observed in the controls; nevertheless, we were still able to observe individual presynaptic terminals clearly identified by Alexa Fluor 633-conjugated phalloidin labeling (Fig. 2E–G). Similarly, controls for the secondary antibody did not result in any nonspecific labeling (Fig. 2H).

• Download figure
• Open in new tab
• Download powerpoint

Figure 2.

Dissociated reticulospinal axons retain functional AZs. A, Representative images showing Alexa Fluor 488 Hydrazide filling of dissociated reticulospinal axon (left) by pressure injection of the dye through a patch pipette (orange asterisk) in whole-cell patch configuration. Note 30 min later, the dye integrity is maintained within the axon (left panel on dye loading, right panel after 30 min), indicating that the axons remain structurally intact during and after dissociation. B, Representative AP (n = 7 axons) elicited in a dissociated reticulospinal axon by stimulating with a sharp microelectrode containing 3 m KCl. C, Representative image of a region in a dissociated reticulospinal axon showing punctate FM1-43-labeled recycling vesicle clusters, labeled by 30 mm KCl bath-perfusion stimulation. D, Antibody labeling against Syt-1. A lumenal domain biotinylated anti-Syt-1 antibody (rabbit, anti-mouse amino acid residues 1–8, SYSY 105 103BT) diluted 1:50 in Ringer was applied by pressure from a pipette over an isolated axon during 30 mm KCl stimulation of the axon. During subsequent stimulation, streptavidin-conjugated dye (Alexa Fluor 488 Hydrazide) was applied by pipette. Fluorescent puncta were again resolved. Control stimulation and application of streptavidin labeled dye but without prior application of antibody against Syt-1 revealed no puncta. E–G, Antibody labeling against N-type (CaV2.2; Ea), P/Q-type (CaV2.1; Fa), and R-type (CaV2.3) VGCC (Ea, Fa, Ga, left panels) demonstrating punctate distribution along the axon membrane. Note: primary antibodies against VGCC subtypes (rabbit, Alomone Labs), secondary antibody Alexa Fluor 633 Hydrazide conjugated (goat, anti-rabbit, Alomone Labs). The primary anti N-type antibody was against an intracellular epitope (C)RHHRHRDRDKTSASTPA, corresponding to amino acid residues 851–867 of rat CaV 2.2 (accession Q02294) located in the intracellular loop between Domains II and III of the VGCC. The primary anti-P/Q-type antibody was against an intracellular epitope (C)PSSPERAPGREGPYGRE, corresponding to amino acid residues 865–881 of rat CaV 2.1 (Accession P54282) located in the intracellular loop between Domains II and III of the VGCC. The primary anti-R-type antibody was against an intracellular epitope (C)SASQERSLDEGVSIDG, corresponding to amino acid residues 892–907 of rat CaV 2.3 (accession Q07652) located in the intracellular loop between Domains II and III of the VGCC. Presynaptic terminal locations were determined in the same axon by labeling presynaptic actin with Alexa Fluor 488 Hydrazide-conjugated phalloidin (Ea, Fa, Ga, center panels). Colocalization of VGCC labeling with locations of presynaptic terminals for each VGCC subtype characterized. Red indicates VGCC labeling, green indicates presynaptic actin labeling, and yellow colocalization (Ea, Fa, Ga, right panels). Scale bar in all images: 10 µm. n for each VGCC subtype: N-type 5 axons from 3 animals; P/Q-type 9 axons from 4 animals; R-type 3 axons from 2 animals. Representative images showing controls for the primary antibodies used for N-type, P/Q-type, and R-type channel staining. Primary antibody was preincubated with specific control antigen for 30 min before staining the preparation (Eb, Fb, Gb, left panels). Note the absence of any VGCC labeling in the controls. Eb, Fb, Gb, right panels, Presynaptic terminals location labeled by Alexa Fluor 488 Hydrazide-conjugated phalloidin labeling of presynaptic actin in same region of the axon as in the left panel. Scale bar in all images: 10 μm. n for each control: N-type control antigen 10 axons from 2 animals; P/Q-type control antigen 5 axons from 2 animals; R-type control antigen 3 axons from 2 animals. H, Representative image of control for secondary antibody (Ha) and presynaptic terminals location labeled by Alexa Fluor 488 Hydrazide-conjugated phalloidin labeling of presynaptic actin (Hb) in same region of the axon. n = 8 axons from 3 animals.

Characterization of VGCC subtype specific currents at individual presynaptic terminals

The acutely dissociated preparations provide unhindered access to direct recording at the release face membrane of single presynaptic terminals. Presynaptic terminals were identified by fluorescent labeling with FM1-43 during evoked depolarization by perfusing 30 mm KCl. This allowed us to target individual AZs for cell-attached patch clamp recordings using fluorescence microscopy (Fig. 1C). We recorded Ca²⁺ currents at the release face membrane of individual fluorescently labeled AZs, with 10 mm [Ca²⁺]_external in patch pipette (Fig. 3A, n = 5 patches), using custom low-noise single channel cell-attached patch clamp recordings (Ramachandran and Alford, 2014). Single AZs were patched in cell-attached configuration, and the holding potential was set to −88 mV (adjusted from the known intracellular membrane potential determined from intracellular sharp electrode recording). The patch solution contained channel specific blockers, blocking all other known ionic currents (Materials and Methods), permitting the isolation of Ca²⁺ currents. We validated that the currents recorded represented Ca²⁺ influx through VGCCs by additionally incorporating Cd²⁺ in the patch solution (Swandulla and Armstrong, 1989). Pharmacological block of all VGCCs with Cd²⁺ (Fig. 3B, n = 8 patches) eliminated all currents, leaving the Gaussian distribution of the amplitude histogram indistinguishable from background noise. Voltage dependent Cd²⁺ block was reversed on repolarization shown by tail currents at the end of the pulse (Fig. 3Bb). Additionally, patch recordings obtained from nonpresynaptic terminal locations, identified by absence of fluorescent FM1-43 puncta along the axon membrane, demonstrated no VGCC currents (Fig. 3C, n = 3 patches), corroborating our previous findings of VGCCs being restricted to presynaptic AZs at these synapses (Photowala et al., 2005; Bleckert et al., 2012).

• Download figure
• Open in new tab
• Download powerpoint

Figure 3.

Cell-attached recordings from AZs reveal multiple VGCC subtypes. Aa, Representative sweep from a cell-attached patch recording showing Ca²⁺ currents recorded from a single AZ following a voltage step to 0 mV (n = 5 patches). For all Ca²⁺ current traces shown, inward currents have a negative directionality. Solid line marked 0 indicates channel closed state, dashed lines indicate incremental channel opening events corresponding to the opening of 1, 2, or 3 channels. Patch pipette contains 10 mm [Ca²⁺]_external as charge carrier. Ab, Amplitude histogram (gray bars) fitted by a sum of three Gaussian function (magenta fit line), indicates the opening of one to three Ca²⁺ channels during the indicated recording. Ba, Similar cell-attached recordings, from single AZs, following a voltage step to 0 mV with 500 μm Cd²⁺ and 10 mm [Ca²⁺]_external in the patch pipette demonstrating absence of inward Ca²⁺ currents. Bb, Amplitude histogram (gray bars; n = 8 patches) fitted with a single Gaussian function (magenta fit line) indicating only a current noise peak and no channel events. Inset, Tail currents measured, on repolarization to −80 mV from the voltage step during release of Cd²⁺ block, plotted as amplitude histogram (gray bars). The amplitude histogram was fitted by a sum of three Gaussian function (magenta fit line), indicating the opening of one to three Ca²⁺ channels on repolarization. Ca, Cell-attached recordings from nonpresynaptic terminal locations (Fig. 1C, white arrowhead), following a voltage step to 0 mV, demonstrating absence of inward Ca²⁺ currents. Cb, Amplitude histogram (gray bars, n = 3) fitted with a single Gaussian function (magenta fit line) indicating only a noise peak and no channel events. D, Depolarization stimulus protocol, applied from a holding potential of −80 mV, to elicit VGCC currents in cell-attached patch recordings. Red bar at the holding potential level indicates where the background current noise was measured (current noise record). Blue bars indicate the voltages at which analysis of single channel currents (current records) was conducted: step depolarizations to −30, 0, and 30 mV, and on repolarization to −80 mV (tail currents). E, Representative examples of N-type (CaV2.2), P/Q-type (CaV2.1), R-type (CaV2.3), and L-type (CaV1) currents, recorded at single AZs, following voltage steps to −30, 0, and 30 mV. Solid line marked 0 indicates channel closed state, dashed lines indicate incremental channel opening events, with maximum number of simultaneously open channels indicated in each representative trace. Fa, Cell-attached recordings from single AZs, following a voltage step to 0 mV, showing absence of depolarization evoked inward currents when all VGCC subtypes (N, P/Q, R, and L) were blocked. Fb, Amplitude histogram (gray bars, n = 4) fitted with a single Gaussian function (magenta fit line) indicating only a current noise peak and no channel events. G, Current-voltage relationships plotted for each of the indicated Ca²⁺ channel subtypes. For each channel subtype, mean single channel amplitude measured at corresponding voltages is shown along with the linear fit (N-type green, P/Q-type purple, R-type orange, L-type blue). Error bars indicate SEM. Linear fits have been extrapolated to intersect the x-axis indicating the reversal potential (E_rev). H, Mean open probability (mean P_open) plotted against membrane voltage (mV) for each of the indicated Ca²⁺ channel subtypes (N-type green, P/Q-type purple, R-type orange, L-type blue).

Multiple channel subtypes have been implicated in depolarization-evoked Ca²⁺ influx for vesicle fusion and neurotransmitter release (Wheeler et al., 1994; Mintz et al., 1995; Poncer et al., 1997; Wu et al., 1999; Cao and Tsien, 2010; Sheng et al., 2012). In lamprey, this is supported by our immunohistochemical analysis as well as previous imaging studies in the intact spinal cord in situ (Photowala et al., 2005). Therefore, we isolated VGCC subtype-specific currents pharmacologically (Materials and Methods), corresponding to each of the four different VGCC subtypes: N-type (CaV2.2, n = 6 patches), P/Q-type (CaV2.1, n = 5 patches), R-type (CaV2.3, n = 6 patches), and L-type (CaV1, n = 4 patches) with 10 mm [Ca²⁺]_external (in patch pipette), eliciting Ca²⁺ currents (Fig. 3D,E). Gaussian fits to amplitude histograms of current recordings, on combined selective blocking of all four VGCC subtypes and other known ionic currents, demonstrated no additional currents with a distribution similar to background noise (Fig. 3F, n = 5 patches). Channel openings were observed only on depolarization to a membrane voltage of −30 mV (Figs. 3E, 4A), indicating the absence of any low-voltage-activated T-type (CaV3) channels at these presynaptic terminals. We measured single channel amplitude (1), from current recordings for each channel subtype that demonstrated the opening of only a single channel, at depolarized voltages of −30, 0, 30 mV and on repolarization to −80 mV (tail currents; Fig. 3D). Single channel amplitudes at 0 mV were 0.28 ± 0.01 (N-type), 0.29 ± 0.03 (P/Q-type), 0.28 ± 0.02 (R-type), and 0.28 ± 0.02 pA (L-type; Fig. 3G; Table 1). Single channel conductance was calculated to be 2.97 ± 0.62 pS (N-type), 1.82 ± 0.36 pS (P/Q-type), 1.94 ± 0.41 pS (R-type), and 2.4 ± 0.75 pS (L-type; Fig. 3G; Table 1). Mean channel opening probability (P_open) at 0 mV was 0.26 ± 0.02 (N-type), 0.29 ± 0.03 (P/Q-type), 0.22 ± 0.02 (R-type), and 0.24 ± 0.02 (L-type; Fig. 3H; Table 2).

• Download figure
• Open in new tab
• Download powerpoint

Figure 4.

Few open channels from a small available VGCC pool mediate Ca²⁺ influx. A, Gaussian fits for the amplitude histograms (gray bars) for the indicated Ca²⁺ channel subtypes (10 mm [Ca²⁺]_external) following voltage steps to −30, 0, and 30 mV. Tail currents were measured, within a window of 20 ms immediately after repolarization to −80 mV, and plotted as amplitude histograms (gray bars). Inset (in N-type −80 mV) shows representative example of tail current channel opening events. Sum of Gaussian fit to the amplitude histogram is shown as solid line (N-type green, P/Q-type purple, R-type orange, L-type blue). Insets in histogram plots provide magnified view of the histogram (note expanded y-axis) showing bin values for higher current amplitudes that are not discernable from the full scaled histogram. B, Representative current traces, from cell-attached patch clamp recordings at single AZs with 90 mm [Ba²⁺]_external in the patch pipette, following voltage steps to −50, −30, 0, and 30 mV. Solid line in all the representative traces indicates the closed state of the channel. Dashed lines indicate incremental channel opening events, with maximum number of simultaneously open channels indicated in each representative trace. Amplitude histograms are shown in gray and the sum of Gaussian fit line in blue. Note the absence of any channel openings at a membrane voltage of −50 mV, one to two channels at −30 mV, and one to three channels at 0 and 30 mV.

Few open channels from a small available VGCC Pool mediate Ca²⁺ influx

We next enumerated Ca²⁺ channels at single presynaptic AZs for each Ca²⁺ channel subtype. The current data from replicate sweeps, for each analyzed voltage, in a patch recording were binned to generate amplitude histograms, which were averaged and fitted with a sum of Gaussians function. Using this approach, we enumerated the maximum number of channels, visually confirmed by inspection of the channel opening events, that simultaneously opened at each analyzed voltage during a patch recording. A maximum count of 10 (4–10, mean 5) N-type, 9 (3–9, mean of 6) P/Q-type, 32 (4–32, mean 12) R-type, and 17 (3–17, mean 10) L-type channels yielded a total maximum VGCC count of 68 channels (14–68 channels, mean 33; Fig. 4A; Table 3) at a presynaptic AZ. Restricting the count to only N + P/Q + R channel subtypes, that are implicated in evoked synaptic transmission at lamprey reticulospinal synapses, we obtained a maximum total of 11–51 channels (mean 23; Fig. 4A; Table 4) that may contribute Ca²⁺ to drive release at a single AZ.

Having determined the available pool of VGCCs at a single AZ, we next asked how many VGCCs open on depolarization, when all channel subtypes are available for opening. We used voltage steps with the same voltage range as seen during presynaptic APs recorded directly in situ (Cochilla and Alford, 1997) and thus the same voltage range to when an AP evokes presynaptic Ca²⁺ influx. We conducted cell-attached recordings from single presynaptic AZs (Fig. 1C,D), with 90 mm [Ba²⁺]_external (in patch pipette). As for our previously described recordings with 10 mm Ca²⁺, Ba²⁺currents were evoked only on depolarization to −30 mV. Notably, on depolarization to 30 mV, which is close to the peak of APs, we observed the opening of only one to seven (mean 4) channels, out of the available pool of a maximum of 68 (mean 33) channels at an AZ (Fig. 4B; Table 5, n = 5 patches), suggesting very few out of the available small pool of channels mediate Ca²⁺ influx at depolarization voltages that are equivalent to peaks of APs.

Molar quantitation corroborates Ca²⁺ entry through few open VGCCs

Evoked openings of VGCCs in cell-attached recordings were determined after acute isolation. A similar approach has been used in calyceal synapses (Sheng et al., 2012). However, acute isolation may alter channel activity. To confirm that evoked Ca²⁺ entry remains the same in isolated axons to those in situ we used quantitative imaging to determine molar quantities of Ca²⁺ that entered each AZ during each AP. We used Ca²⁺ dyes as buffers to quantify total molar quantities of AP-evoked Ca²⁺ entry to the reticulospinal axons maintained in the intact spinal cord in situ as an independent measure of axonal evoked Ca²⁺ entry with no dissociation. Ca²⁺ dyes with calibrated binding properties were injected directly into axons at known concentrations, before recording AP-evoked Ca²⁺ transients (Materials and Methods). Ca²⁺ binding properties of the Ca²⁺ dyes, fura-2, Oregon Green BAPTA1, and Fluo-5F from the lots used experimentally, were determined using Ca²⁺ standards (Invitrogen). Log plots of these data points were used to determine K_d as well as fluorescent maximum/minimum ratios (Fig. 5A; K_d = 326 nm for fura-2, 450 nm for Oregon Green BAPTA1 and 1.12 μm for Fluo-5; measured at pH 7.2 and 10°C).

• Download figure
• Open in new tab
• Download powerpoint

Figure 5.

Quantifying AP-evoked molar Ca²⁺ entry to AZs. A, Dyes from the lots used from the experiments were calibrated using the same light path, temperature, and pH of solution as during experiments. Graph shows plots of log₁₀{(R – R_min)/(R_max – R)} for ratiometric fura-2 or log₁₀{(F – F_min)/(F_max – F)} for single wavelength Fluo-5F and Oregon Green BAPTA1 versus log₁₀[Ca²⁺] from EGTA buffered standards (Life Technologies). K_ds obtained from these plots shown in corresponding colors. B, Imaging was performed in axons in superfused intact spinal cords impaled with microelectrodes containing fura-2 (Life Technologies), Fluo-5F or Oregon Green BAPTA1. Dyes were in buffered KCl solution (pH 7.2). Imaging used either an upright epifluorescence microscope (ratiometric imaging) or confocal microscope (single wavelength dyes). In both cases the arrangement of objective (Olympus LUMPLF 40 × 0.8 NA), spinal cord, and recording/injection microelectrodes were identical. Axons were stimulated with AP trains via current injection through the microelectrode. Ca, Cb, Reticulospinal axons were loaded with dye be pressure injection through the recording microelectrode. Representative Ca²⁺ responses (Ca, Cb) show the effect of increased dye concentration on peak Ca²⁺ amplitude and decay rate (τ) from single exponential fits (black traces) to traces at lower (red) and higher (blue) concentration of dye. Cd, Alexa Fluor 594 Hydrazide-conjugated phalloidin (Life Technologies) labeled presynaptic AZs imaged after subsequent injection into the axon with a second electrode revealing their numbers in the imaged region. D, Dye concentrations in fura-2 filled axons were calculated by comparing fluorescence intensity of dye at known concentration in the microelectrode with dye labeled axons at the same depth in tissue. Da, For fura-2, images were taken with electrode diameter the same as the axon to allow imaging from the same volume exciting at the isosbestic point. Db, For nonratiometric dyes, Alexa Fluor 594 Hydrazide was imaged confocally at the same depth in tissue in adjacent axons and electrodes uses images substantially larger than the microscope point spread function. E, The rate of decay of Ca²⁺ transients (t) was plotted versus buffering capacity of the dye (k_dye) from exponential fits (Ca) over a range of k_dye. These data were well fit (r² = 0.88) with a linear function (dark red line, 95% confidence indicated by encompassing faint red lines) yielding an estimate of the endogenous buffering capacity (k_end) to be 13.1 ± 11.1 from Equation 10. F, Peak Δ[Ca²⁺], was plotted against a range of k_dye and fitted with Equation 11 (dark red line, 95% confidence indicated by encompassing faint red lines) yielding a peak Δ[Ca²⁺] of 31.6 ± 5.2 nm per stimulus from the intercept on the ordinate axis. This fit also yielded value of total Ca²⁺ concentration, bound and unbound (Δ[Ca]_total) of 109.9 ± 19.2 nm and a k_end of 2.5 ± 1.1. Equation 11 is a hyperbolic function. Thus, to confirm this fit, a line is fit to the plot of 1/peakΔ[Ca²⁺] against k_dye. The data are well fit to a line with r² = 0.90. G, The total Ca²⁺ entering bound to dye (Δ[Ca]_total = Δ[Ca²⁺] × κ_dye) for each AP was plotted against κ_dye and fitted by Equation 12 to yield a κ_end of 5.2 ± 2.2, an asymptotic value of Δ[Ca]_total representing full dye binding of all Ca²⁺ entry of 116.9 ± 13.2 nm. From these calculations (Table 6), we conclude a total charge entering the axon at each AZ during one AP is 11.2 fC which represents a current of 2.79 pA over 5 ms (the rise time of the response).

To measure presynaptic Ca²⁺ entry, spinal cords were isolated, intact and superfused with oxygenated Ringer's solution (10°C; Materials and Methods) and imaged under epifluorescence or confocal microscopy (Fig. 5B). Microelectrodes were filled with Ca²⁺ dyes and reticulospinal axons were impaled to fill them with dyes of differing affinities; either the ratiometric dye fura-2 or the nonratiometric dyes Fluo-5F or Oregon Green BAPTA1 (Life Technologies) in buffered 1 m KCl solution (pH 7.2 with 5 mm HEPES; Fig. 5C,D). Dye (from 0.1 to 1 mm in electrode) was pressure injected into the axon to obtain a range of dye buffering capacities (κ_dye; see Eq. 9) in the axons. After loading with dye, the axon was stimulated with a train of APs. The axon was allowed to recover between trials for 3–5 min.

Images were acquired for fura-2 fluorescence on a conventional epifluorescence microscope. Fluorescence transients from excitation at 380 nm were obtained after capturing baseline images from excitation at 350 and 380 nm and absolute Ca²⁺ concentrations calculated from Equation 6 (Materials and Methods). Examples of responses (Fig. 5Ca,Cb) evoked by 10 APs (Fig. 5Cc) at two different axonal fura-2 concentrations are shown to demonstrate that as the concentration, and therefore κ_dye, rises, the peak amplitude of free Ca²⁺ decreases and the time course (τ) of the decay increases. AP-evoked fluorescence transients were also obtained using Oregon Green BAPTA1 and Fluo-5F. For these, imaging was confocal with 488-nm excitation captured in long pass from 510 nm. Fluorescence maxima were measured during repetitive stimulation at 50 Hz until intensity reached an asymptotic maximum. Absolute Ca²⁺ concentrations were obtained from these data and Equation 7 (Materials and Methods). To obtain the amount of Ca²⁺ entering each AZ, we counted AZ numbers within the region of measurement. To achieve this, the Ca²⁺ dye/buffer electrode was withdrawn, and the same axon was re-impaled with an electrode filled with Alexa Fluor 594 Hydrazide-conjugated phalloidin. Phalloidin was pressure injected into the axon and images captured after 10–15 min allowing for the phalloidin to label presynaptic actin (Bleckert et al., 2012). From these images the number of phalloidin labeled puncta were counted (Fig. 5Cd). The mean number of AZs per axon region imaged was 79 ± 7 (0.40 ± 0.04 synapses per linear micrometer of axon).

Dye concentrations in the axons were also measured. For fura-2, at the end of the experiment, fluorescence at the isosbestic point (350- to 360-nm excitation) was obtained from the axon and the electrode in which the dye concentration was known (Fig. 5D). Because the conventional epifluorescence imaging captured out of focus light, the electrode was positioned adjacent to the imaged axon and such that the imaged portion was the same diameter as the imaged axon. to give the same imaged volumes. Axon dye concentrations were calculated as a fraction of the known concentration measured in profile plots (Fig. 5Da). A similar method was used to estimate Oregon Green BAPTA1 and Fluo-5F concentrations. For these confocal images, a known concentration of Alexa Fluor 594 Hydrazide was included in the electrode and co-injected into the axons. Electrodes and axons at the same depth were imaged using 560-nm excitation. The electrode was positioned such that its diameter was greater than the point spread function of the microscope (0.5 µm; Hamid et al., 2019). The fluorescence intensities of the electrode and of the cell were again compared (Fig. 5Db). We used Ca²⁺ dyes as buffers (Harrison and Bers, 1987) for which buffer capacities of dye injected into the axons at known concentrations were calculated. The buffering capacity (κ_dye) during each stimulated transient was calculated using Equation 9 (Augustine and Neher, 1992; Jackson and Redman, 2003; Hamid et al., 2019): κdye=Δ[Cadye]Δ[Ca2+]i=[Dyetotal]kd[(1 + [Ca2+]2kd)(1 + [Ca2+]1kd)],(9) where [Ca_dye] is the concentration of Ca²⁺-bound dye, Δ[Ca²⁺]_i is the evoked change in Ca²⁺ concentration, and [Dye_total] is the total dye concentration. [Ca²⁺]₁ and [Ca²⁺]₂ are the resting Ca²⁺ and peak Ca²⁺ concentrations before and during stimulation respectively (Fig. 5Ca,Cb).

The endogenous buffering capacity of the axon (κ_end) was determined from the dependency of the rate of decay the dye recorded Ca²⁺ transient to the buffering capacity of the dye at that time. The recovery of the stimulus evoked Ca²⁺ transient (Fig. 5Ca) was fitted with an exponential function and the decay rate (τ) measured, over a range of values of κ_dye calculated from Equation 9. Values of τ were plotted against κ_dye and fitted with a linear function (Fig. 5E, r² = 0.88) yielding an estimate of the endogenous buffering capacity (κ_end = 13.1 ± 11.1) from Equation 10, which describes the decay rate (τ) of a Ca²⁺ signal in a cell compartment (Neher and Augustine, 1992): τ=τext(1 + κend+ κ dye).(10)

This value of κ_end gives an estimate independent of the absolute measure of Ca²⁺ transient amplitudes and thus provides a reasonable measure of validity for the remaining calculations. We may also calculate the free Ca²⁺ concentration evoked by stimulation and the total molar quantity of Ca²⁺ that enters the axon. By measuring the peak amplitude of the free Ca²⁺ transient throughout the varicosity over a range of values of κ_dye (Neher and Augustine, 1992; Hamid et al., 2019), we may state: Δ[Ca2+]i=Δ[Ca]total(1 + κdye + κend),(11) where Δ[Ca²⁺]_i varies with κ_dye. Values of Δ[Ca²⁺]_i are computed from our data using Equation 6 or Equation 7 (Materials and Methods) and values of κ_dye from Equation 9. Peak Δ[Ca²⁺], per AP in the volume of axon, was plotted for a range of calculated κ_dye and fitted with Equation 11 (Fig. 5F) yielding a peak free Δ[Ca²⁺] of 31.6 ± 5.2 nm throughout the axon per stimulus from the intercept on the ordinate axis. This extrapolation is the true value of peak Δ[Ca²⁺]_i in the axons when no dye is present. This fit also yielded the value of total Ca²⁺ concentration, bound and unbound (Δ[Ca]_total) of 109.9 ± 19.2 nm that entered the whole axon volume and a κ_end of 2.5 ± 1.1. Equation 11 is hyperpolic, therefore the inverse of evoked free Ca²⁺ was plotted against κ_dye, giving a linear relationship. The straight line fit to this gives r² of 0.9. We can also calculate the total Ca²⁺ bound to dye for each stimulus against κ_dye. At higher dye concentrations the dye/buffer captures a greater proportion of entering Ca²⁺ reaching an asymptote as all entering Ca²⁺ is bound to dye. Thus, we plotted change in Ca²⁺ bound dye concentration (Δ[Ca]dye) against k_dye and fitted this with Equation 12: Δ[Ca]total=Δ[Cadye].{(κend + 1)κdye + 1}.(12)

This fit yields κ_end of 5.2 ± 2.2, and an asymptotic value of Δ[Ca]_total (the total stimulated Ca²⁺ entry) representing full dye binding of all Ca²⁺ entry of 116.9 ± 13.2 nm (Fig. 5G). Values obtained were similar from fits to Equations 11, 12 (mean values, Table 6). Note that during Ca²⁺ entry that occurs only at AZs, the local concentration of Ca²⁺ which evokes neurotransmission is transiently at much higher concentrations, but this high concentration almost immediately disperses throughout the larger axonal volume from which it was imaged. We calculated axon volumes from images, assuming the axons are cylindrical, to determine the molar quantity of Ca²⁺ that entered the axon per AP and the mean molar amount entering at each terminal can then be given by dividing this total by the number of synaptic terminals imaged using phalloidin (Fig. 5Cd). From these calculations (Table 6), we conclude a total charge entering the axon at each AZ during one AP is just 7.98 ± 0.90 fC representing a Ca²⁺ current of 1.6 ± 0.3 pA over the duration of the AP (2–3 ms of depolarization and 2 ms of estimated tail current). Mean channel currents over this voltage range were ∼0.4 pA (from the integral of slopes between −80 and +20 mV in Fig. 3G), indicating opening of a mean of four channels per AZ per AP. This is very similar to results from our cell-attached recordings.

Resolving presynaptic Ca²⁺ channel transients with LLSM

Ca²⁺ entry to lamprey reticulospinal axons is limited to synaptic AZs (Photowala et al., 2005), and we have demonstrated that few channels open at each AZ on presynaptic depolarization. Thus, variability of channel activation between APs may contribute to the stochastic nature of quantal release. To investigate these properties of presynaptic VGCCs in intact synapses physiologically activated by APs, we visualized Ca²⁺ transients at single presynaptic terminals in spinal cords in the intact spinal cord. Prior methods that provide sufficient temporal resolution to image single transients such as confocal line scanning (Takahashi et al., 2001) suffer from insufficient signal-to-noise characteristics to resolve variations in signal between single APs at each AZ. They also cause photodamage because high excitation powers are necessary to resolve the responses. To overcome these difficulties, we used a custom built LLSM (Chen et al., 2014; Potcoava et al., 2021; Fig. 6A). This eliminated out of focus noise and enabled optical slicing at sufficient rates to resolve presynaptic Ca²⁺ transients. It reduced the light dose compared with confocal imaging by ∼1000-fold. This enabled variations in AZ Ca²⁺ signaling to be compared between APs. Additionally, the near absence of photobleaching (Chen et al., 2014) allowed recording of many repetitions of stimulated transients (Chen et al., 2014) allowing us to investigate the stochastic nature of channel dependent presynaptic Ca²⁺ entry at single presynaptic AZs.

• Download figure
• Open in new tab
• Download powerpoint

Figure 6.

Recording presynaptic Ca²⁺ transients with LLSM. A, Lamprey spinal cords were isolated in a custom chamber (Aa) designed to allow superfusion, recording and stimulation of giant axons under two lenses of the LLSM (Ab). Axons filled with Ca²⁺ dye were imaged in single LLSM planes over time (Ac). B, Imaging single planes at 300 Hz during a single AP reveals Ca²⁺ hotspots. Stimulation of the axons with an extracellular stimulation electrode evoked Ca²⁺ transients at multiple discrete locations at the axon surface. An example of one hotspot (Ba), in response to single APs in the plane of focus of the recording, is shown when recording the same plane repeatedly at intervals indicated. Individual planes are shown as ΔF/F in pseudocolor and intensity mapped to the vertical dimension to emphasize the resolution of the hotspot and are averages of responses from five stimuli. Other hotspots not in focus are also seen. The summed Ca²⁺ entry from multiple hotspot locations diffuses throughout the axons within 10 ms of the stimulus. This diffusion is shown across the axon with a profile centered on the hotspot. Bb, A profile plot (between white lines of pseudocolor image at left). This profile is displayed for the same frames shown in Ba. C, The intensity of the center 1-µm diameter (green) and the surrounding 5 µm (red) across all frames is expressed as ΔF/F along with the nonhotspot whole axon response (black). The latter representing diffusion from all hotspots. These responses are overlaid (Ca). The diffusional background is subtracted (Cb) to reveal just the hotspot Ca²⁺ signal and its time course (green and red from the same locations as Ca). D, In repetitive stimulation (five stimuli at 50 Hz), the slower, whole axon diffusional signal was subtracted from the hotspot signal isolating Ca²⁺ hotspots (Da, blue and green, hotspots shown in insets; black, whole axon signal). This approach isolated responses even during bursts of stimuli (Db, colors as for Da).

Lamprey giant synapses are uniquely suited for this study, because they have simple AZs (Gustafsson et al., 2002) and a large axon into which Ca²⁺ can disperse (Cochilla and Alford, 1998; Takahashi et al., 2001), enabling resolution of Ca²⁺ transients from single AZs. To label reticulospinal axons with Ca²⁺-sensitive dye, axons in intact lamprey spinal cords were impaled with microelectrodes containing 5 mm Fluo-5F and pressure injected via the microelectrode (final concentration in axon ∼4 μm (Takahashi et al., 2001; Materials and Methods). We used LLSM (Fig. 6A) to image AP-evoked Ca²⁺ transients at AZs in situ in these axons at 330–800 Hz at fixed z-axis (Fig. 6Ac). Superficially, Ca²⁺ transients appear as single hotspots (Fig. 6Ba), that prior studies demonstrate colocalize to AZs (Photowala et al., 2005), although LLSM gave substantially greater signal-to-noise (mean LLSM rms peak signal-to-noise ratios = 41.1 ± 14.2 n = 10, compared with 4.0 ± 1.3 with line scanning n = 6). LLSM reveals readily resolved transients at single AZs during each AP. We characterized this Ca²⁺ entry and its distribution in 10 axons. To illustrate the response to single presynaptic AP, the average response to 10 sequential stimuli applied at 1 min intervals is shown (coded in color LUTs and the intensity profile represented in the z-axis of selected frames) from before and after stimulation (Fig. 6Ba). Frames were analyzed by subtracting the background signal outside the axon and expressing data as ΔF/F. In the recording shown (Fig. 6Ba), the LLSM lattice plane was placed over a single AZ (circled in red, frame at 3.2 ms), although other slightly out of focus hotspots also contributed to this initial signal (red arrows) whereby many AZs arranged around the axon plasmalemma contribute to the total Ca²⁺ entry. Ca²⁺ entered at AZs during stimulation and diffused throughout the 10-µm diameter structure within 10 ms [mean risetime (τ) of the Ca²⁺ signal at the center of the axon was 9.1 ± 1.6 ms]. This is shown graphically by calculating the profile signal in a region of interest (ROI) spanning the width of the optical section (Fig. 6Bb, white box) and plotting the profile before, immediately after and in subsequent frames (Fig. 6Bb).

To visualize the time course of the evoked responses, the transient taken from 0.5 × 0.5 µm (5 × 5 pixels) at the hotspot center is graphed (Fig. 6Ca, green) as well as the signal from the whole ROI in red (from ROI looping the hotspot at 3.2 ms; Fig. 6Ba). The mean signal from the axon, excluding hotspots is also shown (Fig. 6Ca, marked in black). It originates from the summed Ca²⁺ diffusing from all hotspots around the periphery of the axon, both in and out of focus. From phalloidin labeling data (Fig. 5A), a mean of 0.40 ± 0.04 synapses is present per linear micrometer of axon, thus on average, 20 AZs are present in this 50-µm region of axon. To demonstrate the amplitude and time course of Ca²⁺ entry to hotspots this overall Ca²⁺ signal derived from Ca²⁺ diffusing from all hotspots in this region of the axon was subtracted from the hotspot signals (Fig. 6Cb). We sought to investigate the variability of Ca²⁺ entry between single stimuli, and therefore used the same subtraction approach during bursts of repetitive stimulation. An example is shown for two hotspots (Fig. 6Da, blue and green) in the same axon (Fig. 6D, insets). Individual hotspots showed diffuse Ca²⁺ accumulation. The nonhotspot signal was again subtracted from this to reveal the time course of local Ca²⁺ entry and its amplitude (Fig. 6Db). We then analyzed these individual hotspot signals to determine variation within subregions of each hotspot and between individual AP-evoked hotspots.

We analyzed subregions within AP-evoked Ca²⁺ transients. Peak evoked Ca²⁺ hotspot transients from just one AP are shown for a section of axon (Fig. 7Aa) and the highlighted hotspot (Fig. 7Aa). The larger yellow ROI encompassing the whole hotspot was imaged during sequential bursts of 5 evoked APs (Fig. 7B, blue). Transients were isolated to just hotspots by removing the overall axon signal using the approach described above (Fig. 6Cb,Db). While the whole region showed variability in Ca²⁺ transient peak amplitudes, variations between stimuli were greater between subregions. This was assessed by measuring the amplitudes in a grid of ROIs, each of 5 × 5 pixels (0.5 × 0.5 µm) within the hotspot (Fig. 7Ab, small yellow ROIs). Two of these responses from different subregions during the same stimulus train are overlaid (Fig. 7C, top regions e and g) and peak amplitudes from all the subregions are expressed as a color-coded grid (Fig. 7C, bottom). Each region shows variability, but they do not co-vary in amplitude. Peak amplitudes of the two regions (from Fig. 7Ce,g) were also directly compared for each of 40 stimuli (Fig. 7D). There was no discernible correlation. This was true for all pairs of locations tested at this synapse and in a further four synapses with similar high signal-to-noise recording.

• Download figure
• Open in new tab
• Download powerpoint

Figure 7.

Variation in Ca²⁺ within hotspots. A, Ca²⁺ transient intensity was recorded following single APs in regions within the hotspot to demonstrate variation in responses between APs. Aa, Example of a single un-averaged response to one AP, expanded in Ab to outline a grid of analyzed regions (each 5 × 5 pixels – 0.5 × 0.5 µm) within a ROI encompassing the whole hotspot. B, Stimuli, three trains of five APs shows variations in amplitudes in the whole region (large yellow ROI in Ab) between individual stimuli. C, Variation within the smaller subregions is much greater. Two subregions are compared in the traces (top) and all subregions compared with color coding of peak amplitudes of ΔF/F for each AP in a train of 5. D, Peak values of responses expressed as ΔF/F from the two regions in C (e, g) were compared for each of 50 stimuli, showing no correlation between amplitudes from these two regions between stimuli. E, The degree of correlation between each subregion was plotted against distance between the centers of these subregions, showing no relationship. F, Correlation coefficients were also plotted between each subregion and all others in this AZ for 20 stimuli coding positive correlation of 1 as red, negative correlation of −1 as blue and zero correlation as black. Correlation between subregions was very low (mean correlation coefficient = −0.09). For this one hotspot, data are shown in the left of the figure. Mean correlations for similar analyses in AZs from five axons is shown on the right of the figure confirms this lack of correlation.

To determine whether these local rapid signals showed correlation in variation because of proximity, correlation coefficients between the peak values ΔF/F of Ca²⁺ transients each subregion and the other 8 measured at the AZ were compared by plotting these correlation coefficients against distance between ROI centers (Fig. 7E). Data were obtained from responses from 40 sequential APs. There was no relationship between correlation coefficient and distance between subregions visible in this plot. Results were similar for the other four synapses. To quantify this lack of correlation in all five recorded AZs, correlation coefficients of peak amplitudes between each subregion and the other eight were plotted as a color-coded grid (Fig. 7F). The data for the highlighted AZ are shown in the lower left triangle. Comparing each region with itself gave perfect correlation (red squares) but compared with other subregions correlations were randomly positive, absent, or negative (mean correlation coefficient = 0.03 ± 0.08; Fig. 7F). The mean values of correlation coefficients in all examined AZs were measured and the means displayed in the same way (Fig. 7F, upper right triangle; mean correlation coefficient from all subregions in all AZ = 0.008 ± 0.04, n = 5 AZs, not significantly different from zero, p = 0.83).

We also analyzed Ca²⁺ transients, encompassing the whole AZ, to determine their amplitude variation between APs. Ca²⁺ transients were measured in ROIs that encompassed the entire hotspot outlined in red in the first poststimulus image (Fig. 8A). Trains of stimuli were applied (five stimuli at 50 ms) and the resultant hotspot transients displayed as before but showing unaveraged responses (Fig. 8B). Three sequential stimulus train evoked transients are shown. In six AZs the amplitudes of these events varied between stimuli, and the mean event amplitude reduced throughout the stimulus train (Fig. 8C). To further analyze amplitude variation, amplitudes of all 320 stimuli were plotted as an amplitude histogram that was well fit by five Gaussians that included a peak representing just signal noise (Fig. 8Da). Gaussian fits were constrained such that the amplitude ratios of sequential gaussians were integers and the widths scaled with these amplitudes (not including failures). Gaussian peaks were observed at increments of 0.31 ΔF/F (Fig. 8C, purple). These Gaussian peaks may represent quantal amplitudes following individual channel openings because their numbers correspond well to the numbers of synchronous channel openings seen in cell-attached recordings and calculated from molar Ca²⁺ entry recordings. The amplitude reduction during trains was caused by a reduction in event probability. Amplitude histograms from each stimulus position in the train were plotted and again fit with multi-Gaussians. Similar peak increment values were observed but at lower increments at sequential stimuli (range from 0.33 to 0.29 ΔF/F; Fig. 8Db). For all six axons tested, if each Gaussian corresponds to a channel event, then a mean number of 1.73 ± 007 channels open at each AZ with a minimum of 0 and maximum of 7. Histograms, normalized to a single quantal amplitude of 1, from these six puncta in six axons were averaged to demonstrate the most common amplitude was of 1 quantum (Fig. 8E). Failures were not because of the absence of AP firing, because events were observed at other locations in the same axon on the same stimuli. Results are consistent with small numbers of channel openings with approximately discrete amplitudes detected for each AP and a reduction of opening probability but no change in event sizes at later stimuli in the train.

• Download figure
• Open in new tab
• Download powerpoint

Figure 8.

Variation in Ca²⁺ transients between APs. A, Single AZs were imaged (up to 320 APs in 80 trains) with no bleaching. Images expressed as ΔF/F are shown before stimuli, at the peak of responses to stimuli one to five and after stimulation. B, At individual whole AZs, responses were measured (5 stimuli, 3 repeats). Events are expressed at ΔF/F after whole axon diffuse Ca²⁺ signals were subtracted (see Fig. 6Cb,Db). C, Mean amplitudes of responses to these five stimuli show a reduction throughout the stimulus train. Amplitudes normalized to the peak of the response to stimulus 1. D, Peak of transients from 320 stimuli (80 × trains of 5) were plotted as an amplitude histogram for all 320 APs. These show distinct peaks implying events are multiples of a unit amplitude. Gaussian fits (magenta) were made to this distribution with two constraints; amplitudes between peaks were equal, and variance scaled linearly with increase in amplitude. Db, A similar set of histograms were calculated for each of the first through the fifth stimulus in each train and gaussians were again fit with the same constraints as Ca. While the numbers of low-amplitude responses increased later in the train the unit amplitudes (purple text) remained constant. E, The amplitude data from six analyzed AZs were normalized to the value of the unit amplitude from individual fits. The mean number of events per stimulus were then plotted against these normalized amplitudes giving a similar overall distribution of events.

Numbers of presynaptic Ca²⁺ channels and primed vesicles are the same

The lamprey giant axon provides unique direct access to recording from AZs using sharp electrodes while recording from their postsynaptic targets. This enables both paired-cell recording of direct monosynaptic connection between axons and their individual targets and simultaneous manipulation of the presynaptic biochemistry (Blackmer et al., 2001). We used this unique access to determine the numbers of primed vesicles at each AZ and their release probability (P_r). Thus, we could relate prime vesicle number and number of evoked vesicles to Ca²⁺ channel opening. Paired recordings can be readily made between individual giant axons and their target ventral horn neurons. Each such reticulospinal axon makes a number of connections (up to ∼12 synapses) onto individual target neurons in the ventral horn. We thus made paired recordings by recording reticulospinal axons with sharp microelectrodes and spinal ventral horn neurons using whole-cell patch clamp (Fig. 9A). EPSCs were recorded following evoked presynaptic APs (Fig. 9Ba). From repeated evoked EPSCs, peak evoked amplitude histograms yielded multiple events consistent with multiple AZs between the axons and target neurons (Fig. 9Bb). In this example paired recording, Gaussian fits to this histogram revealed a minimum evoked EPSC amplitude of 8.7 pA (Fig. 9Bb). In five example pairs, similar histograms were created and fitted with similar multiple Gaussian curves. The mean minimum quantal amplitude was 8.1 ± 2.7 pA (Fig. 9Bc, n = 5). Mean probability of release [P_r] was calculated from the Gaussian fits (Fig. 9Bb,Bc) to be 0.22 ± 0.02. P_r was relatively uniform at these synapses and ranged from 0.16 to 0.25 (see Table 7 for summary).

• Download figure
• Open in new tab
• Download powerpoint

Figure 9.

The number of primed vesicles at reticulospinal synaptic terminals. A, Synaptic responses were recorded in paired cell recordings between microelectrode recorded presynaptic axons and whole-cell patch recorded postsynaptic ventral horn neurons. B, Postsynaptic EPSCs (Ba, top) were recorded in voltage clamp following presynaptic APs evoked by 2-ms depolarizing current pulses applied through the presynaptic recording electrode (Ba, bottom). Individual synaptic responses were recorded repetitively at 30-s intervals shown from 30 stimuli. From 200 consecutive EPSCS amplitude histograms were constructed (Bb, example from 1 paired recording) and fitted with multiple Gaussian curves (purple line). Examples of fits from all six recordings are shown (Bc), with the amplitude between response failures and single quantal amplitudes normalized to a value of 1 for each curve (mean quantal amplitude = 8.1 ± 1.1 pA). C, Quantal amplitudes were confirmed by measuring mean amplitudes of spontaneous EPSCs whose frequency was enhanced by presynaptic injection of Ca²⁺ buffered to 200 μm with EGTA. Paired cell recordings were made between postsynaptic ventral horn neurons and presynaptic reticulospinal axons (Ca). Responses shown are means following 10 presynaptic APs. An example of control spontaneous activity recorded in ventral horn neurons is shown (Cb, black). In four paired cell recordings, the presynaptic electrode contained 100 mm EGTA and 100 mm CaCl₂ (free Ca²⁺ in electrode ∼ 100 μm, pH 7.2, 5 mm HEPES, and 1 m KCl). This solution was pressure injected into the presynaptic axon leading to much enhanced spontaneous release with no depolarization of the axon (Cb, red). This increase in frequency is quantified (Cc) in which control (black) cumulative interevent intervals (mean of four recordings) are compared with cumulative intervals after presynaptic Ca²⁺ injection (red, mean 4 paired recordings). Pink and gray lines indicate SEM. Cd, Spontaneous event amplitude histograms were quantified from these four paired (modal amplitude = 6.2 ± 0.6 pA). D, Paired recordings were made to determine the number of APs necessary to deplete the primed vesicle pool. Da, BoNT/B was injected into the presynaptic axon. After 5 min to allow toxin to act, stimuli (1 Hz) were applied and the EPSC measured until it decayed to zero (examples in inset). An exponential fit (magenta) to this gives a decay (τ) of 93 stimuli. Db, Means of similar responses from four paired cell recordings are also plotted. Mean values of τ from all pairs were taken to give the total number of APs (100 ± 17) to deplete the primed vesicle pool. E, Schematic model of the terminal. A small number of primed vesicles are associated with an equal and equally small number of Ca²⁺ channels available at the presynaptic terminal. Of these channels, only one to seven open in one AZ on each AP. This provides part of the reason for low probability of release.

We confirmed the amplitude of unitary events by recording asynchronous events between paired reticulospinal axons and their target neurons. We cannot relate raw spontaneous activity to spontaneous event amplitudes from reticulospinal axons because there are too many nonreticulospinal synapses onto the postsynaptic cell. However, asynchronous release was enhanced from individual reticulospinal presynaptic axons by injecting high Ca²⁺ concentrations buffered with EGTA to cause a tenfold increase in total spontaneous event frequency recorded in target neurons paired to these axons. To achieve this, paired recordings were again made with microelectrodes in the axon and whole-cell recording of the ventral horn target neurons. The presynaptic microelectrode contained ∼200 μm free Ca²⁺ at pH 7.2 in the presynaptic microelectrode. Evoked responses obtained by stimulating the axon immediately after obtaining the paired recording confirmed synaptic connectivity (Fig. 9Ca). Subsequent pressure injection (140 kPa, 200 ms repeated up to 20 times) caused a marked increase in the frequency of evoked events (Fig. 9Cb,Cc). This approach yielded a modal amplitude of these asynchronous events (6.6 ± 1.5 pA, n = 4), not significantly different from the minimal evoked amplitude obtained from histograms (Fig. 9B; p = 0.58), supporting the hypothesis that minimal evoked amplitudes represent unitary quantal events.

To understand the relationship between total numbers of VGCCs at AZs and numbers of primed vesicles we then calculated the number of APs required to deplete the primed vesicle pool by preventing vesicle priming with botulinum toxin B (BoNT/B). BoNT/B prevents neurotransmission by selectively cleaving synaptobrevin but cannot cleave targets after formation of ternary SNARE complexes during priming (Pellegrini et al., 1994). Thus, after BoNT/B injection into presynaptic axons the stimulus dependent rate of loss of axonal response represents the exhaustion of the BoNT/B resistant primed vesicle pool. In paired recordings, again made between microelectrode recorded reticulospinal axons and whole-cell recorded targets, axons were pressure injected with light chain BoNT/B (Gerachshenko et al., 2005) in the microelectrode filling solution. After BoNt/B injection, and after 5 min to allow cleavage of unprimed synaptobrevin (Gerachshenko et al., 2005), APs were repeatedly stimulated to evoke EPSCs (Fig. 9Da). The time course (τ) of EPSC amplitude decay expressed as number of APs was 100 ± 17 (Fig. 9D). If we consider P_r (Fig. 9B), the mean number of primed vesicles per AZ is 22, which corresponds very closely to the mean number of Ca²⁺ channels at the AZ (23 channels).