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1 Department of Neurobiology, David Geffen School of Medicine at UCLA, University of California, Los Angeles, 650 Charles E. Young Drive South, Los Angeles, CA 90095, USA
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(Received 6 April 2004;
accepted after revision 10 August 2004;
first published online 12 August 2004)
Corresponding author F. E. Schweizer: Department of Neurobiology, David Geffen School of Medicine at UCLA, University of California, Los Angeles, 650 Charles E. Young Drive South, Los Angeles, CA 90095, USA. Email: felixs{at}ucla.edu
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Introduction |
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Ribbon synapses are characterized by the presence of a presynaptic, electron-dense structure that is either plate-like (synaptic ribbon, e.g. found in photoreceptors (Sjostrand, 1958)) or spherical in shape (synaptic body, e.g. present in auditory hair cells (Smith & Sjostrand, 1961) and fish lateral line hair cells (Hama, 1965)). Club-shaped synaptic bodies have been described in fish electroreceptors (Wachtel & Szamier, 1966) and intriguing changes in the shape of the synaptic body during development from spherical to plate-like have been observed in the mouse cochlea (Sobkowicz et al. 1982). Synaptic bodies are studded with small synaptic vesicles and lie in close proximity to the active zone, two features that led to the proposal of their function as conveyor belts, delivering vesicles to the docking sites prior to release (Bunt, 1971; Gleisner et al. 1973). The conveyor belt model is consistent with the hypothesis that docked vesicles form a pool that is released with short latency following onset of Ca2+ influx. Those vesicles associated with the synaptic body could constitute a second pool that is released with slower kinetics, limited by the time required for vesicle transport to the plasma membrane (Mennerick & Matthews, 1996; von Gersdorff et al. 1996; Neves & Lagnado, 1999).
We chose mechano-sensitive hair cells from the sacculus of leopard frogs for our studies since their morphological and physiological properties have been characterized in unsurpassed detail. Also, the lack of processes makes it possible to gain electrical access to the release sites and compare the physiological properties of fast exocytosis with the reported morphology. The 3-dimensional ultrastructure of the afferent synapse between saccular hair cells and primary afferents from the VIIIth nerve has been described (Lenzi et al. 1999, 2002). On average, there are 32 vesicles docked at each active zone, while 380 vesicles are tethered to the synaptic body. An additional ‘cloud’ of outlying, cytoplasmic vesicles that are neither tethered to the synaptic body nor the plasma membrane surrounds the active zone. The rise in intracellular [Ca2+] in response to a depolarization is sharply restricted to the hair cell active zone since voltage-gated Ca2+ channels responsible for synaptic signalling are found almost exclusively in this region (Roberts et al. 1990) and a highly mobile Ca2+ buffer (Edmonds et al. 2000) prevents the local increase of free Ca2+ ions from spreading to adjacent regions (Roberts, 1993, 1994; Issa & Hudspeth, 1996; Zenisek et al. 2003). Given these data, it has been proposed that three morphologically defined vesicle pools fulfil physiologically distinct functions: docked vesicles support phasic release, ribbon-associated vesicles support tonic release and the outlying, cytoplasmic vesicles adjacent to the ribbon replenish vesicles leaving the ribbon (see, for example, von Gersdorff & Matthews, 1997; Neves & Lagnado, 1999; Holt et al. 2004). Docked vesicles would thus be expected to fuse with fast kinetics and ribbon-associated vesicles, requiring translocation toward the membrane, would constitute a pool fusing with slower kinetics, limited by the speed of vesicle movement. The outlying pool might not be detectable as a kinetically distinct, third component if replenishment is not rate limiting. In this study, we tested whether the number of fast-fusing vesicles is limited to the number of morphologically docked vesicles. We defined fast fusion as fusion occurring within 20 ms, i.e. fusion prior to the time it would take a molecular motor to move a vesicle by one vesicle diameter (> 20 ms; see Discussion). This pragmatic definition should limit the ‘fast phase’ to vesicles docked at the plasma membrane. By measuring cell membrane capacitance to assay vesicle fusion with the plasma membrane we found two phases of exocytosis: a fast phase that is exhausted in 20 ms and a slower phase that continued for at least 5 s, consistent with the above model. Surprisingly, we find that the fast phase is supported by the fusion of about 9 times more vesicles than those previously observed as docked at the presynaptic membrane. Our results therefore indicate that fast exocytosis at this ribbon synapse (< 20 ms) is predominantly mediated by vesicles that are not docked at active zones at the time of stimulus onset.
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Methods |
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Normal Ca2+ Ringer solution contained (mM): 112 NaCl, 2 KCl, 1.8 CaCl2, 0.7 MgCl2, 3 D-glucose and 5 Hepes (pH with NaOH to 7.25). For low Ca2+ Ringer solution, MgCl2 was omitted and CaCl2 was lowered to 0.1 mM. The osmolality of the external solutions was adjusted to 220–225 mmol kg–1. Unless stated otherwise, all reagents were obtained from Fluka (Sigma-Aldrich Corp.). We used hair cells from the sacculus of small (5.5–7.5 mm in length) leopard frogs (Rana pipiens, wild-caught male and female; various suppliers). Frogs were chilled on ice and killed using the double pithing method according to protocols approved by the UCLA Institutional Animal Care and Use Committee. Sacculi were removed from the inner ear and incubated for 6 min in low Ca2+ Ringer solution (on ice) to which 1 mM MgCl2 and 1 mM EGTA had been added. Sacculi were then transferred to low Ca2+ Ringer solution containing subtilisin (Protease Type XXIV; Sigma) at 50 µg ml–1 for 12–15 min (at room temperature). Cells were dissociated onto concanavalin-A-coated glass coverslips by gently scraping the macular epithelium with an eyelash attached to the end of a handheld probe. Cells were visualized on an inverted microscope (Zeiss, Axiovert 25 or S100; Carl Zeiss Inc. Oberkochen, Germany) using a 32 x objective.
Electrophysiological recordings
Recordings were made at room temperature (
23°C) using the perforated patch configuration of the whole-cell voltage clamp method. Patch pipettes were made with borosilicate glass (TW150F-4; World Precision Instruments, Sarasota, FL, USA) and pulled to tip resistances of 1.5–3.5 M
measured in our standard recording solutions. To reduce stray capacitance, tips were coated with HIPEC R6101 (Dow Corning, Midland, MI, USA; pre-incubated for 6 h at 90°C). Approximately 0.5 mm of the pipette tip was front-filled with intracellular solution containing (mM): 114 caesium aspartate (L-aspartate from Sigma), 0.08 CaCl2, 2 MgCl2, 5 Hepes, 1 EGTA (pH 7.25, 220–225 mmol kg–1). The pipette was back-filled with intracellular solution to which nystatin (50 µg ml–1; stored as 1000 x stock in DMSO at 20°C; Sigma) had been added. Recordings were made with an Axopatch 200 A (Axon Instruments) patch clamp amplifier or an Optopatch (Cairn Research, Faversham, UK) using jCLAMP software (Scisoft Co., New Haven, CT, USA). Membrane voltages were corrected for a measured liquid junction potential error of –13 mV between the intracellular and extracellular solutions. Series resistance of 14.2 ± 3.2 M (mean ± S.E.M.; n = 49) was uncompensated. Unless indicated otherwise, data were filtered at 30 kHz and sampled at 100 kHz with further filtering imposed in software. Ca2+ currents shown in Figs 2 and 3 were leak subtracted using a P/5 protocol and, for each depolarization, the nominal voltage was corrected for the voltage error due to the series resistance. Peak currents at the nominal voltages were then obtained by cubic spline interpolation and the average I–V plot (Fig. 2B) was constructed by averaging the interpolated current values at voltages between –120 and +20 mV. I–V plots from individual cells were fitted with the equation:
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| (1) |
the voltage at which the half-maximal current is achieved, and k is the slope factor indicating the steepness of the voltage dependence. The average values from the fit were: gmax = 0.011 ± 0.001 nS, Vrev = 39.2 ± 1.5 mV, V = –41.9 ± 1.2 mV and k = 7.05 ± 0.14 mV. An I–V curve calculated from these average values is superimposed onto the data (dashed line in Fig. 2B). The conductance versus voltage plot (Fig. 2C) was obtained by measuring the chord conductance between the test voltage and the reversal potential for the current. The chord conductance was calculated at all voltages as the current divided by the driving force and normalized using gmax. The data were then averaged and fitted with a Boltzman distribution:
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Capacitance measurements were made using a dual sine-wave voltage (DSWV) stimulus protocol when the series resistance was stable and less than 20 M (Donnelly, 1994; Rohlicek & Schmid, 1994; Gillis, 1995). Cells were held at –75 mV, and a DSWV (r.m.s. amplitude: 6 mV; maximal negative and positive excursions: –81.75 mV and –63 mV, respectively) of 390.5 and 781 Hz was superimposed on the applied holding potential. Admittance analysis was then performed with jCLAMP software on the Fast Fourier Transform (FFT) of the current response every 2.56 ms. The analysis yielded estimates for membrane capacitance (Cm), series resistance (Rs), membrane resistance (Rm) and the holding current (I) during the course of the experiment (Santos-Sacchi et al. 1998) at every time point. In contrast to some other techniques for measuring capacitance (for a review see Gillis, 1995), this method employs the same algorithm throughout the recording, does not require cancellation of baseline Rs and Cm, and makes no assumptions about Rm staying constant during the recording (Santos-Sacchi, 2004). To validate the dual sine-wave method, we measured baseline capacitance in 10 cells using both the dual sine-wave approach and the RC cancellation circuitry of the Optopatch. The values obtained with both methods were indistinguishable (2-sine wave: 14.69 ± 0.83 pF; RC cancellation: 14.7 ± 0.74 pF; n = 10). To verify that large changes in capacitance can be accurately measured with this technique, we used a model cell (Axon Instruments; nominal values: Cm = 33 pF, Rs = 10 M, Rm = 500 M) and placed a capacitor of nominally 4.7 pF in series with Cm. Using the DSWV method we determined a baseline capacitance of 37.14 ± 0.02 pF. Rapid removal of the 4.7 pF capacitor elicited a drop in the measured capacitance of 4.64 ± 0.05 pF, in excellent agreement with its nominal value. Not surprisingly, this large change in capacitance could not be accurately measured using a single sine wave and the built-in lock-in capabilities of the Optopatch amplifier (measured Cm change: 0.48 ± 0.05 pF), since the phase angle of the measurement has to be readjusted when large changes in Cm occur (Bookman et al. 1991; Gillis, 1995). However, when the automatic track-in feature of the Optopatch was enabled (Johnson et al. 2002) an accurate measurement could be made (measured Cm change: 4.61 ± 0.07 pF). A similar conclusion was recently reached by Thoreson et al. (2004) who compared the DSWV method implemented in jCLAMP with the square wave method. Furthermore, our values for total membrane capacitance of frog saccular hair cells are also consistent with values published in the literature (e.g. 8–16 pF (Smotherman & Narins, 1998); 15 ± 3 pF (Lioudyno et al. 2000); 7.1–17.3 pF (Holt et al. 2001); 16 pF used for model (Catacuzzeno et al. 2003)). We conclude that the DSWV method allows for accurate and reliable detection of large capacitance changes and offers the advantage over single sine-wave approaches since Cm, Rs and Rm can be continuously monitored during the course of the experiment.
Additional analysis on the parameters was performed off-line with Igor Pro 4.0 (WaveMetrics, Inc., Lake Oswego, OR, USA) and Origin 6.0 (MicroCal, Northampton, MA, USA). In some experiments, hair cells were held at –85 mV, enabling us to use a DSWV of larger amplitude (r.m.s. amplitude: 13 mV; maximal negative and positive excursions: –99.8 mV and –59 mV, respectively) without activating voltage-dependent conductances, and thus improve the signal to noise ratio of our estimates. In paired pulse experiments (Fig. 6) we increased the temporal resolution of our parameter estimates to once every 0.64 ms using a higher frequency DSWV (1562.5 and 3125 Hz). The DSWV was turned off during depolarization to avoid interference with Ca2+ channel activation.
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Our interpretations depend, to some extent, on correctly estimating the number of fused vesicles. We used a value of 37 aF per vesicle based on the vesicle diameter of 39 nm measured from three-dimensional reconstructions of electron micrographs of frog saccular hair cells (Lenzi et al. 1999). While shrinkage of the preparation prior to EM analysis might have led to an underestimate of the vesicle diameter (Lenzi et al. 1999) a good correspondence between vesicle diameters determined from electron micrographs and by membrane capacitance measurements has recently been reported (Klyachko & Jackson, 2002). We have assumed the ‘canonical’ membrane capacitance of 1 µF cm–2, although recent measurements of the specific membrane capacitance for various cell types have indicated that the correct value may be lower (
0.8 µF cm–2) (Solsona et al. 1998; Gentet et al. 2000; Roth & Hausser, 2001). We are thus probably underestimating, rather than overestimating, the actual number of vesicles fusing.
Our interpretations also depend on a correct estimate of the number of active zones per frog saccular hair cell. This number has been determined by different investigators using diverse methods. Issa and Hudspeth, using the fluorescent dye fluo-3 to label synaptic bodies, estimated the number at 18 ± 4 (Issa & Hudspeth, 1994), a result recently confirmed using evanescent wave microscopy (Zenisek et al. 2003). Roberts and colleagues used the dual approach of focal recordings, to identify ion channel clusters, and electron microscopy to identify active zones. Their results suggested that hair cells contain 20 channel clusters and 19 active zones (Roberts et al. 1990). In turtle hair cells, a maximum of six active zones per cell have been reported (Tucker & Fettiplace, 1995). For our interpretations, we have taken the conservative and generally accepted value of 20 active zones per frog saccular hair cell.
Statistical methods
We utilized bootstrapping methodology to test the confidence of our conclusions from Fig. 4 by simulating the null hypothesis without making any assumptions about the statistical distribution of the data (Efron & Tibshirani, 1991). For a given duration of depolarization and stimulation strength, data were placed into a vector in Resampling Stats (http://www.resample.com). These data were randomly sampled N times with replacement (N is the total number of data points), a mean was calculated and this was repeated 15 000 times, allowing the construction of a frequency histogram of the re-sampled means and the estimation of statistical significance (P < 0.05). For other data a standard t test was performed as the test for statistical significance. For averaged data, values are given as means ± S.E.M.
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Results |
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We measured exocytosis as a change in cell surface area by monitoring cell membrane capacitance (Neher & Marty, 1982). To avoid wash-out of endogenous Ca2+ buffers (Edmonds et al. 2000; Moser & Beutner, 2000) we used the perforated-patch configuration of the patch clamp technique for all experiments. To avoid confounding effects of repetitive stimulation, only responses to the very first depolarization (or pairs of depolarization) for each cell were included in the analysis. The cell membrane capacitance (Cm) of unstimulated hair cells was between 8 and 22 pF (14.9 ± 3.3 pF, n = 49; see Methods) and remained stable unless the cell was stimulated by depolarization. Figure 1A shows a typical example of a recording from a hair cell with an initial cell membrane capacitance of 14 pF. Depolarization from the holding potential of –75 mV to –20 mV for 25 ms activated an inward current (Fig. 1A; I), mostly carried by Ca2+ ions since K+ conductances were blocked by internal caesium ions and these cells do not express voltage-dependent Na+ conductances (Lewis & Hudspeth, 1983). Upon repolarization, the membrane capacitance increased by 400 fF (Fig. 1A; Cm). Using an average capacitance of 37 aF per vesicle (Lenzi et al.), the observed increase corresponds to the release of 540 vesicles at each of the 20 active zones (Roberts et al. 1990; Issa & Hudspeth, 1994; Zenisek et al. 2003).
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As indicated in Fig. 1A, the series resistance (Rs) was stable during the experiment; however, the input resistance (Rm) was higher immediately after repolarization and quickly relaxed back to baseline. The transient increase in resistance, which is paralleled by an outward current (I), probably reflects voltage-dependent closing of inward rectifier K+ channels as it was mostly blocked by the addition of 1 mM CsCl to the external solution (Fig. 1B) (Holt & Eatock, 1995). Most experiments were performed without external caesium and since external caesium did not appear to alter capacitance changes (20 ms depolarization in Cs+: 301 ± 125 fF, n = 5; 25 ms depolarization without Cs+: 248 ± 67 fF, n = 12; P = 0.7), data with and without caesium were pooled.
Kinetics of Ca2+ current
The initial rate of exocytosis following depolarization depends on a number of factors including the amplitude and the rate of activation of the Ca2+ current (ICa). Therefore, we examined the voltage dependence of the amplitude and rise time of ICa. In agreement with previous reports (Armstrong & Roberts, 1998; Rodriguez-Contreras & Yamoah, 2001), the Ca2+ current in hair cells showed little inactivation (Fig. 2A) and began to activate with weak depolarizations to around –65 mV with a half-maximal current at –50 mV and a peak inward current at –20 mV (Fig. 2B). We also estimated channel activation by calculating the conductance at all test voltages (see Methods). Again, full activation was achieved at –20 mV whereas at –50 mV only 25% of the maximal conductance had been activated (Fig. 2C). Figure 3A shows two recordings of ICa in response to depolarizing steps to –50 and –20 mV. The rising phase of each of these currents was well fitted by a third order exponential:
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is the maximum value for ICa and 
is the time constant associated with the movement of 1 of 3 hypothetical gating charges within the Ca2+ channel (Fig. 3A and B) (Hudspeth & Lewis, 1988). Mean values for were 687 ± 156 µs (n = 7) and 316 ± 47 µs (n = 9) for jumps to –50 and –20 mV, respectively. The average 10–90% rise times were 1.5 ms and 0.7 ms for steps to –50 and –20 mV, respectively. Activation of ICa was very fast at all potentials tested (Fig. 3B) and thus appears too fast to significantly contribute to the time course of exocytosis (see below). Depolarizations to –50 mV and to –20 mV were used in subsequent experiments since they elicit fast activation of sustained ICa with different amplitudes. Fast phase of exocytosis
To determine the secretory capacity of hair cells, we depolarized the voltage clamped cells to –20 mV for a range of durations. Figure 4A shows examples of the observed capacitance changes. The magnitude of the capacitance change increases with longer depolarizations; however, longer depolarizations trigger less release than might be expected from a linear relationship suggesting saturation of the fast exocytotic process. A plot of the mean capacitance increase for stimuli in the range of 2–30 ms (Fig. 4B, filled triangles; n = 3–13) indeed demonstrates a fast but limited phase of exocytosis. A 20 ms depolarization to –20 mV elicited an average capacitance increase of 253 ± 78 fF (n = 8; range 109–783 fF; 4 ms: mean = 54 ± 19 fF, n = 8, range 2–124.5 fF; 10 ms: mean: 226 ± 87 fF, n = 9, range 5.7–844 fF). These data were well described by a ‘fast’ exponential of order 4.8 that saturates at 209 fF with a time constant of 2.7 ms (Fig. 4B).
This amplitude corresponds to the fusion of 282 vesicles at each of the 20 active zones (Roberts et al. 1990; Issa & Hudspeth, 1994; Zenisek et al. 2003) about an order of magnitude more than the 32 vesicles that are docked (Lenzi et al. 1999). Even assuming dense hexagonal packing of docked presynaptic vesicles, the active zone could accommodate only 80 vesicles corresponding to 60 fF of potential capacitance increase, at least 3 times less than we observed. Therefore, the amplitude of the fast phase that we detect cannot be accounted for by the fusion of vesicles pre-docked at active zones.
At the ribbon synapse in bipolar cell terminals it has been suggested that the number of synaptic vesicles that fuse with short latency depends on stimulus strength, that is, on the amplitude of the Ca2+ current (von Gersdorff & Matthews, 1997). Therefore, in addition to eliciting maximal Ca2+ currents by stepping to –20 mV, we examined capacitance changes resulting from weaker stimuli. Depolarization to –50 mV recruited only half of the maximal Ca2+ current (Fig. 2B), due to the opening of fewer Ca2+ channels (75% fewer on average than at –20 mV, Fig. 2C). A 20 ms depolarization to –50 mV elicited an average capacitance increase of 220 ± 31 fF (n = 7; range 138–348 fF), not significantly different (P < 0.001) from the increase at –20 mV (4 ms: mean 10 ± 6 fF, n = 3, range 0–19 fF; 10 ms: mean 93 ± 21 fF, n = 3, range 58–132 fF). Similarly, the amplitude of the fast component for depolarization to –50 mV obtained from fitting (213 fF) was virtually identical to the amplitude of the fast component at –20 mV (209 fF) (Fig. 4B, dotted lines). Thus, either stimulus recruited about 9 times more vesicles for fast release than are identified morphologically as docked. In contrast to the correspondence in amplitudes, the rates of release were significantly different: the fast exponential component of release at –50 mV had a time constant of 5.5 ms, about twice the value at –20 mV (2.7 ms). To estimate the maximal release rates at –50 and –20 mV we differentiated the curves fitted to the change in Cm as a function of duration of the stimulus (Fig. 5). The maximal release rate at –20 mV was 32 fF ms–1, corresponding to a peak vesicle fusion rate of 43 vesicles ms–1 at each active zone. The values at –50 mV were 16 fF ms–1 and 22 vesicles ms–1, respectively (Fig. 5). In summary, while the rate of exocytosis does depend on stimulus strength, the size of the limited, fast pool does not. This pool of fast-fusing vesicles greatly exceeds the reported number of vesicles docked at actives zones (Lenzi et al. 1999).
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Longer depolarizations to –20 mV of up to 5 s in duration trigger an additional phase of exocytosis that appeared to continue at a constant rate of 1.1 pF s–1, corresponding to the fusion of about 1500 vesicles s–1 at each active zone (Fig. 4C, filled triangles; average increase at 1 s: 1.7 ± 0.5 pF; n = 8; range 0.8–4.1 pF). Depolarizations to –50 mV also yielded a slow component of release (Fig. 4C; filled circles; average increase at 1 s: 326 ± 16 fF; n = 3; range 301–357 fF), proceeding at a slower rate of 0.14 pF s–1, corresponding to the fusion of 190 vesicles s–1 at each active zone. Therefore, the rate of exocytosis from the slow pool also depends on stimulus strength.
Paired pulse plasticity
As indicated in Fig. 4B, a 10 ms depolarization to –20 mV elicits the release of approximately 90% of the fast pool of vesicles. Under physiological conditions these cells respond to stimulation in the range of 20–200 Hz (Koyama et al. 1982; Hudspeth & Lewis, 1988; Lewis, 1988), corresponding to bouts of exocytosis every 5–50 ms. We therefore tested the efficacy of a second 10 ms stimulus delivered after a 10 ms interstimulus interval (ISI). We observed a marked depression of the second response relative to the first for a 10 ms ISI (64% depression; Fig. 6). Recovery from paired pulse depression was then investigated by increasing the ISI from 10 to 100 ms, and the data were fitted with a single exponential with a time constant of 29 ms. The observed recovery time course of the fast component of release is consistent with the functional requirement that hair cells produce bouts of exocytosis at frequencies to which they are maximally sensitive (Glowatzki & Fuchs, 2002). Interestingly, longer depolarizations of 100 ms elicited much more long-lasting depression. For example, with an ISI of 100 ms, the amplitude of the response to the second stimulus was only 54% of the initial response (Fig. 6, filled triangle; 71% recovery for 200 ms ISI (data not shown)). This longer lasting depression is consistent with adaptation to a prolonged stimulus (Furukawa & Matsuura, 1978).
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Discussion |
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A previous study in frog saccular hair cells (Parsons et al. 1994) reported a slow phase of exocytosis but failed to detect a fast phase. While the nature of this discrepancy is unclear, a relatively large amplitude sine wave (40 mV peak to peak, Vh: –70 mV: maximal positive excursion –50 mV, versus maximal positive excursion –63 mV in this study; see Methods; both studies report similar series resistance values) might have triggered the release of the fast pool before data acquisition was initiated. Such depression of the readily releasable pool of vesicles by prolonged depolarization has recently been reported for chick cochlear hair cells (Eisen et al. 2004), but other experimental differences might also contribute.
In agreement with data from frog sacculus (Parsons et al. 1994), mouse (Moser & Beutner, 2000) and chick cochlear hair cells (Eisen et al. 2004) we did not observe a fast component of endocytosis following depolarization-induced exocytosis. While a fast component of endocytosis was observed in response to flash photolysis of caged calcium that raised the global intracellular free calcium concentration above 15 µM, no fast endocytosis was observed at lower free calcium concentrations (Beutner et al. 2001). This might indicate that calcium is elevated only locally in response to depolarization (see below) and that calcium does not rise above 15 µM at endocytic sites. However, since capacitance measurements detect net changes in surface area, we cannot exclude the existence of endocytosis during the depolarization. In such a case, our measurements would represent an underestimate of the number of vesicles fusing, especially during the slow phase but conceivably also during the fast phase of exocytosis.
Fusion of vesicles outside of the active zone
An alternative hypothesis that could account for the near 10-fold discrepancy between the number of fast-fusing vesicles and vesicles docked at active zones is that ‘docked’ extrasynaptic vesicles are also fusing during the stimulus. Several observations suggest that this is not the case. First, voltage-gated Ca2+ channels, responsible for synaptic signalling, are almost exclusively located at the active zone (Roberts et al. 1990; Issa & Hudspeth, 1994; but see Rodriguez-Contreras & Yamoah, 2001) and Ca2+ entry hot spots have been detected and determined to colocalize with active zones (Issa & Hudspeth, 1994; Zenisek et al. 2003). Second, Ca2+ ‘spillover’ from the active zone to vesicles docked outside of the active zone is not likely to raise the Ca2+ concentration high enough to support vesicle fusion due to the strong endogenous calcium buffering capacity of hair cells (Roberts, 1993, 1994; Edmonds et al. 2000; Schneggenburger & Neher, 2000). Third, if the 5000 ‘excess fusing vesicles’ not docked at active zones (300 total fusing vesicles minus 32 docked at each of 20 active zones) were each tightly associated with at least one calcium channel with an amplitude of 0.5 pA (Rodriguez-Contreras & Yamoah, 2003) to support vesicle fusion, then the extrasynaptic calcium current alone would have to be considerably larger (> 2.5 nA) than the measured current (600 pA). Fourth, at –50 mV, the open probability of single Ca2+ channels is significantly smaller than at –20 mV (Rodriguez-Contreras & Yamoah, 2003). Indeed, only about 25% of the channels activated at –20 mV are activated at –50 mV (Fig. 2C). Therefore, if extrasynaptic vesicles and thus single channels were significantly contributing to exocytosis, then the amplitude of the fast phase at –50 mV should be smaller than at –20 mV, which is in direct contrast to our experimental observations (Fig. 3B). Fifth, extrasynaptic fusion events would have to outnumber synaptic events almost 10: 1, drastically reducing or even eliminating the specificity of synaptic transmission. Sixth, the fast pool is refilled very rapidly (Fig. 6), which would be unlikely if it were composed primarily of outlying vesicles. In addition, direct observation of fusion events has indicated the existence of fusion hot-spots (Zenisek et al. 2000, 2003; Holt et al. 2004). Furthermore, active transport is too slow to allow for a significant number of outlying docked vesicles to be recruited into the active zone during the fast phase of exocytosis (see below). Such a recruitment could be responsible for the general depletion of vesicles seen after 30 min of stimulation (Lenzi et al. 2002), much longer than the 20 ms considered here. It is worth noting that at the frog neuromuscular junction, fusion at the active zone was elicited with stimuli of short duration (‘0 min’) while extrasynaptic fusion events were only observed if the stimulation lasted for more than 1 min (Ceccarelli et al. 1988). Thus, while we can certainly not exclude that a few outlying vesicles contribute to our signal, it is exceedingly unlikely that fusion of vesicles occurs predominantly outside of active zones.
The near 10-fold difference between the number of morphologically docked and fast-fusing vesicles can also not be explained by an undercount of docked vesicles. Assuming hexagonal tight packing, only 80 vesicles, approximately one third of the fast-fusing vesicles, could be docked in the area where free Ca2+ can reach concentrations high enough to support fusion (active zone with radius of 138 nm plus a rim of 40 nm (Roberts et al. 1990; Lenzi et al. 1999)). Taken together, our data indicate that at least three to nine times more vesicles than those docked at the active zone can fuse with fast kinetics, and points to the unconventional hypothesis that a non-docked source of vesicles contributes to the fast phase of release.
Capacitance increases that are larger than expected have also been elicited in cochlear hair cells, but in response to flash photolysis of caged calcium (Beutner et al. 2001). However, this large pool could not be depleted by prior depolarization and the authors argue that fusion of vesicles localized outside of the active zone and not associated with calcium channels contribute a majority of the vesicles fusing in response to flash photolysis. Flash photolysis raises Ca2+ concentrations at active zones with a spatio-temporal profile that is distinct from that induced by depolarization. In addition, Ca2+ concentration is increased at sites away from Ca2+ channels and active zones, which may account for an abundance of extrasynaptic fusion (Beutner et al. 2001). It will therefore be interesting to test whether uncaging of calcium in frog saccular hair cells could release an even larger number of vesicles, namely outlying membrane-bound vesicles in addition to the large depolarization-dependent pool observed here.
Fast-fusing vesicles and compound fusion
Morphological data demonstrate the existence of a small pool of docked vesicles at the active zone of hair cells. Our data are consistent with the idea that these vesicles make up an ‘ultrafast’ pool of vesicles (Mennerick & Matthews, 1996) that we did not resolve. Indeed, based on resampling statistics, more vesicles fused in less than 2 ms than the number docked, and more vesicles fused in less than 4 ms than the maximum number theoretically packed at the active zone. This emphasizes the major finding reported here, namely that a large excess of vesicles fuse during a time interval too short to allow for vesicle movement or for diffusion of calcium to distant sites.
What then are the anatomical correlates of these fast-fusing vesicles? Electron micrographs demonstrate the existence of a pool of about 380 vesicles that are bound to the synaptic body in frog saccular hair cells (Lenzi et al. 1999). Perhaps the synaptic body-associated vesicles contribute to the fast phase of release, with the synaptic body acting as a conveyor belt to supply vesicles to the release sites (von Gersdorff, 2001). However, molecular motor proteins such as the kinesin KIF3A that is associated with synaptic ribbons of photoreceptors (Muresan et al. 1999) have transport speeds of only 0.05–2 µm s–1 (Hirokawa, 1998). During the 20 ms of the fast phase, such a motor can transport vesicles at most a mere 40 nm, i.e. only about one vesicle diameter. Given a diameter of the synaptic body of 450 nm (Lenzi et al. 1999), those vesicles located on the synaptic body farthest from the active zone would have to move at rates of more than 22 µm s–1 or 10 times faster than the fastest known kinesin (Hirokawa, 1998). In fact, actual movement of synaptic vesicles has been estimated to be much slower (50 nm s–1 (Rizzoli & Betz, 2004); 0.5 µm s–1 (Klopfenstein & Vale, 2004); 0.75 µm s–1 (Rea et al. 2004)). For this reason we consider it unlikely that the ribbon actively delivers vesicles to the presynaptic membrane for fusion during the fast phase of exocytosis. The ribbon more probably serves as a conveyor belt during the slower sustained phase of release and/or to maintain the structural integrity of the synapse during massive membrane turnover (Bunt, 1971; Lenzi et al. 2002; Parsons & Sterling, 2003; Holt et al. 2004).
Since there are not enough vesicles within 40 nm of the plasma membrane to account for the amplitude of the fast phase, we instead propose that vesicles further away from the membrane fuse with vesicles closer to or even docked to the presynaptic membrane, a process known as compound fusion (Palade, 1975). Such compound fusion was previously suggested as a mechanism for release at ribbon-type synapses (Heidelberger et al. 1994; Heidelberger, 1998; Parsons & Sterling, 2003) and is consistent with the large invaginations occasionally observed at release sites of frog saccular hair cells (Lenzi et al. 1999, 2002). Such invaginations are also compatible with a lack of visible ‘inflation’ of the hair cells even in response to massive surface membrane increase (Fig. 4C). Compound fusion can also account for the large spontaneous postsynaptic events recorded in single afferent fibres in the frog vestibular system (Rossi et al. 1977) and for the very large postsynaptic events in recordings from single boutons contacting cochlear hair cells whose frequency, but not amplitude, changed during depolarization of the hair cells (Glowatzki & Fuchs, 2002). Such large events could drive the postsynaptic membrane quickly to threshold in order to maintain precise timing information (Glowatzki & Fuchs, 2002) and our experiments support the proposal (Parsons & Sterling, 2003) that they might be due to compound fusion.
Given the tight spatio-temporal calcium buffering in hair cells (Roberts, 1993, 1994), what could trigger the fusion of more distant vesicles in a compound exocytosis scheme? We propose three entirely speculative possibilities. (1) While calcium cannot diffuse very far in unconstrained space (Roberts, 1994), the space at the active zone of hair cells is rather restricted by the presence of vesicles and the synaptic body. This might allow for a larger and faster calcium increase close to the synaptic body, while still restricting lateral diffusion of calcium. (2) The synaptic body could directly be involved in signal transduction and might serve as a ‘calcium wire’ to deliver the stimulus deep into the terminal (a calcium ion absorbed at one end could ‘push’ another calcium ion out at the other end). (3) The cisternae formed by the fusing vesicles might serve as an additional source for a fusion stimulus away from the plasma membrane.
The fast-fusing pool of vesicles is depleted within 20 ms in response to both strong and weak stimuli (Fig. 4B). Consistent with a depletion of the fast pool, we also find strong paired pulse depression at this synapse (Fig. 6). Recovery from paired pulse depression is remarkably fast with a time constant of 29 ms. This is a much faster rate of recovery than has been reported for the ribbon-containing retinal bipolar cells (Mennerick & Matthews, 1996), and faster than was reported in cochlear hair cells (Moser & Beutner, 2000). Fast recovery might allow the synapse to release sufficient amounts of transmitter to trigger postsynaptic action potentials even in response to repetitive stimuli that mimic behaviourally relevant stimuli more closely than single step depolarizations (Koyama et al. 1982; Hudspeth & Lewis, 1988; Lewis, 1988).
Our experiments suggest that vesicles not initially docked to the plasma membrane can fuse with fast kinetics. Thus, in contrast to current models, stable docking does not appear to be a strict requirement for fast neurotransmitter release at graded synapses. It remains to be elucidated whether vesicles not docked at active zones can fuse with fast kinetics at other types of synapses or whether the ribbon plays a unique role in the fusion of these vesicles.
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(dashed lines; 
Cm) are plotted against the time of depolarization to –20 mV (
) and to –50 mV (•). The data for depolarizations up to 500 ms were fitted (dashed lines) with the equation: 
, where A, 
