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Myosin V functions as a vesicle tether at the plasma membrane to control neurotransmitter release in central synapses

  1. Dario Maschi
  2. Michael W Gramlich
  3. Vitaly A Klyachko  Is a corresponding author
  1. Washington University, United States
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Cite this article as: eLife 2018;7:e39440 doi: 10.7554/eLife.39440

Abstract

Synaptic vesicle fusion occurs at specialized release sites at the active zone. How refilling of release sites with new vesicles is regulated in central synapses remains poorly understood. Using nanoscale-resolution detection of individual release events in rat hippocampal synapses we found that inhibition of myosin V, the predominant vesicle-associated motor, strongly reduced refilling of the release sites during repetitive stimulation. Single-vesicle tracking revealed that recycling vesicles continuously shuttle between a plasma membrane pool and an inner pool. Vesicle retention at the membrane pool was regulated by neural activity in a myosin V dependent manner. Ultrastructural measurements of vesicle occupancy at the plasma membrane together with analyses of single-vesicle trajectories during vesicle shuttling between the pools suggest that myosin V acts as a vesicle tether at the plasma membrane, rather than a motor transporting vesicles to the release sites, or directly regulating vesicle exocytosis.

https://doi.org/10.7554/eLife.39440.001

Introduction

Quantal vesicle release at the synaptic active zone (AZ) represents a unitary event of information transmission at synapses. Recent nanoscale resolution studies revealed that vesicle release is organized in multiple discreet release sites, which are distributed throughout the AZ and spatially coincide with the clusters of presynaptic docking factors (Maschi and Klyachko, 2017; Tang et al., 2016). While the spatial organization and molecular architecture of release sites are beginning to emerge, the mechanisms that dynamically regulate the release site properties and reuse remain poorly understood.

Following a fusion event, refilling of a release site with a new vesicle is believed to represent a critical rate-limiting step in the release site reuse capacity thereby governing the ability of synapses to sustain release during repetitive activity (Neher, 2010). How this process is organized and regulated at the AZ is largely unknown due to the difficulty of directly visualizing the refilling process at the AZ, whosedimensions are typically at or below the diffraction limited resolution of conventional microscopy.

Presynaptic terminals are rich in actin filaments (Cingolani and Goda, 2008) and recent studies suggested that refilling of release sites in cerebellar synapses is actin-dependent (Miki et al., 2016). The major unanswered question is what role actin cytoskeleton and actin-based vesicle transport play in release site refilling: an active, structural, or both? An active, actin-dependent role occurs via molecular motors, for which only myosin V has been identified as a presynaptic vesicle-associated protein in central neurons (Prekeris and Terrian, 1997; Takamori et al., 2006; Watanabe et al., 2005). Major synaptic defects were observed in cortical neurons of myosin V dominant-negative mutant mice flailer (Yoshii et al., 2013) and due to myosin V knockdown (Correia et al., 2008). Myosin V null mice exhibit severe seizures and human mutations in myosin V gene cause severe nervous system dysfunction known as Griscelli syndrome (Hammer and Wagner, 2013; Kneussel and Wagner, 2013; Pastural et al., 1997). These findings point to a major role of myosin V in synaptic function; yet whether or how myosin V regulates neurotransmitter release remains largely unexplored. Single-vesicle tracking experiments showed that a majority of recycling vesicles undergo large-scale motion within synaptic boutons (Forte et al., 2017; Gramlich and Klyachko, 2017; Kamin et al., 2010; Lee et al., 2012; Park et al., 2012; Peng et al., 2012; Westphal et al., 2008) including a substantial component of directed actin-dependent motion (Forte et al., 2017; Gramlich and Klyachko, 2017; Peng et al., 2012). Myosin V may thus regulate release by supporting vesicle transport to the release sites. In addition to being a processive motor, myosin V is also known to function as a tether and to interact with the SNARE proteins in a Ca2+-dependent manner to promote the SNARE complex formation (Krementsov et al., 2004; Ohyama et al., 2001; Prekeris and Terrian, 1997; Watanabe et al., 2005), leading several studies to suggest a role for myosin V in vesicle docking and exocytosis (Desnos et al., 2007; Eichler et al., 2006; Porat-Shliom et al., 2013; Rudolf et al., 2011). Whether in central synapses myosin V plays a role in the vesicle transport to refill release sites, or at the later stages of the refilling process by tethering the vesicle to the release site machinery, or in the vesicle exocytosis itself is poorly understood.

Here, we sought to address these questions by employing a nanoscale detection of individual vesicle release events at the AZ in the hippocampal boutons, together with single-vesicle tracking to visualize release site refilling and reuse. Our results uncover a major role for myosin V in release site refilling, but not the exocytosis process itself. Surprisingly, rather than a unidirectional vesicle flow towards the release sites, we observed a dynamic vesicle shuttling between a plasma membrane pool and an inner pool, which is regulated by neuronal activity and requires myosin V as a vesicle tether rather than a transporting motor. These results, supported by ultrastructural analyses, suggest a major role for myosin V in regulating neurotransmitter release by controlling vesicle retention at the release sites rather than vesicle transport to the release sites or the exocytosis process itself.

Results

Inhibition of myosin V reduces release site re-use in hippocampal synapses

To understand the role of myosin V in presynaptic release mechanisms, we employed a nanoscale imaging modality to examine the effects of myosin V inhibition on spatiotemporal features of individual release events in hippocampal boutons. Our imaging approach takes advantage of a pH-sensitive indicator pHluorin targeted to the vesicle lumen via vGlut1 (vGlut1-pHluorin) (Balaji and Ryan, 2007; Leitz and Kavalali, 2011; Voglmaier et al., 2006) to permit detection of single vesicle release events with a 27 nm precision (Maschi and Klyachko, 2017). vGlut1-pHluorin was expressed in cultures of excitatory hippocampal neurons using a lentiviral infection at DIV3 and imaging was performed at DIV 16 – 19 at 37°C. Robust detection of individual release events evoked by 1 AP stimulation at 1 Hz was achieved within individual synapses at 40 ms/frame rate throughout the observation time period of 120 s (Figure 1A). Hierarchical clustering algorithms were used to define individual release sites within each bouton using a cluster diameter of 50 nm (Maschi and Klyachko, 2017).

Figure 1 with 1 supplement see all
Inhibition of myosin V impairs the reuse of release sites in hippocampal synapses.

(A) Sample spatial distribution of release events within a single hippocampal bouton evoked by 1 Hz stimulation. Scale bar: 1 µm. (B) Effect of myosin V inhibition with Myo1 or PBP on release site reuse was evaluated using a paired-pulse protocol as a probability that the same site is reused for two sequential stimuli 1000 ms apart, normalized to the same measurement in control condition. (C) Effect of myosin V inhibition with Myo1 on the probability that the same site is reused for two stimuli at different inter-stimulus intervals (in the range of 1 – 10 s) normalized to the same measurement in control conditions and binned for two sequential intervals. (D) Effect of myosin V inhibition on the average number of clusters/release sites detected in individual boutons plotted as a function of the number of release events observed (left). Pooled average number of clusters for boutons with 15 to 19 detected events in different conditions indicated (right). (E) Large-Area Scanning Electron Microscopy (LaSEM) of hippocampal boutons in culture showing that no significant changes in the AZ size were detected as a result of myosin V inhibition with Myo1. Examples of control and Myo1 treatment are shown (left). Scale bar: 200 nm. (F) Effect of myosin V inhibition on the distribution of distances of release events to AZ center evoked at 1 Hz. **=P < 0.01, *=P < 0.05, two-sample t-test; ns – not significant.

https://doi.org/10.7554/eLife.39440.002

To avoid potential developmental effects of interfering with myosin V function, we used the most acute approach to inhibit myosin V in developed neurons using two different and highly specific agents that arrest the myosin V ATP/ADP cycle (MyoVin-1 (Myo1) and Pentabromopseudilin (PBP) (Fedorov et al., 2009; Gramlich and Klyachko, 2017; Islam et al., 2010). Using this approach, we examined the effects of myosin V inhibition on vesicle release in basal conditions and during repetitive stimulation to distinguish two hypothesized functions of myosin V in synaptic transmission:

  • Myosin V may regulate refilling of the release sites with a new vesicle. In this case, myosin V inhibition would not strongly affect the basal release probability from a resting state when release sites are presumed to be largely occupied. In contrast, we expect to observe a significant effect of myosin V inhibition on vesicle release probability during repetitive activity. This effect should be more pronounced as stimulation frequency increases when demand for the new vesicles is higher.

  • Myosin V may function downstream of release site refilling, as a mediator of the exocytosis process itself. In this case, the requirement for myosin V should be apparent independently of the stimulation frequency. Thus, inhibition of myosin V is expected to reduce the vesicle release probability from a resting state as well as during repetitive activity, and the magnitude of the effect should be similar for a wide range of stimulus frequencies.

To discern these models, we first examined effect of myosin V inhibition on basal release probability, which we estimated based on a number of single release events evoked by 100 APs at 1 Hz. Inhibition of myosin V with either MyoVin-1 or PBP had no or only a small effect on basal release probability (Table 1), arguing against the major role for myosin V in the exocytosis process itself.

Table 1
Table of all data values and statistical analyses.

Data table columns are formatted as (i) corresponding figure location; (ii) conditions being statistically compared and separated by ‘/’; (iii) number of samples (synapses, dishes, cultures) used for each test; (iv) mean values and errors for each condition separated by ‘/’ and corresponding to conditions in column (i); (v) statistical test used for comparison; (vi) P-value resulting from the statistical comparison.

https://doi.org/10.7554/eLife.39440.004
Figure numberConditionsNSynNDishesNculturesmean ± semStastical testP val
Basal Pr
(related to 1)
Ctrl/Myo-1367/25915/154/30.064 ± 0.001/0.056 ± 0.002Two-sample t-test0.004
Ctrl/PBP367/83915/114/30.064 ± 0.001/0.067 ± 0.001Two-sample t-test0.15
1(B)Ctrl/Myo-1367/25915/154/31.000 ± 0.006/0.503 ± 0.004Two-sample t-test0.02
Ctrl/PBP367/83915/114/31.000 ± 0.006/0.5971 ± 0.003Two-sample t-test0.02
1(C)1–4/7–10367/25915/154/30.63 ± 0.06/1.036 ± 0.1028Two-sample t-test0.003
1(D)Ctrl/Myo-1367/25915/154/311.1 ± 0.2/10.9 ± 0.3Two-sample t-test0.58
Ctrl/PBP367/83915/114/311.1 ± 0.2/11.0 ± 0.1Two-sample t-test0.56
1(E)Ctrl/Myo-1137/813/33/3378 ± 10/364 ± 13Two-sample t-test0.38
1(F)Ctrl/Myo-1367/25915/154/3120 ± 3/129 ± 4Two-sample t-test0.04
Ctrl/PBP367/83915/114/3120 ± 3/132 ± 3Two-sample t-test0.01
2(B)Ctrl/Myo-1-9/103/30.50 ± 0.06Two-sample t-test<0.001
Ctrl/PBP-9/73/30.64 ± 0.04Two-sample t-test<0.001
Slope
(related to 2B)
Ctrl/Myo-1-9/103/30.52 ± 0.05Two-sample t-test<0.001
Ctrl/PBP-9/73/30.62 ± 0.04Two-sample t-test<0.001
2(C)Ctrl/Myo-1-36/3411/11y = 0.75269–0.17806 x (Linear Fit)Linear Fit<0.001
2(F)1 Hz/10 Hz, Ctrl367/25415/154/6120 ± 3/126 ± 2Two-sample t-test0.04
1 Hz/10 Hz, Myo-1259/86215/103/3129 ± 4/151 ± 6Two-sample t-test0.004
1 Hz/10 Hz, PBP839/98811/113/3132 ± 3/145 ± 4Two-sample t-test0.01
3(C)Ctrl/Myo-1, KCl86/673/33/343.19 ± 0.02/59.32 ± 0.03Two-sample KS-test<0.001
3(D)Ctrl/Myo-1, KCl86/673/33/337 ± 3/13 ± 3Two-sample t-test<0.001
5(C)
Baseline
Ctrl(-)/Ctrl(+)29/12227/645/120.048 ± 0.017/0.05 ± 0.0012Two-sample KS-test0.42
Ctrl(-)/Myo-1(+)29/6927/405/80.048 ± 0.017/0.047 ± 0.01Two-sample KS-test0.42
Ctrl(-)/PBP(+)29/2127/475/80.048 ± 0.017/0.048 ± 0.008Two-sample KS-test0.42
Ctrl(-)/EGTA(+)29/5127/305/60.048 ± 0.017/0.03 ± 0.006Two-sample KS-test0.88
Ctrl(-)/DMSO(+)29/4427/345/50.048 ± 0.017/0.046 ± 0.01Two-sample KS-test0.43
Ctrl(+)/Myo-1(+)122/6964/4012/80.05 ± 0.0012/0.047 ± 0.01Two-sample KS-test0.43
Ctrl(+)/PBP(+)122/2164/4712/80.05 ± 0.0012/0.048 ± 0.008Two-sample KS-test0.43
Ctrl(+)/EGTA(+)122/5164/3012/60.05 ± 0.0012/0.03 ± 0.006Two-sample KS-test0.88
Ctrl(+)/DMSO(+)122/4464/3412/50.05 ± 0.0012/0.04608 ± 0.01053Two-sample KS-test0.13
Myo-1(+)/PBP(+)69/2140/478/80.047 ± 0.01/0.04831 ± 0.00749Two-sample KS-test0.88
Myo-1(+)/EGTA(+)69/5140/308/60.047 ± 0.01/0.03201±0.00639Two-sample KS-test0.43
PBP(+)/EGTA(+)21/5147/308/60.048 ± 0.008/0.03 ± 0.006Two-sample KS-test0.43
5(C)
20 Hz stimulation
Ctrl(-)/Ctrl(+)29/12227/645/120.026 ± 0.002/0.013 ± 0.0003Two-sample KS-test<0.001
Ctrl(-)/Myo-1(+)29/6927/405/80.026 ± 0.002/0.029 ± 0.003Two-sample KS-test0.16
Ctrl(-)/PBP(+)29/2127/475/80.026 ± 0.002/0.03 ± 0.002Two-sample KS-test0.63
Ctrl(-)/EGTA(+)29/5127/305/60.026 ± 0.002/0.013 ± 0.002Two-sample KS-test<0.001
Ctrl(+)/Myo-1(+)122/6964/4012/80.013 ± 0.0003/0.02907 ± 0.00264Two-sample KS-test0.02
Ctrl(+)/PBP(+)122/2164/4712/80.013 ± 0.0003/0.03 ± 0.002Two-sample KS-test0.02
Ctrl(+)/EGTA(+)122/5164/3012/60.013 ± 0.0003/0.013 ± 0.002Two-sample KS-test0.63
Ctrl(+)/DMSO(+)122/4464/3412/50.013 ± 0.0003/0.012 ± 0.0006Two-sample KS-test0.66
Myo-1(+)/PBP(+)69/2140/478/80.029 ± 0.003/0.03 ± 0.002Two-sample KS-test0.16
Myo-1(+)/EGTA(+)69/5140/308/60.029 ± 0.003/0.013 ± 0.002Two-sample KS-test0.02
PBP(+)/EGTA(+)21/5147/308/60.03 ± 0.002/0.013 ± 0.002Two-sample KS-test0.007
5(B)MC-Model (-)10010008.50E-02
MC-Model (+)10010003.00E-02
5(E)
Baseline
Ctrl(-)/Ctrl(+)60/34327/645/120.02 ± 0.003/0.02 ± 0.01Two-sample KS-test0.14
Ctrl(-)/Myo-1(+)60/17427/495/80.02 ± 0.003/0.02 ± 0.002Two-sample KS-test0.59
Ctrl(-)/PBP(+)60/5527/345/80.02 ± 0.003/0.02 ± 0.005Two-sample KS-test0.14
Ctrl(-)/EGTA(+)60/6327/305/60.02 ± 0.003/0.016 ± 0.005Two-sample KS-test0.14
Ctrl(+)/Myo-1(+)343/17464/4012/80.02 ± 0.01/0.02 ± 0.002Two-sample KS-test1
Ctrl(+)/PBP(+)343/5564/3412/80.02 ± 0.01/0.02 ± 0.005Two-sample KS-test0.59
Ctrl(+)/EGTA(+)343/6364/3012/60.02 ± 0.01/0.016 ± 0.005Two-sample KS-test0.31
Myo-1(+)/PBP(+)174/5540/348/80.02 ± 0.002/0.02 ± 0.005Two-sample KS-test0.89
Myo-1(+)/EGTA(+)174/6340/308/60.02 ± 0.002/0.016 ± 0.005Two-sample KS-test0.14
5(E)
20 Hz stimulation
Ctrl(-)/Ctrl(+)60/34327/645/120.025 ± 0.001/0.02 ± 0.0006Two-sample KS-test0.59
Ctrl(-)/Myo-1(+)60/17427/495/80.025 ± 0.001/0.025 ± 0.0005Two-sample KS-test1
Ctrl(-)/PBP(+)60/5527/345/80.025 ± 0.001/0.024 ± 0.003Two-sample KS-test0.31
Ctrl(-)/EGTA(+)60/6327/305/60.025 ± 0.001/0.015 ± 0.001Two-sample KS-test0.001
Ctrl(+)/Myo-1(+)343/17464/4012/80.02 ± 0.0006/0.025 ± 0.0005Two-sample KS-test0.89
Ctrl(+)/PBP(+)343/5564/3412/80.02 ± 0.0006/0.024 ± 0.003Two-sample KS-test0.31
Ctrl(+)/EGTA(+)343/6364/3012/60.02 ± 0.0006/0.015 ± 0.001Two-sample KS-test0.001
Myo-1(+)/PBP(+)174/5540/348/80.025 ± 0.0005/0.024 ± 0.003Two-sample KS-test0.14
Myo-1(+)/EGTA(+)174/6340/308/60.025 ± 0.0005/0.015 ± 0.001Two-sample KS-test0.001
Displacement
(related to 6B)
Ctrl(-) Last 5/First 5 s122751.17 ± 0.12Two-sample t-test0.001
Ctrl(+) Last 5/First 5 s3564121.08 ± 0.05Two-sample t-test0.02
Myo-1(+) Last 5/First 5 s244081.18 ± 0.12Two-sample t-test<0.001
6(D) Change in VelocityCtrl(-)/Ctrl(-)122751.54 ± 0.13Two-sample t-test<0.001
Ctrl(+)/Ctrl(+)3564121.2 ± 0.016Two-sample t-test0.022
Myo-1(+)/Myo-1(+)244081.31 ± 0.02Two-sample t-test<0.001
PBP(+)/PBP(+)174781.27 ± 0.03Two-sample t-test0.002
EGTA(+)/EGTA(+)203061.29 ± 0.02Two-sample t-test0.002
6(E) Change in AngleCtrl(-)/Ctrl(-)122751.02 ± 0.04Two-sample KS-test1
Ctrl(+)/Ctrl(+)3564120.95 ± 0.04Two-sample KS-test1
Myo-1(+)/Myo-1(+)244080.88 ± 0.03Two-sample KS-test0.65
PBP(+)/PBP(+)174781.03 ± 0.04Two-sample KS-test1
EGTA(+)/EGTA(+)203061.06 ± 0.04Two-sample KS-test1
Figure 1—figure supplement 1(A)Ctrl/DMSO-15/84/31.00 ± 0.17/0.97 ± 0.16Chi-squared test0.89
Figure 2—figure supplement 1(A)Ctrl/DMSO-7/93/31.0 ± 0.1/0.86 ± 0.09Two-sample t-test0.43
Figure 2—figure supplement 1(B)Ctrl/Myo-1-9/103/32.74 ± 0.5/3.42 ± 0.98Two-sample t-test0.56
Ctrl/PBP-9/73/32.74 ± 0.5/1.47 ± 1.18Two-sample t-test0.3
Figure 2—figure supplement 1(C)Ctrl/Myo-1, KCl86/673/33/338 ± 3/14±3Two-sample t-test0.40
Figure 2—figure supplement 1(E)Ctrl/Myo-1-9/103/32.05 ± 0.16/2.06 ± 0.11Two-sample t-test0.93
Ctrl/PBP-9/73/32.05 ± 0.16/2.39 ± 0.07Two-sample t-test0.09
Figure 2—figure supplement 1(F)Ctrl/Myo-1-9/103/33.6 ± 0.1/3.42 ± 0.06Two-sample t-test0.2
Ctrl/PBP-9/73/33.6 ± 0.1/3.51 ± 0.07Two-sample t-test0.57
Figure 2—figure supplement 1(G)Ctrl/Myo-1-9/103/3y = 0.50915–0.0016552 x (Linear Fit)Linear Fit0.57
Ctrl/Myo-1; 1 st vs 2–5-9/103/30.52 ± 0.06/0.50 ± 0.02Two-sample t-test0.81
Figure 2—figure supplement 1(H)Ctrl/PBP-9/73/3y = 0.61391 + 0.00052562 x (Linear Fit)Linear Fit0.93
Ctrl/PBP; 1 st vs 2–5-9/73/30.61 ± 0.04/0.62 ± 0.02Two-sample t-test0.85
Figure 3—figure supplement 1(A)Ctrl/Myo-1, KCl86/673/33/343.23 ± 0.02/41.15 ± 0.02Two-sample KS-test<0.001
Figure 3—figure supplement 1(B)Ctrl/Myo-1137/813/33/334 ± 2/33 ± 3Two-sample t-test0.84
Figure 4—figure supplement 1(H)Ctrl Disappear/Re-appear16/1664/6412/121.14 ± 0.105/0.88 ± 0.062Two-sample t-test0.04

Next, we examined how inhibition of myosin V affects the reuse of individual release sites using a paired-pulse protocol at 1000 ms (Maschi and Klyachko, 2017). In this analysis, for each stimulus pair, we identified a subset of boutons in which release events were detected for both stimuli in the pair; we then determined the probability that the two release events occur at the same release site. This protocol measures vesicle refilling of the release site on short timescales. We found that both myosin V inhibitors, MyoVin-1 or PBP, markedly reduced the reuse probability of release sites (Figure 1B; Table 1), while DMSO alone had no effect (Figure 1—figure supplement 1, Table 1). Furthermore, we observed that this effect of myosin V inhibition dissipated with increase in time between release events, and was no longer apparent when release events were separated by more than ~5 s (Figure 1C, Table 1). These findings suggest that myosin V regulates rapid refilling of release sites. However, on longer timescales, once the release sites are refilled, the release process itself appears not to depend on myosin V.

The change in release site reuse probability could reflect the role of myosin V in refilling the sites with a new vesicle, but this effect could also arise if the number of release sites is altered by myosin V inhibition. We thus examined whether inhibition of myosin V affects the number and basic structural organization of release sites. The number of clusters/release sites in individual boutons was not affected by myosin V inhibition (Figure 1D, Table 1). Ultrastructural analysis using a Large-Area Scanning Electron Microscopy (LaSEM) further indicated that the AZ size was also not altered by myosin V inhibition (Figure 1E, Table 1). Interestingly, we noted a small, but significant effect of myosin V inhibition on increasing the average distance from release events to the AZ center (Figure 1F, Table 1). One possible interpretation of this result is that peripheral release sites are engaged more frequently when myosin V is inhibited (see below for additional analyses).

Thus far these results favor the first model that myosin V has an important role in the refilling of release sites rather than the exocytosis itself.

Myosin V controls release site refilling during high-frequency stimulation

If myosin V plays a role in refilling of the release sites, inhibition of myosin V should have a larger effect on release magnitude at higher stimulation frequencies when demand for vesicles is increased. To determine if this is the case, we measured the magnitude of release at individual boutons (as given by the total vGlut1-pHluorin signal) evoked by high-frequency trains. Both MyoVin-1 and PBP reduced the magnitude of synaptic release evoked by 20 stimuli at 50 Hz to half compared with the control (Figure 2A,B, Table 1), while DMSO alone had no effect (Figure 2—figure supplement 1A, Table 1). Inhibition of myosin V also slowed the kinetics of release to a similar degree (as evident by the change in the slope of the vGlut1-pHluorin signal increase during stimulation, Table 1), which is consistent with slower refilling of the release sites when myosin V is inhibited. We further examined how release magnitude is affected in a wide range of stimulus frequencies from 0.1 Hz to 50 Hz. We found that inhibition of myosin V had a larger effect on the magnitude of release at higher stimulus frequencies while it did not have any measurable effect on release at a very low 0.1 Hz stimulation frequency (Figure 2C, Table 1). The observation that release amplitude is normal at very low stimulus frequencies supports the notion that the exocytosis process itself is not strongly affected by myosin V inhibition.

Figure 2 with 1 supplement see all
Myosin V controls release site refilling during high-frequency stimulation.

(A) Examples of vGlut1-pHluorin responses at single hippocampal boutons to 20 stimuli trains at 50 Hz for Ctrl (black) and Myo1 (red). (B) Ratio of vGlut1-pHluorin responses in (A) at individual boutons for Myo1 or PBP normalized to the control. (C) Effect of myosin inhibition with Myo1 on the amplitude of vGlut1-pHluorin responses to 20 stimuli trains at individual boutons as a function of the train frequency, averaged across all boutons in the movie and normalized to a control at a corresponding frequency. (D–E) Effect of myosin inhibition with Myo1 (D) or PBP (E) on the distribution of distances of release events to AZ center evoked at 1 Hz vs 10 Hz. (F) Analysis of data in (D,E) showing a mean distance of release events to the AZ center per bouton at 1 Hz or 10 Hz for Ctrl, Myo1 and PBP, respectively. ***=P < 0.001, **=P < 0.01, *=P < 0.05, two-sample t-test; ns – not significant.

https://doi.org/10.7554/eLife.39440.005

In addition to these temporal requirements for myosin V in supporting release, our result above suggested that spatial distribution of release may be altered by myosin V inhibition. We thus further examined this function of myosin V during high-frequency stimulation. Inhibition of myosin V significantly exacerbated the use of peripheral release sites during high frequency stimulation (Figure 2D,E, F, Table 1) suggesting its role in controlling not only temporal but also the spatial distribution of release site reuse. One plausible explanation for the spatial effects of myosin V inhibition is its involvement in endocytosis independently of its role in refilling of release sites. If this is the case, inhibition of myosin V could cause an increase in the distance of peripheral release sites to the center because of an activity-dependent accumulation of vesicle components on the plasma membrane and an increase in the AZ size. Ultrastructural EM analysis showed no significant changes in the AZ size by myosin V inhibition both at baseline (Figure 1E) and during KCl-induced depolarization (Figure 2—figure supplement 1B) arguing against major effects of myosin V inhibition on endocytosis. To confirm this observation, we further examined if inhibition of myosin V affects endocytosis in live and active synapses. The decay of vGlut1-pHluorin signal following a stimulus train is determined by endocytosis and subsequent vesicle reacidification. Changes in the decay of the vGlut1-pHluorin signal in our measurements can thus be interpreted to reflect predominantly changes in endocytosis (Atluri and Ryan, 2006). We found that inhibition of myosin V had no measurable effect on the decay of the vGlut1-pHluorin signal following 50 Hz trains (Figure 2—figure supplement 1C-E). Moreover, if inhibition of myosin V blocks endocytosis, application of sequential stimulus trains would be expected to cause surface accumulation of VGlut1-pHluorin and a corresponding increase in the bouton fluorescence. We compared the amplitudes of VGlut1-pHluorin signal in five consecutive 50 Hz trains separated by 20 s each, and did not observe any measurable changes from one train to the next under control conditions or in the presence of myosin V inhibitors (Figure 2—figure supplement 1F-H). Combined, these results indicate that the shift in spatial localization of release events upon myosin V inhibition is not caused by an increase in the AZ size and is unlikely to be mediated by altered endocytosis.

Together these observations support the first model in which myosin V plays a major role in regulating the refilling of the release sites rather than the exocytosis or endocytosis processes themselves. Our results further suggest a role for myosin V in regulating spatial distribution of release, with a shift towards more peripheral release sites when myosin V is inhibited.

Inhibition of myosin V causes a vesicle docking defect during sustained activity

If myosin V indeed plays a role in release site refilling, we predicted that inhibition of myosin V should reduce the number of docked vesicles at the AZ during sustained activity. We tested this idea using a scanning electron microscopy (LaSEM) of hippocampal cultures treated or not with MyoVin-1 for 20 min and stimulated via a KCl-induced depolarization for 10 min (with a 10 min delay after beginning of MyoVin-1 treatment), and fixed immediately thereafter (Figure 3A). Experiments were accompanied by the corresponding control measurements without KCl depolarization. We quantified the effect of myosin V inhibition on vesicle localization by dividing the AZ into 0.5 nm segments and calculating the distance from each segment to the closest vesicle (Figure 3B). The distribution of these minimal distance values provides a measure of the vesicle location relative to the AZ, and has an additional advantage of being independent of the total number of vesicles detected in each bouton. We observed that vesicles were localized significantly farther away from the AZ when myosin V was inhibited during sustained depolarization with KCl (Figure 3A,C, Table 1), but this effect was not apparent in cultures not treated with KCl (Figure 3—figure supplement 1A, Table 1).

Figure 3 with 1 supplement see all
Inhibition of myosin V causes a vesicle docking defect during sustained activity.

(A) LaSEM of individual hippocampal boutons in cultures depolarized by KCl application (55 mM) for 10 min in the presence or absence of Myo1 (20 min), immediately followed by fixation. (B–D) Membrane opposite to the PSD was divided into 0.5 nm sections (B) and the distance from each section to the closest vesicle was determined and plotted as a cumulative histogram (C). This subset of closest vesicles was subsequently used to estimate the relation between docked and tethered vesicle populations (D). We considered vesicle as ‘docked’ when the distance from the AZ section to the vesicle center was under 30 nm and ‘tethered’ when the distance was under 100 nm. Scale bar: 200 nm. ***=P < 0.001; **=P < 0.01; two-sample KS-test (C) or two-sample t-test (D). ns = not significant.

https://doi.org/10.7554/eLife.39440.007

To further quantity changes in vesicle localization caused by inhibition of myosin V, we examined the ratio of ‘docked’ vesicles and more loosely ‘tethered’ vesicles: docked vesicles were defined as vesicles adjacent to the AZ, with the vesicle center within 30 nm from the AZ (which is equivalent to vesicle membrane being less than ~5 nm from the AZ assuming a vesicle diameter of 50 nm, and within an error of measurement of our LaSEM that has a pixel size of 5 nm), while ‘tethered’ vesicles included all vesicles with a center within 100 nm from the AZ. Only the subset of vesicles identified above as being the closest to the AZ were used for this analysis to be independent of the total number of vesicles detected in each bouton. We found that the ratio of ‘docked’ to ‘tethered’ vesicles at the AZ was markedly reduced by myosin V inhibition during sustained depolarization with KCl (Figure 3D, Table 1), but not in the absence of KCl (Figure 3—figure supplement 1B, Table 1).

Mechanistically, the role of myosin V in controlling vesicle localization and release site refilling can involve its function as a vesicle transporting motor to the release sites and/or as a tether of vesicles at the release sites or both. Below we employed a single vesicle tracking approach to discern these possibilities.

Tracking individual synaptic vesicles during recycling supports a model of continuous vesicle shuttling between two vesicle pools

To better understand the release site refilling process, we used our established approach for tracking individual synaptic vesicles during recycling and translocation to the AZ (Forte et al., 2017; Gramlich and Klyachko, 2017; Peng et al., 2012). Individual vesicles were labeled with a lipophilic FM-like dye SGC5 via compensatory endocytosis using a pair of stimuli at 100 ms. Due to very low basal release probability (~0.06, Table 1) of hippocampal synapses at 37°C, this protocol labels either none or at most a single vesicle in vast majority of boutons (Forte et al., 2017; Peng et al., 2012); a small subset of boutons with two or more detected vesicles were excluded from further analyses. This imaging approach permits us to track individual vesicles with ~20 nm precision within hippocampal boutons (Figure 4A) (Forte et al., 2017; Peng et al., 2012).

Figure 4 with 1 supplement see all
Vesicle disappearance/reappearance events.

(A) Sample track of a vesicle that disappeared during observation. Track is color-coded by instantaneous vesicle speed. (B) Parameters of vesicle image from (A) before and after vesicle disappearance indicated by a dotted line. Amplitude of the vesicle image (PSF) normalized to image bit-depth (214-1) (Top), and localization error of vesicle position as a function of time (Bottom). (C) Half-width of a Gaussian fit to the whole-synapse image (σ, Top) and whole-synapse integrated intensity (Bottom) before (Black) and after (Red) vesicle detection was lost, for the same vesicle as in A. Raw images (insets) of vesicle image at different time points show that vesicle is still present in the synapse after detection is lost. (D) Sample track of a vesicle that undergoes multiple disappearance/reappearance events. Track is color-coded with initial track shown in black, the first reappearance in red, the second reappearance in blue. (E) Parameters of vesicle track from (D). Vesicle position over time plotted as a distance from center of the bouton (Top). Amplitude of the vesicle image (Middle) and localization error of vesicle position (Bottom) as a function of time. Periods of vesicle disappearance are highlighted in grey. The color coding is the same as in (D).

https://doi.org/10.7554/eLife.39440.009

We previously found that the majority of vesicles undergo transitions between epochs of directed, diffusive and stalled motion during recycling (Forte et al., 2017). We and others also previously reported that vesicle mobility decreased with time, which has been interpreted to represent vesicle settling within one of the functional vesicle pools (Kamin et al., 2010), a membrane-bound readily-releasable pool (RRP) or an inner recycling or reserved pool. Here, we observed that a large subset of these settled vesicles (>50% during 2 min observation) became more mobile and then disappeared during our observation window (Figure 4A,B). The same (or possibly another) vesicle was also often observed (re-) appearing in the same bouton with some delay (Figure 4D,E). These appearance and disappearance events were often observed multiple times in the same bouton (Figure 4D,E) and both occurred at constant rates, with the rate of disappearance ~2 – 3 fold higher than the rate of appearance (Figure 5B-E, Table 1; the fraction of disappearing vesicles per second (δ-rate): 0.046 ± 0.016 fraction/sec; fraction of appearing vesicles per second (α-rate): 0.022 ± 0.010 fraction/sec; see Materials and Methods for quantification details). We note that because disappearance is the only criteria for the vesicle inclusion in our initial analysis, the initial conditions for calculating the disappearance and appearance rates are not equivalent. As a result, the fractional curves in (Figure 5B,D) representing these two processes are visually different, although the actual rates are constant in both cases. Specifically, only vesicles that were detected at T = 0 were counted in disappearance analysis and only a single disappearance event was counted per bouton. To be able to distinguish the disappearance and reappearance events, the appearance analysis was limited to the same subset of synapses in which vesicle disappearance was observed first; but this analysis placed no limitation on the number of vesicle appearance events counted in the same bouton. As a result of these initial conditions, the number of labeled vesicles that could disappear is maximal at T = 0 and is continuously drawn down with time leading to a visually apparent reduction in the disappearance rate (Figure 5B). This effect is caused by un-labeled vesicles replacing the labeled ones and also disappearing at the same rate but without being counted. This reduces the apparent disappearance rate, while the actual rate remains constant. In contrast, these limits on initial conditions do not affect the appearance rate because the number of labeled vesicles that can disappear is constantly replenished and appearance events are counted every time even if they appear multiple times in the same bouton. Consequently, the appearance rate remains linear.

Activity- and myosin V- dependent vesicle shuttling between a membrane and an inner pools.

(A) Model hypothesis of vesicle exchange between two pools resulting in observed disappearance and re-appearance. At T = 0 all labeled vesicles are assumed to be in Pool I; vesicles undergo transition toward Pool II at rate (δ) and re-appear with rate (α). With time (T >> 0) the experimentally observed δ–rate is lower because un-labeled vesicles replace the labeled ones and disappear at the same rate but without being counted. (B) Vesicle disappearance as a function of time plotted as a fraction of total vesicles observed. Ctrl-No Stim represents vesicle disappearance rate in the absence of activity at any time during observation; other data show effects of stimulation (+Stim, 20 Hz applied at 10 – 20 s period) without (white) or with (red) myosin-V inhibition (Myo-1), or EGTA-AM (blue). Computational model is shown as solid lines. Note that the X-axis starts at T = 5 s because all tracks were required to be observed for at least 5 s to be included in analysis. (C) Mean disappearance rate (δ) from exponential-recovery fits to data in (B) at baseline (5–10 s) and during stimulation (10 – 20 s). Error-bars are mean-residual of fits to data. (D) Same as (B) but for vesicle appearances in different conditions indicated plotted as a fraction of total vesicles observed. Linear fits to data show constant rates (solid lines). (E) Same as (C) for the mean appearance rate (α) from linear fits to data in (D). Error-bars are mean-residual of linear fits to data. ***=P < 0.001; two-sample KS-test. ns = not significant.

https://doi.org/10.7554/eLife.39440.011

Because vesicle disappearance and reappearance is often observed multiple times in the same bouton, we hypothesized that this process represents motion of the same vesicle in and out of focus, since very few boutons have more than one labeled vesicle present, and there is no any additional source of the dye other than the vesicle that was initially labeled. These vesicle disappearance/appearance observations may thus be mechanistically explained by a model hypothesis of vesicle transitions between two distinct pools (Figure 5A), one of which is in the focal plane, and the other one is outside of the focal plane. If the total number of vesicles exchanged between the two pools is constant in our model, the α-rate and δ-rate infer the fraction of total vesicles in each pool, then the 2 – 3 fold differences in the α- and δ-rates can be accounted for by assuming that the Pool II is 2 – 3 times larger than Pool I. Experiments below are designed to provide evidence to support this 2-pool model and will examine how vesicle mobility between (and retention at) these pools are regulated by activity and myosin V. We will then provide evidence that the smaller Pool I is a RRP-like membrane pool localized close to the source of calcium influx into the synapse, while the larger Pool II is farther away from the calcium influx and thus represents an inner pool.

While we hypothesize that vesicle disappearance can be explained by loss of detection due to large-scale displacement away from the focal plane within the synaptic bouton, vesicle disappearance can also be caused by either exocytosis or a vesicle leaving the synapse via axonal transport in a process known as an inter-synaptic vesicle exchange (Gramlich and Klyachko, 2017). To distinguish these possibilities we quantified changes in the integrated intensity of the entire synapse in which vesicle disappearance occurred (Figure 4C). The total synapse intensity arises predominantly from the intensity of the labeled vesicle and a nonspecific background; it is thus expected to undergo a stepwise decrease upon vesicle fusion and loss of the dye. We found that the integrated synapse intensity remained unchanged at the time point of vesicle disappearance (Figure 4C and Figure 4—figure supplement 1D) for the vast majority of disappearing vesicles (~97%). Further, the few cases when integrated intensities did decrease matched the gradual intensity decrease profile observed for inter-synaptic vesicle exchange events (Figure 4—figure supplement 1A-D). These results suggest that the vast majority of disappearance events are not caused by exocytosis, but rather by loss of detection due to vesicle moving out of focus.

The shape of the vesicle image is represented by the point-spread function (PSF), which becomes increasingly broader as vesicle goes out of focus (Figure 4C) eventually causing loss of detection when a preset threshold is reached. This notion is evident in the corresponding increase in the PSF σ (half-width) at the time points of vesicle disappearance and a decrease in the PSF σ at time points of subsequent vesicle reappearance (Figure 4—figure supplement 1H). These changes were not caused by differential bleaching of vesicle signal vs background (Figure 4—figure supplement 1I). We were also able to correlate the observed range of vesicle PSF σ values (~120 – 150 nm) with the range of detectable vesicle motion along the Z-axis of ~100 nm (Figure 4—figure supplement 1G), suggesting that the two vesicle pools are at least 100 nm apart from each other. To further clarify the causes of vesicle disappearance we re-analyzed the trajectories of disappearing vesicles under less strict detection criteria by allowing detection of the more out of focus vesicles with broader images (Figure 4—figure supplement 1E,F). We found that with less stringent detection criteria, the vast majority of disappearing vesicles could be tracked beyond the initial disappearance point, supporting the notion that loss of tracking occurred due to vesicle motion out focus rather than fusion. Finally, we observed that the vesicle velocity and displacement both increased significantly in the last 2 s before disappearance, consistent with vesicle acceleration immediately prior to disappearance (Figure 6D, Table 1). These results provide further evidence that vesicle disappearance was caused by loss of detection due to vesicle motion away from the focal plane rather than vesicle fusion.

Inhibtion of myosin V does not affect vesicle mobility before disappearance.

(A) Spatial analysis of vesicle displacement based on the radius of a circle encompassing 95% of vesicle trajectory for the first 5 s (50 frames, black circle) and for the last 5 s (red circle) before disappearance. (B) Cumulative distributions for the spatial analysis in (A) in control with stimulation conditions compared for all disappearing vesicles and fit to exponential recovery function (solid line). (C) Same as (B) in the presence of Myo-1 and stimulation. (D) The ratio of instantaneous speed during the last 2 s and the first 2 s of observation. Error-bars are determined from residuals of cumulative fits. (E) The ratio of angular displacement during the last 2 s and the first 2 s of observation. Error-bars are determined from SEM. ***=P < 0.001; *=P < 0.05; two-sample t-test (D) or two-sample KS-test (E).

https://doi.org/10.7554/eLife.39440.012

Using less strict detection criteria and thus the extended ability to track vesicle trajectories beyond the initial disappearance point, we also confirmed our integrated intensity analysis above showing that a small fraction (<3%) of the disappearance events was caused by vesicle leaving the synapse and entering the axon (Figure 4—figure supplement 1A-D), which is a process we have previously reported (Gramlich and Klyachko, 2017). Thus vesicle disappearance is not caused by travel out of the synapse in >97% of the cases. We note that generally, such inter-synaptic vesicle exchange events occur more frequently than the 3%, but the majority of these events do not result in loss of tracking and are automatically excluded from all our current analyses because their displacement is beyond the spatial limits we set for the synapse size (displacement >0.8 µm from the synapse center).

Taken together these results indicate that in the vast majority of cases, the observed vesicle disappearance is not caused by exocytosis or travel out of the synapse, but rather by vesicle acceleration and motion to another pool located out of focus; subsequent vesicle re-appearance represses vesicle transition back.

Vesicle shuttling between the pools is activity- and myosin V dependent

We previously found that a component of vesicle mobility during recycling is regulated in an activity-dependent manner; however which step in the vesicle cycle this component corresponds to is unknown. Here we found that the rate of vesicle disappearance was reduced 2 – 3-fold during high-frequency stimulation (Figure 5B,C, Table 1). This activity-evoked change in the rate had a very fast onset (<0.5 s) but persisted long after the end of stimulation (Figure 5B), consistent with the rapid rise of presynaptic calcium at the beginning of stimulus trains and a slow calcium clearance following the cessation of stimulation (Neher and Sakaba, 2008). Importantly, we observed that inhibition of myosin V with either Myo1 or PBP eliminated the effect of activity on vesicle disappearance during stimulation, without affecting the baseline disappearance rate before stimulation (Figure 5B,C, Table 1), while DMSO alone had no effect (Table 1). In contrast, the rate of subsequent vesicle appearance in the same set of boutons was not strongly affected by either high-frequency stimulation or myosin V inhibition (Figure 5D,E, Table 1). These results suggest that myosin V mediates the activity-dependence of vesicle disappearance, but not vesicle re-appearance.

To better understand the vesicle disappearance, we used Monte Carlo simulations to model this process (Figure 5A, and solid lines in 5B, Table 1). Vesicle disappearance rate was calculated using a computational model that simulates individual vesicle disappearances from a pool of vesicles (see cartoon in Figure 5A and Appendix for model details). A single constant disappearance probability captures the baseline vesicle disappearance rate (Pδ, baseline= 8.5×10−2 fraction/sec). This result suggests that vesicles are randomly exchanged in and out of focus without preference for residence time (i.e. newly arrived vesicles are just as likely to leave as established vesicles). The model shows that the experimentally observed reduction in disappearance rate upon stimulation can be modeled with an instantaneous 3-fold reduction in disappearance probability (from Pδ, base-line = 8.5×10−2 fraction/sec, to Pδ, stim = 3.0×10−2 fraction/sec) and the effect of myosin V inhibition is captured by an increase in the disappearance probability during stimulation back to the baseline rate (Pδ=8.5×10−2 fraction/sec). These results support the above observations that the vesicle disappearance is rapidly modulated by activity and that this modulation is myosin V dependent.

Vesicle shuttling represents transitions between a membrane pool and an internal pool

How are vesicle disappearance and reappearance related to vesicle location inside the synaptic bouton and specifically the AZ? We hypothesized that the smaller Pool I, which is rapidly regulated by activity/calcium influx, is an RRP-like membrane vesicle pool localized close to the source of calcium influx into the synapse, while the larger Pool II represents an inner pool, further away from the calcium sources. The 2 – 3 fold difference in size between the Pools I and II in our model is also consistent with a smaller membrane-bound RRP and larger internal recycling pool described previously (Denker and Rizzoli, 2010).

To test this model prediction and relate the location of the two pools to the source of calcium influx, we used a slow calcium chelator EGTA. EGTA is too slow to buffer rapid calcium rise in the close proximity to voltage-gated calcium channels at the plasma membrane, but is effective in buffering subsequent slow calcium elevation in the interior of the bouton due to diffusion. Indeed, it has been shown that EGTA does not affect vesicle release, but effectively blocks facilitation and augmentation, the two forms of short-term synaptic enhancement that dependent on residual calcium elevation during repetitive activity (Deng and Klyachko, 2011; Regehr, 2011). We thus pre-incubated neurons with a cell-permeable EGTA-AM for 20 min. Calcium buffering with EGTA had no effect on the activity-dependent change in the disappearance rate during high-frequency stimulation (Figure 5B,C, Table 1) suggesting that vesicle disappearance occurs from a pool located in a close proximity to the source of calcium influx, that is the AZ. In contrast, the appearance rate in the same set of boutons was significantly reduced by EGTA indicating efficiency of EGTA treatment (Figure 5D,E, Table 1). These results suggest that in contrast to vesicle disappearance, the appearance represents transition from a pool located farther away from the source of calcium influx. We note that although we did not detect a measurable effect of activity on the appearance rate, it is possible that the calcium-dependence of this process is already close to saturation even in the basal conditions, but is unmasked by lowering the basal calcium levels by EGTA.

The finding that EGTA strongly affects only the vesicle appearance rate (and thus vesicles in Pool II) also provides a simple way to estimate the ratio of vesicles that start in each pool in beginning of our observations. Since disappearance is the only criteria for the vesicle inclusion in our initial analysis and due to the arbitrary relationship between the focal plane and the AZ at any given synapse, there is no a priori information that would allow us to make the initial assignment of the vesicles to Pool I or Pool II. Our measurements of the disappearance rate thus reflect a mixture of vesicles that are in Pool I and Pool II at the beginning of observation. To perform this analysis, we assumed that only the vesicle appearance rate is truly affected by EGTA (based on our observation that disappearance rate for EGTA and Ctrl during stimulation are the same), while the small effect of EGTA on the disappearance rate (vesicles in Pool I) is caused by contribution from Pool II vesicles. Within these assumptions, the model (Appendix) and EGTA results suggest that among all vesicles assumed to be in Pool I in the beginning of observations, 75 ± 11% actually start in Pool I, while 25 ± 11% are physically located in Pool II. This result is consistent with the Pool II being larger, and having a slower exit rate, because the probability that a given labeled single vesicle in our experiment will leave the Pool II is smaller than if it would be in Pool I and thus less likely to be included in our disappearance analysis.

Together these results support the model that vesicle shuttling occurs between the smaller membrane and larger internal pools, with vesicle transitions away from (but not towards) the plasma membrane pool being both activity- and myosin V dependent. In contrast with widely assumed model of unidirectional vesicle flow during recycling towards the AZ, our results suggest an ongoing vesicle shuttling and exchange between an inner pool and a membrane pool.

The role of myosin V in vesicle tethering at the membrane pool vs vesicle transport

Our results suggest that vesicle disappearance rate (translocation away from the membrane pool) is strongly reduced by activity in a manner that requires myosin V. Myosin V, a vesicle-associated protein, may function in this process as a vesicle-transporting motor and/or as a tethering/docking factor that retains vesicles in a membrane pool in an activity-dependent manner. To distinguish these possibilities, we examined whether inhibition of myosin V alters vesicle mobility immediately prior to disappearance.

First, we employed a spatial analysis of vesicle displacement we developed previously based on calculating the radius of a circle encompassing 95% of the vesicle track (Figure 6A) (Forte et al., 2017; Peng et al., 2012). We compared the spatial extent of vesicle motion over the first 5 s of observations and the last 5 s before disappearance. Vesicle displacement increased significantly just before disappearance (Figure 6B), and the magnitude of the increase was similar in all conditions (Table 1). Importantly, this increase in displacement was not affected by inhibition of myosin V (Figure 6C, Table 1).

We corroborated this result by examining changes in the instantaneous vesicle velocity during a 2 s period immediately before disappearance compared to the first 2 s of observation because analysis of instantaneous vesicle velocity permits higher time resolution than analysis of spatial displacement (which provides a more global assessment of vesicle mobility). Consistent with the above results, the vesicle velocity increased significantly just before disappearance under all conditions, and again this increase was not affected by inhibition of myosin V (Figure 6D, Table 1), suggesting that the activity-induced increase in vesicle mobility was not driven by myosin V.

Finally we examined the changes in vesicle angular displacement immediately prior to disappearance relative to the beginning of observation, which provides a measure of how directed the vesicle motion is (Gramlich and Klyachko, 2017). The instantaneous angular displacement was the same under all conditions and was not affected by inhibition of myosin V (Figure 6E, Table 1).

The combined total displacement, velocity, and angular displacement results suggest that vesicles that are initially tethered at the plasma membrane, become un-tethered and more mobile just before tracking is lost. Activity regulates this process by increasing the vesicle retention (by strengthening tethering) at the membrane pool in a myosin V dependent manner. These analyses further indicate that myosin V does not function in this process to drive vesicle transport away from the membrane pool. Thus the requirement for myosin V in activity-dependent vesicle retention strongly suggests the role for myosin V as a vesicle tether at the plasma membrane.

Discussion

We took advantage of nanoscale detection of individual vesicle fusion events together with single-vesicle tracking during recycling to examine the mechanisms governing release site refilling in hippocampal synapses. Our results support three key observations: (i) Myosin V plays a major role in refilling of the release sites during repetitive stimulation, but not directly in vesicle release process. Myosin V also regulates spatial distribution of release by preferentially promoting release at more central release sites. (ii) Recycling vesicles undergo a continuous bi-directional shuttling between a membrane pool and an inner pool and the rate of vesicle ‘undocking’ but not ‘docking’ is regulated by neural activity (iii) myosin V functions as a tether that retains vesicles at the plasma membrane in an activity-dependent manner, rather than a motor driving vesicle transport to the release sites. These results uncover a complex dynamic mechanism that governs vesicle availability for release in central synapses.

Myosin V in release site refilling

As highly processive motors, class V myosins have been recognized to play multiple roles in synaptic development and function, particularly in dendrites (Kneussel and Wagner, 2013; Rudolf et al., 2011). Myosin Va supports neuronal mRNA transport (Yoshimura et al., 2006), while myosin Vb mediates transport of recycling endosomes into dendritic spines and is required for induction of several key forms of long-term synaptic plasticity (Rudolf et al., 2011). Although presynaptic terminals are also rich in actin filaments, the role of actin-dependent transport and specifically myosin V in presynaptic processes remains debatable (Kneussel and Wagner, 2013). Myosin Va is the only member of the myosin family identified as a synaptic vesicle associated protein and is known to directly interact with the SNARE machinery, including syntaxin 1A and synaptobrevin, in a calcium-dependent manner (Krementsov et al., 2004; Ohyama et al., 2001; Prekeris and Terrian, 1997; Watanabe et al., 2005). Yet the role(s) of myosin V/Va in presynaptic processes remains controversial due to contradictory results from different knockout studies (Kneussel and Wagner, 2013; Rudolf et al., 2011; Schnell and Nicoll, 2001; Yoshii et al., 2013). This discrepancy could arise from developmental or compensatory effects and from the difficulty of separating pre- and postsynaptic effects of myosin V inhibition in electrophysiological recordings of postsynaptic currents. We were able to bypass both of these complications using direct recordings of vesicle release and recycling in presynaptic boutons together with an acute approach to inhibit myosin V in developed neurons.

Myosin V has been implicated in the exocytosis of secretory granules in non-neuronal cells (Eichler et al., 2006; Porat-Shliom et al., 2013; Rudolf et al., 2011) and of large dense-core vesicles in neurons (Bittins et al., 2009; Kögel et al., 2010). Here we found that in the case of synaptic vesicles, the process of exocytosis itself was not directly affected by myosin V inhibition in hippocampal synapses. In contrast, our results support a major presynaptic role for myosin V in the release site refilling process as a vesicle tether at the plasma membrane. This function of myosin V is supported by our EM results demonstrating a marked docking defect caused by inhibition of myosin V during sustained stimulation. This is in line with the published EM observations that the number of docked secretory granules in neuroendocrine cells is reduced by myosin V inhibition (Desnos et al., 2007). Importantly, our finding of increased retention/tethering of vesicles at the membrane during high-frequency stimulation is consistent with the calcium-dependent interaction of myosin V with the SNARE machinery (Krementsov et al., 2004; Ohyama et al., 2001; Prekeris and Terrian, 1997; Watanabe et al., 2005). While elucidating the specific molecular interaction mediating this function of myosin V is beyond the scope of the current study, the submicromolar calcium-dependent binding of myosin V to syntaxin-1A (Watanabe et al., 2005) may serve as a mechanism by which vesicles are specifically targeted to the release sites at the plasma membrane.

We note that our results do not argue against a possible additional role of actin cytoskeleton and myosin motors in transporting vesicles during some of the earlier steps of the recycling process, prior to vesicle tethering/docking. Indeed, we and others described a directed actin-dependent component of vesicle motion (Forte et al., 2017; Kisiel et al., 2014; Peng et al., 2012) and another member of the myosin family, myosin II, has been implicated in supporting multiple stages in the vesicle recycling, including vesicle translocation (Chandrasekar et al., 2014; Chandrasekar et al., 2013; Hayashida et al., 2015; Kisiel et al., 2014; Miki et al., 2016; Peng et al., 2012; Seabrooke et al., 2010; Takagishi et al., 2005). Notably, myosin II does not possess a significant vesicle-transporting ability (Kneussel and Wagner, 2013; Porat-Shliom et al., 2013) and is likely to act indirectly by generating tension and promoting actin dynamics which is required for processive motion of other myosin isoforms (Semenova et al., 2008). In contrast, myosin V is a highly processive motor and we previously found that it supports vesicle transport in the axon during inter-synaptic vesicle exchange in central neurons (Gramlich and Klyachko, 2017). Inhibition of myosin V also slightly, but significantly, reduced directionality of vesicle motion inside the synaptic boutons (Gramlich and Klyachko, 2017). Myosin V may thus play a dual role in vesicle recycling both as a tether at the release sites and as a transporting motor. Yet the specific stage(s) of the recycling process mediated by actin-dependent transport and myosin V or other members of the myosin family remains to be elucidated.

Myosin V in the differential engagement of central vs peripheral release sites

Our analyses of spatial distribution of release events within individual AZs suggest that central release sites are engaged more frequently under basal conditions and this preferential re-use is determined, in part, by myosin V. Moreover, we previously found that during high-frequency stimulation reuse of release sites shifts towards periphery, and here we observed that this shift is exacerbated by inhibition of myosin V. These results suggest that release sites have a spatial gradient of basal reuse probability from the center to periphery, and that the spatial shift in release site reuse is activity- and myosin V-dependent. Our analyses suggest that this spatial shift is unlikely to arise from myosin V involvement in endocytosis. The center vs periphery differences in release site reuse may arise, for example, if density of actin filaments is larger near the center of the AZ, which could increase the basal probability of vesicle tethering at the more central release sites. A shift of release towards periphery during stimulation could arise because central release sites, which are engaged first, would have a reduced availability during the progression of sustained high-frequency activity. It remains to be determined whether the spatial effect of myosin V inhibition is a reflection of myosin V’s function in vesicle tethering or relies on an independent mechanism. Notably, differential spatial distribution of functionally distinct modes of release has been observed for kiss-and-run (closer to the AZ center) vs full fusion events (more peripheral) (Park et al., 2012). While our tools do not currently permit distinguishing these modes of release, our findings of an activity-dependent shifts toward increased reuse of peripheral release sites is consistent with the shift from prevalent kiss-and-run to full fusion observed with increase in calcium elevation or increased stimulation frequency (Harata et al., 2006; Richards, 2010; Zhang et al., 2009). Our results further predict that by spatially controlling release site reuse, myosin V may play a role in setting the balance between kiss-and-run and full fusion modes of release.

A myosin V- and activity-dependent reversible vesicle tethering in release site refilling

A widely held view of vesicle recycling in central synapses assumes a unidirectional flow of vesicles from the sites of endocytosis to the RRP directly or via another recycling/reserve pool(s) (Rizzoli, 2014). Our results suggest an important revision to this view with a model of sustained bidirectional vesicle shuttling between a membrane pool and an inner pool. Within this model, vesicles that are tethered at the membrane pool by myosin V have a residential time on the order of twenty seconds in resting conditions and undergo repeated cycles of detaching and transitioning to the inner pool and back to the membrane. Our results further indicate that elevated activity shifts the equilibrium towards vesicle retention at the plasma membrane in a Myosin V-dependent manner by reducing the detachment rate 2 – 3 fold, presumably by strengthening myosinV/SNARE interactions. Although to the best of our knowledge this bi-directional shuttling process has not been described in central synapses, vesicle undocking has been reported in hippocampal synapses to occur at rates three times higher than the occurrence of spontaneous exocytosis (Murphy and Stevens, 1999). Taking the rate of spontaneous exocytosis of ~1 – 2 events per minute at 37°C under our experimental conditions (Peng et al., 2012), this gives a rough estimate for the vesicle residence times at the plasma membrane of ~10 – 20 s, similar to our results. Our observations are also reminiscent of the vesicle docking/undocking observed in ribbon synapses, which had an escape rate of 1 in 4 s, comparable to our estimates (Chen et al., 2013; Zenisek, 2008; Zenisek et al., 2000). Notably, much faster vesicle ‘approach and bounce’ events have also been observed in the Calyx of Held synapses using TIRF with a residence time of 88 ms (Midorikawa and Sakaba, 2015). Such rapid events, if present in our model system, would not be reliably detectable in our measurements because their duration is comparable to our time resolution. Interestingly, the same single-vesicle tracking measurements in the calyx of Held revealed a tethering step prior to vesicle docking with a time constant of 3 – 4 s (Midorikawa and Sakaba, 2015) which is comparable to our estimates for the vesicle residence time at the membrane pool. Thus cycles of vesicle approaching, tethering and reversal might be a common feature in different classes of synapses. The two-pool shuttling observed here could also provide a mechanistic basis for the two-step actin-dependent model of RRP refilling, which was recently proposed for cerebellar synapses (Miki et al., 2016).

The important limitation of the current study is in defining the functional identity of the two vesicle pools involved in vesicle shuttling/tethering. Our EGTA experiments provide evidence that the activity/myosin V-dependent pool is localized in the close proximity to the sources of calcium influx, that is the plasma membrane, but to what extent this pool represents the functionally defined RRP remains to be determined. Our estimate of the relative sizes of the two pools is also comparable to the differences between functional RRP and recycling pools defined previously (Harata et al., 2001). Nevertheless, our single vesicle tracking measurements do not distinguish whether vesicles travel along or perpendicular to the AZ plane. The effect of EGTA could thus in principle be also explained by two vesicle pools both localized at the AZ, but at a systematically distinct distance to the source of calcium influx, that is calcium channels. However, this model is unlikely for two reasons: (i) a large proportion (~60%) of the calcium channels are mobile in the AZ plane with the median surface area explored by individual channel molecules of ~200 – 250 nm (Schneider et al., 2015) comparable to dimensions of the entire AZ (Schikorski and Stevens, 1997); (ii) the large spatial domain of vesicle motion that we observed in our tracking experiments, which commonly spans several hundred nanometers over our observation period and includes epochs of fast diffusion and directed motion (Forte et al., 2017); such extensive vesicle mobility is difficult to reconcile with the motion restricted along the AZ given the very limited AZ dimensions and the vesicle crowding at the AZ which unlikely to allow fast diffusion or directed motion. Further studies are needed to define the precise spatial localization of these two pools and their correspondence to the RRP and recycling pool. Revealing spatial identity of these pools would require a 3D localization of the AZ with a fluorescence marker in each individual bouton and a simultaneous dual-color imaging of vesicle motion/tethering. Such measurements are currently difficult to perform because even a very small bleed through of the AZ signal to the vesicle channel increases the background noise and reduces localization accuracy. Furthermore, in the absence of a single labeling method that permits simultaneous monitoring of both vesicle motion and release, it is not currently feasible to unambiguously combine these two measurements for the same vesicle, and thus to directly link vesicle motion and tethering inside the synaptic boutons with the functional vesicle pool identity. Nevertheless, our findings provide the first step towards understanding of the mechanisms governing release site refilling and vesicle availability for release.

Materials and methods

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Reagent type
(species) or
resource
DesignationSource or
reference
IdentifiersAdditional
information
Chemical
compound,
drug
MyoVin-1 (Myo-1)EMD
Millipore
Cat. No.
475984
Chemical
compound,
drug
Pentabrom
opseudilin (PBP)
FISHER
SCIENTIFIC
Cat. No.
501015859
Chemical
compound,
drug
EGTA-AMFISHER
SCIENTIFIC
Cat. No. E1219
Chemical
compound,
drug
SGC5VWRCat. No. 89410–772
Chemical
compound,
drug
DL-AP5
Sodium salt
TocrisCat. No.
3693
Chemical
compound,
drug
CNQX disodium
salt
TocrisCat. No.
1045
Chemical
compound,
drug
HEPESSigmaCat. No.
H4034
Chemical
compound,
drug
D-(+)-GlucoseSigmaCat. No.
G 7021
Chemical
compound,
drug
Calcium
chloride
dihydrate
SigmaCat. No.
223506
Chemical
compound,
drug
Magnesium
chloride
hexahydrate
SigmaCat. No.
M9272
Chemical
compound,
drug
Minimum
Essential
Media (MEM) - No
Phenol Red
thermofisherCat. No.
51200–038
Chemical
compound,
drug
Characterized Fetal
Bovine Serum
hycloneCat. No.
SH30071.03
Chemical
compound,
drug
Penicillin-
Streptomycin
(5,000 U/mL)
thermofisherCat. No.
15070063
Chemical
compound,
drug
N-2 Supplement
(100X)
thermofisherCat. No.
17502048
Chemical
compound,
drug
Donor Equine
Serum
hycloneCat. No.
SH30074.03
Chemical
compound,
drug
Sodium Pyruvate
(100 mM)
thermofisherCat. No. 11360–070
Chemical
compound,
drug
Neurobasal-
A Medium
thermofisherCat. No. 10888–022
Chemical
compound,
drug
B-27 Supplement
(50X),
serum free
thermofisherCat. No. 17504–044
Chemical
compound,
drug
GlutaMAX
Supplement
thermofisherCat. No. 35050061
Chemical
compound,
drug
Earle’s
Balanced
Salts
sigmaaldrichCat. No. E3024
Chemical compound, drugCorning
Collagen I, Rat
Fisher
Scientific
Cat. No. 354236
Chemical
compound,
drug
Cover
Glasses
Fisher
Scientific
Cat. No. 12-545-80
Chemical compound, drugPapainWorthington
Biochemical
Cat. No. LS003126
Chemical compound, drugPDL (poly-D-lysine)BD Bios
ciences
Cat. No. 40210
Chemical compound, drugTrypsin-EDTA (0.05%), phenol redtherm
ofisher
Cat. No. 25300–054
Biological
sample
(Rattus norvegicus,
Female)
Sprague-Dawley
Timed-Pregnant rat,
E15 pups
Charles
River
Pups of both genders
Software,
algorithm
MATLABMathWorksRRID:SCR_001622
Software,
algorithm
u-track2.0Jaqaman,
K., et al.
(2008),
Nat.Meth.
5, 695–702
Gaudenz
Danuser Lab
Software,
algorithm
ImageJhttps://imagej.nih.gov/ij/RRID:SCR_003070
Software,
algorithm
Fijihttp://fiji.scRRID:SCR_002285
Software,
algorithm
inkscapeRRID:SCR_014479
 OtherVGluT1-
pHluorin
Drs. Robert
Edwards and
Susan Voglmaier
(UCSF)
Genetically
encoded optical
indicator of vesicle
release and recycling

Neuronal cell cultures

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Neuronal cultures were produced from the hippocampus of E16-17 rat pups of mixed gender as previously described (Peng et al., 2012). Hippocampi were dissected from E16-17 pups, dissociated by papain digestion, and plated on coated glass coverslips containing an astrocyte monolayer. Neurons were cultured in Neurobasal media supplemented with B27. All animal procedures conformed to the guidelines approved by the Washington University Animal Studies Committee (protocol approval # 20170233).

Lentiviral infection

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VGlut1-pHluorin was generously provided by Drs. Robert Edward and Susan Voglmaier (UCSF) (Voglmaier et al., 2006). Lentiviral vectors were generated by the Viral Vectors Core at Washington University. Hippocampal neuronal cultures were infected at DIV3.

Fluorescence microscopy

Neurotransmitter release measurements

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All experiments were conducted at 37°C within a whole-microscope incubator (In Vivo Scientific) at DIV16–19 as we described previously (Maschi and Klyachko, 2017). Neurons were perfused with bath solution (125 mM NaCl, 2.5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, 15 mM Glucose, 50 mM DL-AP5, 10 mM CNQX, pH adjusted to pH 7.4). Fluorescence was excited with a Lambda XL lamp (Sutter Instrument) through a 100 × 1.45 NA oil-immersion objective and captured with a cooled EMCCD camera (Hamamatsu). With this configuration the effective pixel size was 80 nm. Focal plane was continuously monitored, and focal drift was automatically adjusted with 10 nm accuracy by an automated feedback focus control system (Ludl Electronics). Field stimulation was performed by using a pair of platinum electrodes and controlled by the software via Master-9 stimulus generator (A.M.P.I.). Images were acquired using two frames with an acquisition time of 40 ms, one 45 ms before stimulation and one coincidently (0 ms delay) with stimulation. In some experiments (Figure 2A-C) imaging was performed using a cooled sCMOS camera (Hamamatsu). With this configuration, the effective pixel size was 60 nm and images were acquired at a frame rate of 200 ms.

Single-vesicle tracking

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Sparse vesicle labeling and functional synapse localization were performed following our previously developed procedures (Forte et al., 2017; Gramlich and Klyachko, 2017; Peng et al., 2012) The same bath solution as above was used for the dye loading and imaging, except 0.2 mM CaCl2, 1.0 mM MgCl2 were used to wash excess dye from the sample. 10 µM SGC5 (Biotium) were added to the bath solution for the dye loading step. Samples were imaged for 50 – 70 s, at an exposure rate of 80 msec (with a total frame rate of 10 Hz). Samples were stimulated for 10 s at 20 Hz with a 10 s delay after the first frame.

Pharmacology

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MyoVin-1 (Millipore), Pentabromopseudalin (PBP, Fisher Scientific) or EGTA-AM (Millipore) were diluted in DMSO (Sigma-Aldrich) and stored at −20°C. Samples were incubated in imaging solution with 30 µM Myo-1 for 5–10 min or 5 µM PBP for 5 min, or 250 µM EGTA-AM for 20 min before dye loading. The effective final DMSO concentration was <0.5%. Extended exposure to MyoVin-1 or PBP caused cell death, thus the bath solution during the experiment did not include Myo-1 or PBP. Our control measurements indicated that continuous presence of these blockers during the experiments did not have additional effects on vesicle motility beyond the effects of pre-incubation (data not shown).

Large-Area scanning electron microscopy (LaSEM)

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Cultures were fixed in a solution containing 2.5% glutaraldehyde and 2% paraformaldehyde in 0.15 M cacodylate buffer with 2 mM CaCl2, pH 7.4 that had been warmed to 37 ˚C for one hour. In experiments with KCl-induced depolarization, fixation was performed immediately following KCl application, and care was taken to complete the fixation procedure within a few seconds. The samples were then stained according the methods described by Deerinck et al., 2010. In brief, coverslips were rinsed in cacodylate buffer 3 times for 10 min each, and subjected to a secondary fixation for one hour in 2% osmium tetroxide/1.5% potassium ferrocyanide in cacodylate buffer for one hour, rinsed in ultrapure water 3 times for 10 min each, and stained in an aqueous solution of 1% thiocarbohydrazide for one hour. After this, the coverslips were once again stained in aqueous 2% osmium tetroxide for one hour, rinsed in ultrapure water 3 times for 10 min each, and stained overnight in 1% uranyl acetate at 4 ˚C. The samples were then again washed in ultrapure water 3 times for 10 min each and en bloc stained for 30 min with 20 mM lead aspartate at 60 ˚C. After staining was complete, coverslips were briefly washed in ultrapure water, dehydrated in a graded acetone series (50%, 70%, 90%, 100% x2) for 10 min in each step, and infiltrated with microwave assistance (Pelco BioWave Pro, Redding, CA) into Durcupan resin. Samples were flat embedded in a polypropylene petri dish and cured in an oven at 60 ˚C for 48 hr. Post resin curing, the coverslips were exposed with a razor blade and etched off with concentrated hydrofluoric acid. Small pieces of the resin containing the cells was then cut out by saw and mounted onto blank resin stubs before 70 nm thick sections were cut in the cell culture growing plane and placed onto a silicon wafer chips. These chips were then adhered to SEM pins with carbon adhesive tabs and large areas (~330×330 µm) were then imaged at high resolution in a FE-SEM (Zeiss Merlin, Oberkochen, Germany) using the ATLAS (Fibics, Ottawa, Canada) scan engine to tile large regions of interest. High-resolution tiles were captured at 16,384 × 16,384 pixels at 5 nm/pixel with a 5 µs dwell time and line average of 2. The SEM was operated at 8 KeV and 900 pA using the solid-state backscatter detector. Tiles were aligned and export using ATLAS 5.

Image and data analysis

Vesicle fusion analyses

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Fusion event localization

The fusion event localization at subpixel resolution was performed using MATLAB and the uTrack software package that was kindly made available by Dr. Gaudenz Danuser lab (Aguet et al., 2013; Jaqaman et al., 2008). The input parameters for the PSF were determined using stationary green fluorescent 40 nm beads. See (Maschi and Klyachko, 2017) for further details.

Whole synapse intensity quantification

Individual synapses were identified as fluorescence intensity peaks that increased in intensity upon stimulation using Matlab. Whole-synapse vGlut1-pHluorin intensity was measured over a region of interest (ROI) 10 pixels (600 nm) in radius from the center of each bouton. For a given stimulation frequency, five trains at 20 s intervals were applied. The data was pooled for the five consecutive trains in the same bouton because no difference between subsequent trains was observed (Figure 2—figure supplement 1F,G,H) and because the magnitude of changes measured using only the first train was indistinguishable from that averaged across trains (Figure 2—figure supplement 1G,H).

Hierarchical clustering analysis

Hierarchical clustering was performed using built-in functions in MATLAB as we described previously (Maschi and Klyachko, 2017).

Release site re-use probability analysis

a two-pulse stimulation paradigm was used as we described previously (Maschi and Klyachko, 2017). For each pair of stimuli, a subset of boutons was identified in which release events were detected for both stimuli in the pair; the probability of observing the two release events at the same release site was then calculated.

EM analyses

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Synapses identification for EM analysis

we used three characteristic features for manual synapse identification: the presence of a synaptic vesicle cluster, the postsynaptic density and the uniform synaptic gap between pre and postsynaptic membranes.

AZ to closest vesicle analysis of EM data

membrane opposite to the PSD was divided in small sections (0.5 nm) and the distance from each small section to the closer vesicle was determined using Matlab. This subset of membrane ‘closest’ vesicles was subsequently used to determine the relation between ‘docked’ and ‘tethered’ vesicle fractions. ‘Docked’ vesicles were defined as those with the distance from the membrane to the vesicle center less than 30 nm and ‘tethered’ vesicle as those with the distance less than 100 nm (Figure 3B).

Single-Vesicle detection and tracking

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Single-vesicle tracking

The feature identification and subpixel localization were performed using MATLAB and uTrack software package kindly provided by Dr. Gaudenz Danuser lab (Aguet et al., 2013; Jaqaman et al., 2008) following our previously developed procedures (Forte et al., 2017; Gramlich and Klyachko, 2017; Peng et al., 2012). Localization of functional synapses was performed using ImageJ. Quantification of vesicle motion was performed using the three-frame moving average of vesicle position to mitigate the effects of noise.

Integrated synapse intensity analysis

We quantified the total integrated intensity of a synapse containing a single labeled vesicle to determine changes in intensity after vesicle detection was lost. Analysis was performed in ImageJ. First, we drew a 10 × 10 pixel box (800 × 800 nm), corresponding to the typical size of a synapse (Schikorski and Stevens, 1997, Schikorski and Stevens, 1999), around the center of a vesicle at T = 0. Second, we summed the intensity of all pixels within the box for each frame in the movie (I(T)). Third, we normalized the data by the first frame intensity (I(T)/I(0)).

Vesicle detection with reduced stringency

To determine whether disappearing vesicles could be tracked longer with less stringent detection criteria, we loosened the full width at half max (FWHM) restriction on the width of the vesicle image by a factor of 2 (from 1.5 to 3 pixels). Consequently, we were able to track disappearing vesicles longer (Figure 4C and Figure 4—figure supplement 1E,F), and also track few vesicles that left their synapses and traveled to the axon (Figure 4—figure supplement 1C,D).

Identification of vesicles that exited from synapse

We determined if vesicle detection was lost because vesicles exited the synapse to the axon using definitions and analysis we described previously (Gramlich and Klyachko, 2017) (Figure 4—figure supplement 1A,B). To quantify the change in integrated synapse intensity upon vesicle exit to the axon, we used tracks of the vesicles known to exit a synapse without loss of detection: Figure 4—figure supplement 1A,B shows the intensities of 4 different synapses with apparent vesicle exit, aligned relative to vesicle exit, so that T = 0 is defined as the time a vesicle is >600 nm from its position in the first frame (Gramlich and Klyachko, 2017). The intensity remains constant before exit followed by a 20–30% reduction after the vesicle exits the synapse. The rate at which the intensity drops depends upon the speed of the vesicle as it exits the synapse, with faster vesicles exhibiting a faster drop in intensity. Thus we used the average curve (black line in Figure 4—figure supplement 1B) as the baseline to determine if loss of detection was due to vesicle exit from the synapse to the axon. We found three vesicles which disappearance was consistent with the exit to the axon.

Vesicle disappearance and appearance analysis

The subgroup of vesicle tracks used in the disappearance and appearance rate analysis was determined as follows:

  1. All tracks that started at T = 0, were within 600 nm of a synapse and were tracked longer than 50 frames were chosen for disappearance group (S1).

  2. Any synapse that had more than one track at T = 0 was excluded.

  3. Any track that traveled more than 800 nm from the center of a synapse during observation was excluded.

  4. Tracks that appear in a synapse were chosen for the appearance group (S2) only if another track disappeared in the same synapse at an early time point.

  5. Appearing tracks were a minimum of 20 frames in duration.

  6. Appearing tracks were not chosen if they began outside of the synapse.

The rate of disappearance was calculated as the cumulative fraction of vesicles which detection was lost in group S1 as a function of time. The number of lost tracks per time (N1(t)) were calculated. Lost tracking was calculated as the number of tracks reporting ‘NaN’ during tracking. Some tracks have gaps, or missing frames reported as ‘NaN,’ which results in variance in N1(t). The total lost tracks per frame was then divided by the total number of tracks in the group:

F1(t)=N1(t)|S1|

The rate of appearance was determined as the cumulative fraction of appearances in group S2 as a function of time. At each frame (t), the total number of new track appearances (N2(t)) were counted and added the total number of appearances per time (F2(t)). If a track appeared more than once following another track in the same synapse, then the new appearance was also counted in the cumulative total. Thus, the appearance rate measures the fraction of appearances as a function of time:

F1(t)=1|S2|i=0tN2(t)

Error-bars on disappearance and appearance rates were determined from averages of their best fit residuals. Disappearance and Appearance rates were fit to exponential and linear functions, respectively. The difference from the best fit line and raw data was then determined. The average of those differences were then reported with the value for each condition (Figure 5 C,E).

Vesicle disappearance and appearance oversampling correction

Vesicle disappearance and appearance distributions were sampled at a rate of 10 frames per second. However, the typical disappearance rate was on the order of 1 vesicle per second (1 vesicle per 10 frames) resulting in significant oversampling. Thus, we averaged the oversampled distributions with a five-frame moving average and plotted every fifth data point. Further, we performed statistical analysis on the averaged data to prevent over-sampling bias of the statistics.

Statistical analyses

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Statistical analyses were performed in Matlab. Statistical significance was determined using two tailed Student's t-test, Kolmogorov-Smirnov (K-S) test, or chi-squared test where appropriate. Data is reported as mean ±SEM or±Residual from fits to data, as indicated in the text.

Appendix 1

Computational model of vesicle disappearance

We computationally simulate vesicle disappearance rates following a previously published dynamic Monte-Carlo model (Gramlich and Klyachko, 2017), using the python 3.3 language. First, we defined the following variables:

Pδ,0Baseline disappearance probability
Pδ,SDisappearance probability during stimulation
T0Time of vesicle disappearance
TSTime stimulation starts
NtotTotal number of vesicles simulated
FlabeledFraction of Ntot vesicles labeled at T0

Second, at the beginning of each time-step, we obtained two random numbers (R1, R2) between [0,1] using the python numpy random number generator random.rand(). Third, if R1 is less than Pδ,0/S and R2 is less than the fraction of current labeled vesicles then a labeled vesicle is chosen to be removed and replaced with an un-labeled vesicle; otherwise an un-labeled vesicle is chosen to be replaced with another un-labeled vesicle.

The fraction of labeled vesicles remaining in the simulation was plotted as a function of time-steps. The fraction of simulated vesicles that disappeared was then compared to the experimentally observed fractions in (Figure 5B). When best-fit probability was found that matched experimentally observed data, the simulated δ-rate was calculated:

δ=ProbabilityofvesicledisappearanceperframeNumberofsimulatedvesicles=pδNTotal

Calculation of the initial fraction of vesicles in Pools I and II.

To calculate the fraction of vesicles in each pool at T = 0 based on the effect of EGTA-AM on the basal disappearance rate the following definitions/assumptions were made:

  1. The true disappearance rate (from Pool I) is (δCT)

  2. The observed rate of disappearance in the presence of EGTA-AM is (δEGTA), the observed rate of appearance is (αEGTA).

  3. ρ is a fraction of labeled vesicles in Pool I; thus 1- ρ is a fraction of labeled vesicles in Pool II.

  4. Observed changes in disappearance rate are due to contribution from vesicles in Pool II.

  5. The change in observed rate of disappearance in the presence of EGTA-AM (δEGTA) is determined by a combination of a vesicle fraction in Pool I (ρ) and a contribution from the vesicle fraction in Pool II (1- ρ).

These assumptions can be combined in an equation:

δEGTA=ρδCT+(1ρ)αEGTA

This equation results in the solution:

ρ=δEGTAαEGTAδCTαEGTA

Using the range of values observed experimentally, we obtained:

δEGTA = 0.038 ± 0.006 fraction per second;

αEGTA = 0.014 ± 0.005 fraction per second;

δCT = 0.047 ± 0.017 fraction per second;

ρ = 74.5 ± 11%

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Decision letter

  1. Yukiko Goda
    Reviewing Editor; RIKEN, Japan
  2. Richard Aldrich
    Senior Editor; The University of Texas at Austin, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Myosin V functions as a vesicle tether at the plasma membrane to control neurotransmitter release in central synapses" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Richard Aldrich as the Senior Editor. The following individual involved in review of your submission has agreed to reveal his identity: Alain Marty (Reviewer #3).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

Here Maschi et al. examine how active zone release sites are replenished with new vesicles following exocytosis, specifically focusing on the role for myosin V in this process, using nano-scale imaging to track the behavior of single vesicles. They identify that vesicle replenishment is sensitive to myosin V inhibition and provide evidence that vesicle undocking from the plasma membrane – i.e. the rate of vesicle exit from the field of view – is suppressed by activity, which in turn, helps strengthen the tether in a myosin V-dependent manner. The study is timely and imaginative, and it reports a series of provocative new observations. Methods are appropriate, analysis and modeling are generally well done (except for some caveats below), and the writing is clear. Overall, the study should be of general interest to the readership of eLife. However, in its present form, the paper has two major issues.

Firstly, myosin V inhibitor is shown to have effects on the measurements of single release site exocytosis (Figure 1), activity-dependent vesicle recruitment to the release site (Figure 2), and single vesicle tracking inside the presynaptic terminal (Figure 3-4). The authors present the case that these effects are related to each other. However, there remains a possibility that such effects represent distinct actions of myosin V that are based on interactions with different molecular processes, similarly to a role for myosin V in inter-bouton trafficking of vesicles that have been previously reported by the Klyachko group. For example, the results in Figure 1-2 suggest a very local action of myosin whereas the results of Figure 3-4 suggest a longer range action. Secondly, the analysis of vesicular movement based on standard deviation, and the accompanying double pool model contains several weakness that require careful consideration. The specific points relating to the concerns are outlined below. The reviewers have agreed that these concerns are addressable, in principle, by additional experiments, re-analysis of data, and re-writing of the text over a two-month period.

Essential revisions:

1) Figure 2: The two pool model involving tethered vesicles and vesicles remote from the docked pool (inner recycling pool or reserve pool) is not necessarily compatible with myosin V inhibitor-dependent spatial distribution of release site re-use (Figure 2D-F), and this point requires further clarifications. What is the relationship between myosin V-dependent tethering and the location of release sites with respect to VGCCs responsible for the calcium influx that drives exocytosis? An attractive interpretation of the results in Figure 2D-E would be that myosin V is involved in endocytosis. By inhibiting endocytosis, Myo1 would then enhance the size of the AZ in an activity dependent manner, thus increasing the distance of peripheral release sites to the center.

2) Figure 2A-C: A control should be added showing the (lack of?) effects of a previous stimulation in control conditions, without addition of Myo1.

3) Figure 2H-I: The control with Myo1 but no KCl is missing.

4) Figure 3 and 4: Orientation of the synapse with respect to the imaging plane. The analysis makes the implicit assumption that the AZ is orthogonal to the Z-axis. In Figure 1, the argument can be made that AZs displaying several release sites must be mainly orthogonal to the Z-axis, introducing a sample bias in favor of synapses displaying a horizontal orientation. With the protocol of Figure 3-4 no such argument can be made, so that a change in the Z-axis may represent a movement parallel to the AZ plane. This challenges the identification of pool I and pool II as representing two sets of SVs positioned at different distances of the AZ membrane.

In addition to the assumption of a horizontal orientation, the analysis implicitly assumes that the imaged SV is initially in pool I, near the AZ membrane. Since pool II is larger than pool I, it would seem more likely that it would be initially in pool II, away from the AZ membrane. One way out of this conundrum may be to assume an experimental bias towards pool I by selection of slowly moving vesicles.

5) Figure 3 and 4: Lack of calibration of σ changes. The main argument linking disappearance to z movement is that σ increases. However the extent of z movement associated with the observed change in σ remains unclear. In view of the results of Figure 2 the z movement is expected to take place in a range of 10s of nm, but since the size of the point spread function is on the order of 1 micron in the Z-axis, the z movements corresponding to the observed σ changes are likely to be larger, in a range of 100s of nm. This issue could perhaps be settled by measuring σ values as a function of z for a fluorescent bead of small size.

6) Figure 3 and 4: Interpretation of the σ change. The proposed change in z position is one possibility, but an alternative interpretation is that the SGC5 fluorescence bleaches more rapidly than the background fluorescence. To distinguish between these possibilities, it would be good to see whether σ returns near its initial value when the SV reenters pool I (e.g. in Figure 3C).

7) Figure 3: The data in Figure 3B suggest a continuous drift. By contrast, the scheme of Figure 3D suggests abrupt changes with probabilities linked to α and δ. A possible explanation for the drift could be a movement of the preparation with respect to the imaging plane. Yet another interpretation would be a diffusion process, without the intervention of distinct vesicular pools. This point needs to be clarified.

8) Figure 4: Regarding EGTA's effect on disappearance and appearance rates, in Figure 4B it seems like EGTA has a role in baseline disappearance rate (although it appears to be non-significant, the extent of reduction comparing +Ctrl and +EGTA in Figure 4B-Basal looks similar to the level of change shown in Figure 4D-Activity for the same groups, the significance seems to be a consequence of reduced dispersion in 4D compared to 4B). Would this mean that calcium regulates the stability/immobility of docked synaptic vesicles in basal conditions?

9) The plots showing spatial displacement of vesicles in Figure 4E are very straightforward and allow a fast understanding for the reader. The authors should include the same plot for inhibition of Myosin V (the data is only presented in Table 1), to emphasize the result and make the conclusion easier to understand.

10) Although not mentioned in the present study, the undocking of vesicles was previously characterized by Murthy and Stevens (Murthy and Stevens, 1999) in hippocampal synapses, who reported that undocking was three times more prevalent than the occurrence of spontaneous exocytosis, similarly to the present findings.

11) Figure 1: The method used to obtain the control data in the upper panel of Figure 1B should be explicitly stated. Likewise, the method used to generate the plot in Figure 1C should be explained.

12) Figure 2: Here the main issue is time. The reloading mechanism discussed in Figure 1 and Figure 2A-F operates on a time scale of a few seconds. In Figure 2H-I however, a prolonged KCl stimulation (10 min) is applied. Notably, this might be harsh for the cultured neurons, and the authors should state the concentration of KCl used. Furthermore, the time between the end of stimulation and fixation is not specified, and this time is presumably again on the order of minutes. One day the experiments of Figure 2H-I will need to be repeated with the better time resolution of the flash and freeze technique. This is not needed for this paper, but the protocols used for EM experiments should be more precisely outlined, and caution should be applied when discussing the links between EM and single vesicle exocytosis results.

https://doi.org/10.7554/eLife.39440.016

Author response

Essential revisions:

1) Figure 2: The two pool model involving tethered vesicles and vesicles remote from the docked pool (inner recycling pool or reserve pool) is not necessarily compatible with myosin V inhibitor-dependent spatial distribution of release site re-use (Figure 2D-F), and this point requires further clarifications. What is the relationship between myosin V-dependent tethering and the location of release sites with respect to VGCCs responsible for the calcium influx that drives exocytosis? An attractive interpretation of the results in Figure 2D-E would be that myosin V is involved in endocytosis. By inhibiting endocytosis, Myo1 would then enhance the size of the AZ in an activity dependent manner, thus increasing the distance of peripheral release sites to the center.

We thank the reviewers for this important point and completely agree. We performed a series of analyses to examine a possibility that Myosin V is involved in endocytosis:

1) We directly examined the reviewers’ suggestion what inhibition of myosin V could increase the size of the AZ in an activity-dependent manner thus increasing the distance of peripheral release sites to the center. Our scanning EM analysis showed that AZ size remained unchanged by myosin V inhibition both at baseline (Figure 1E) and during KCl-induced depolarization(Figure 2—figure supplement 1B). This result suggests that inhibition of myosin V does not cause activity-dependent changes in AZ size.

2) We examined if inhibition of myosin V affects dynamics of endocytosis in live and active synapses. The decay of vGlut1-pHluorin signal following a stimulus train is determined by endocytosis and subsequent vesicle reacidification. Changes in the decay of the vGlut1-pHluorin signal in our measurements can thus be interpreted to reflect predominantly changes in endocytosis (Atluri and Ryan, 2006). We found that inhibition of myosin V had no measurable effect on the decay of the vGlut1-pHluorin signal following 50Hz trains (Figure 2—figure supplement 1C-E).

3) If inhibition of myosin V blocks endocytosis, application of sequential stimulus trains would be expected to cause surface accumulation of VGlut1-pHluorin and a corresponding increase in the bouton fluorescence. We compared the amplitudes of VGlut1-pHluorin signal in five consecutive 50Hz trains separated by 20sec each, and did not observe any measurable changes from one train to the next under control conditions or in the presence of myosin V inhibitors (Figure 2—figure supplement 1F, G, H). While an indirect evidence, this result supports the above findings that acute inhibition of myosin V does not significantly affect vesicle endocytosis under our experimental conditions.

Combined, these results indicate that the shift in spatial localization of release events upon myosin V inhibition is not caused by an increase in the AZ size and is unlikely to be mediated by altered endocytosis. These new findings are now presented in the second paragraph of the Results subsection “Myosin V controls release site refilling during high-frequency stimulation”.

We also note that the spatial shift in the location of release Events upon myosin V inhibition was observed not only during stimulus trains but also at low frequency stimulation (Figure 1F). Under these conditions, release is very infrequent and no changes in the size of the AZ are observed (Figure 1E). The observed shift in the distribution of release events upon myosin V inhibition could thus arise without changes in the physical localization of release sites. This effect could arise, for example, from a reduced release capacity of individual release sites we observed, leading to a more spatially distributed release. Indeed, we previously described a comparable phenomenon of reduced release site re-use capacity and a shift in the distance of release events towards AZ periphery during high-frequency stimulation, and this effect was observed without myosin V inhibition (Maschi and Klyachko, 2017). While we cannot exclude the possibility that the spatial and temporal effects of myosin V inhibition arise from distinct myosin V functions, the above considerations and the parallels with the effects of activity point to a common myosin V function as the simplest and the most logical interpretation of our findings. We have expanded the Discussion of the revised manuscript to include these considerations as follows:

“Our analyses suggest that this spatial shift is unlikely to arise from myosin V involvement in endocytosis. […] It remains to be determined whether the spatial effect of myosin V inhibition is a reflection of myosin V’s function in vesicle tethering or relies on an independent mechanism”.

2) Figure 2A-C: A control should be added showing the (lack of?) effects of a previous stimulation in control conditions, without addition of Myo1.

We thank the reviewers for this point and have performed the suggested analysis in two different and complementary ways:

1) We examined responses to 5 consecutive trains of 20 stimuli at 50Hz used in Figure 2A-B and calculated changes in synaptic responses from one train to the next. We found that the difference between subsequent responses in control conditions was negligible (~2%) (Figure 2—figure supplement 1F). Moreover, the change in response from one train to the next was also indistinguishable in control conditions vs. in the presence of myosin V inhibitors (Figure 2—figure supplement 1F).

2) For each of the 5 consecutive stimulus trains, we plotted the magnitude of changes in synaptic response in the presence of myosin V inhibitors relative to control, and did not observe any measurable differences from one train to another (Figure 2—figure supplement 1G, H).

Finally, we note that the magnitude of changes measured using only the first train was indistinguishable from that averaged across trains (Figure 2—figure supplement 1G, H).

Together these analyses provide strong evidence for lack of measurable effects of previous stimulation in our measurements.

We also realized that the measurements in Figure 2C were not described clearly. Measurements for each frequency were performed in a separate set of dishes and normalized to its own control. Therefore there is no effect of previous stimulation in these experiments carried from one frequency to another. The n values reported in the Table 1 represent the total number of dishes for all frequencies combined, since this entire dataset was used to perform the linear regression analysis shown in Figure 2C. We have clarified the description of this experiment in the Results and the Figure 2 legend.

3) Figure 2H-I: The control with Myo1 but no KCl is missing.

We thank the reviewers for this point and note that this control was actually performed (Figure 3—figure supplement 1A, B), but it was not described clearly. We have improved the description of this experiment in Results subsection “Inhibition of myosin V causes a vesicle docking defect during sustained activity”, to clarify this point.

4) Figure 3 and 4: Orientation of the synapse with respect to the imaging plane. The analysis makes the implicit assumption that the AZ is orthogonal to the Z-axis. In Figure 1, the argument can be made that AZs displaying several release sites must be mainly orthogonal to the Z-axis, introducing a sample bias in favor of synapses displaying a horizontal orientation. With the protocol of Figure 3-4 no such argument can be made, so that a change in the Z-axis may represent a movement parallel to the AZ plane. This challenges the identification of pool I and pool II as representing two sets of SVs positioned at different distances of the AZ membrane.

We thank the reviewers for highlighting this limitation of our model. Given the previous literature on calcium channel distribution and mobility within the AZ (Schneider et al., 2015), we believe the simplest model that accounts for our observations is two pools at different distances from the AZ, for the following reasons:

1) The different exchange rates between the two pools suggest that they are different in size and dynamics.

2) Observation that disappearance and appearance rates are differentially affected by EGTA suggest that the two pools are at different distances from the AZ’s calcium channels.

3) Differential activity-dependence of the exchange rates between the two pools supports the notion that these pools have a different distance to the source of calcium influx, the AZ.

We agree with the reviewers that our vesicle tracking measurements do not distinguish whether vesicles travel along or perpendicular to the AZ plane. The effect of EGTA could thus in principle be also explained by two vesicle pools both localized at the AZ, but at a systematically distinct distance to the source of calcium influx, i.e. calcium channels. However, this model is unlikely for two reasons: (i) a large proportion (~60%) of the calcium channels are mobile in the AZ plane with the median surface area explored by individual channel molecules of ~200-250 nm (Schneider et al., 2015) comparable to dimensions of the entire AZ (Schikorski and Stevens, 1997); (ii) the large spatial domain of vesicle motion that we observed in our tracking experiments, which commonly spans several hundred nanometers over our observation period and includes epochs of fast diffusion and directed motion (Forte et al., 2017); such extensive vesicle mobility is difficult to reconcile with the motion restricted along the AZ given the very limited AZ dimensions and the vesicle crowding at the AZ which unlikely to allow fast diffusion or directed motion.

To address the reviewers’ point we now discuss this limitation of our model and these considerations in the last paragraph of the Discussion subsection “A myosin V- and activity-dependent reversible vesicle tethering in release site refilling”.

In addition to the assumption of a horizontal orientation, the analysis implicitly assumes that the imaged SV is initially in pool I, near the AZ membrane. Since pool II is larger than pool I, it would seem more likely that it would be initially in pool II, away from the AZ membrane. One way out of this conundrum may be to assume an experimental bias towards pool I by selection of slowly moving vesicles.

We thank the reviewers for this point and agree. Because disappearance is the only criteria for the vesicle inclusion in our initial analysis and due to the arbitrary relationship between the focal plane and the AZ at any given synapse, there is no a priori information that would allow us to make the initial assignment of the vesicles to Pool I or Pool II. As a result, our measurements of the disappearance rate reflect a mixture of vesicles that are initially in Pool I and Pool II at the beginning of observation. We considered the reviewers’ suggestion to assume bias towards Pool I by selection of slowly moving vesicles to address this issue. However, we found that it was not possible to group vesicles based on their speeds in any systematic manner because vesicle speeds are highly variable during the observation time due to multiple intertwining epochs of diffusion, directed motion and pausing with speeds of different epochs varying several fold along the individual vesicle trajectories (Forte et al., 2017).

However, our finding that EGTA strongly affects only the vesicle appearance rate (and thus vesicles in Pool II) together with the finding that EGTA also has a small effect on the baseline disappearance rate (which we interpreted within our model as a contribution from vesicles in Pool II) provides a simple way to estimate the ratio of vesicles that start in each pool in beginning of our observations. To perform this analysis, we assumed that only the vesicle appearance rate is truly affected by EGTA (based on our observation that disappearance rate for EGTA and Ctrl during stimulation are the same), while the small effect of EGTA on the baseline disappearance rate (vesicles in Pool I) is caused by contribution from Pool II vesicles. Within these assumptions, the model (Appendix) and EGTA results suggest that among all vesicles assumed to be in Pool I in the beginning of observations, 75 ± 11% actually start in Pool I, while 25 ± 11% are physically located in Pool II. This result is consistent with the Pool II being larger, and having a slower exit rate, because the probability that a given labeled single vesicle in our experiment will leave the Pool II is smaller than if it would be in Pool I and thus less likely to be included in our disappearance analysis. These considerations support our model by demonstrating that the majority of the disappearing vesicles are initially located in Pool I. We have added these considerations to the third paragraph of the Results subsection “Vesicle shuttling represents transitions between a membrane pool and an internal pool”.

5) Figure 3 and 4: Lack of calibration of σ changes. The main argument linking disappearance to z movement is that σ increases. However the extent of z movement associated with the observed change in σ remains unclear. In view of the results of Figure 2 the z movement is expected to take place in a range of 10s of nm, but since the size of the point spread function is on the order of 1 micron in the Z-axis, the z movements corresponding to the observed σ changes are likely to be larger, in a range of 100s of nm. This issue could perhaps be settled by measuring σ values as a function of z for a fluorescent bead of small size.

We thank the reviewers for suggesting the σ calibration experiments and have performed these measurements (Figure 4—figure supplement 1G). This calibration allows us to correlate the observed range of vesicle σ values (~120-150 nm) with the range of detectable vesicle motion along the Z-axis of about 100 nm (Figure 4—figure supplement 1G). However, we also note that this concern likely arose from lack of clarity in our description of the results in Figure 2, rather than a problem with the results themselves. The reviewers referred to the results in Figure 2 to estimate relevant distances for the expected vesicle Z-movements between the two pools. However, these measurements in Figure 2 represent changes in the vesicle distance to the AZ upon myosin V inhibition, but do not represent distances in vesicle localization in Pools I and II under normal conditions. Therefore our data in Figure 2 does not provide any estimates for how far apart the pools are expected to be. We also note that only the σ changes in XY plane are used as one of the detection criteria, with the typical σ value of ~120nm. Most importantly, the actual numerical changes in σ values are not used in any of the analyses in the manuscript. We have clarified these points in the fifth paragraph of the Results subsection “Tracking individual synaptic vesicles during recycling supports a model of continuous 238 vesicle shuttling between two vesicle pool”.

6) Figure 3 and 4: Interpretation of the σ change. The proposed change in z position is one possibility, but an alternative interpretation is that the SGC5 fluorescence bleaches more rapidly than the background fluorescence. To distinguish between these possibilities, it would be good to see whether σ returns near its initial value when the SV reenters pool I (e.g. in Figure 3C).

We thank the reviewers for this point and have performed both of the suggested analyses.

First, we performed the analysis of bleaching and showed that the σ changes were not caused by differential bleaching of vesicle signal vs. background (Figure 4—figure supplement 1I).

Second, we compared the σ changes just before vesicle disappearance and right after reappearance (Figure 4—figure supplement 1H). σ values increased just before disappearance, and then decreased just after re-appearance.

Together these results support our interpretation that vesicle going out of focus due to changes in zposition is the predominant cause of vesicle disappearance/loss of detection.

7) Figure 3: The data in Figure 3B suggest a continuous drift. By contrast, the scheme of Figure 3D suggests abrupt changes with probabilities linked to α and δ. A possible explanation for the drift could be a movement of the preparation with respect to the imaging plane. Yet another interpretation would be a diffusion process, without the intervention of distinct vesicular pools. This point needs to be clarified.

We appreciate the reviewers’ point and we believe it is based on a misnomer we used in the figure description: Y-axis in the original Figure 3B was labeled ∆X, but this represents the localization error of the vesicle position (which can increase with vesicle moving away from the focal plane) and not the actual position itself and therefore does not show drift. We performed extensive controls for the drift in our imaging system and found that the drift of the preparation in our measurements is extremely small (6-8nm) (Maschi and Klyachko, 2017; Forte et al., 2017) and is below the localization uncertainty (~15-20nm). We apologize for the confusion in the labeling of original Figure 3B and have re-labeled the figure axis (the new Figure 4B and 4E) as “localization error” to avoid confusion.

8) Figure 4: Regarding EGTA's effect on disappearance and appearance rates, in Figure 4B it seems like EGTA has a role in baseline disappearance rate (although it appears to be non-significant, the extent of reduction comparing +Ctrl and +EGTA in Figure 4B-Basal looks similar to the level of change shown in Figure 4D-Activity for the same groups, the significance seems to be a consequence of reduced dispersion in 4D compared to 4B). Would this mean that calcium regulates the stability/immobility of docked synaptic vesicles in basal conditions?

We appreciate the reviewers’ point. Our measurements of the disappearance rate reflect a mixture of vesicles that are in Pool I and Pool II at the beginning of observation. Because EGTA only strongly affects the appearance rate (and thus vesicles in Pool II), we interpreted the effect of EGTA on baseline disappearance rate to reflect a mixture of vesicles that are in Pool I and Pool II at the beginning of observation. We estimated the ratio of vesicles that start in each pool (as we described in response to point 4 above), assuming that only the vesicle appearance rate is truly affected by EGTA (based on our observation that disappearance rate for EGTA and Ctrl during stimulation are the same), while the small effect of EGTA on the disappearance rate (vesicles in Pool I) is caused by contribution from Pool II vesicles. Within these assumptions, our results suggest that among all vesicles assumed to be in Pool I in the beginning of observations, 75 ± 11% actually start in Pool I, while 25 ± 11% are physically located in Pool II. As we pointed out in our response to the concern #4 above, this result is consistent with the Pool II being larger, and having a slower exit rate, because the probability that a given labeled single vesicle in our experiment will leave the Pool II is smaller than if it would be in Pool I and thus less likely to be included in our disappearance analysis. We have added these considerations to the third paragraph of the Results subsection “Vesicle shuttling represents transitions between a membrane pool and an internal pool” and Appendix.

9) The plots showing spatial displacement of vesicles in Figure 4E are very straightforward and allow a fast understanding for the reader. The authors should include the same plot for inhibition of Myosin V (the data is only presented in Table 1), to emphasize the result and make the conclusion easier to understand.

We thank the reviewers for this suggestion and have included the same plot for inhibition of myosin V accordingly (Figure 6B, C).

10) Although not mentioned in the present study, the undocking of vesicles was previously characterized by Murthy and Stevens (Murthy and Stevens, 1999) in hippocampal synapses, who reported that undocking was three times more prevalent than the occurrence of spontaneous exocytosis, similarly to the present findings.

We thank the reviewers for bringing this oversight to our attention. We have added the citation and corresponding findings to the first paragraph of the Discussion subsection “A myosin V- and activity-dependent reversible vesicle tethering in release site refilling”.

11) Figure 1: The method used to obtain the control data in the upper panel of Figure 1B should be explicitly stated. Likewise, the method used to generate the plot in Figure 1C should be explained.

We thank the reviewers for this suggestion and have expanded the description of the methods in the Results and in the Materials and methods subsections “Whole synapse intensity quantification” and “Release site re-use probability analysis” accordingly.

12) Figure 2: Here the main issue is time. The reloading mechanism discussed in Figure 1 and Figure 2A-F operates on a time scale of a few seconds. In Figure 2H-I however, a prolonged KCl stimulation (10 min) is applied. Notably, this might be harsh for the cultured neurons, and the authors should state the concentration of KCl used. Furthermore, the time between the end of stimulation and fixation is not specified, and this time is presumably again on the order of minutes. One day the experiments of Figure 2H-I will need to be repeated with the better time resolution of the flash and freeze technique. This is not needed for this paper, but the protocols used for EM experiments should be more precisely outlined, and caution should be applied when discussing the links between EM and single vesicle exocytosis results.

We appreciate the reviewers’ point and have expanded and clarify the protocols for the EM experiments and stated the concentration of KCl (55 mM). We also clarified that the fixation was performed immediately following KCl application with only a few second delay. Finally, we avoided making strong conclusions based on the links between the EM and single-vesicle imaging data.

https://doi.org/10.7554/eLife.39440.017

Article and author information

Author details

  1. Dario Maschi

    1. Department of Cell Biology and Physiology, Washington University, Missouri, United States
    2. Department of Biomedical Engineering, Washington University, Missouri, United States
    Contribution
    Software, Formal analysis, Investigation, Methodology
    Contributed equally with
    Michael W Gramlich
    Competing interests
    No competing interests declared
  2. Michael W Gramlich

    1. Department of Cell Biology and Physiology, Washington University, Missouri, United States
    2. Department of Biomedical Engineering, Washington University, Missouri, United States
    Contribution
    Software, Investigation, Methodology
    Contributed equally with
    Dario Maschi
    Competing interests
    No competing interests declared
  3. Vitaly A Klyachko

    1. Department of Cell Biology and Physiology, Washington University, Missouri, United States
    2. Department of Biomedical Engineering, Washington University, Missouri, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Methodology, Writing—original draft, Project administration
    For correspondence
    klyachko@wustl.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3449-243X

Funding

CIMED Center at Washington University

  • Vitaly A Klyachko

National Institute of Neurological Disorders and Stroke (NS105776)

  • Vitaly A Klyachko

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work was supported in part by grants to VAK from NINDS (RO1 NS105776) and CIMED center at Washington University. We acknowledge the assistance of Matthew Joens and Dr. James Fitzpatrick at the Washington University Center for Cellular Imaging in EM studies.

Ethics

Animal experimentation: All animal procedures were in compliance with the US National Institutes of Health Guide for the Care and Use of Laboratory Animals. All animal procedures conformed to the guidelines approved by the Washington University Animal Studies Committee (protocol approval # 20170233).

Senior Editor

  1. Richard Aldrich, The University of Texas at Austin, United States

Reviewing Editor

  1. Yukiko Goda, RIKEN, Japan

Publication history

  1. Received: June 23, 2018
  2. Accepted: October 11, 2018
  3. Accepted Manuscript published: October 15, 2018 (version 1)
  4. Version of Record published: October 31, 2018 (version 2)

Copyright

© 2018, Maschi et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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