Skip to main content
NCBI home page
The Journal of Physiology logoLink to The Journal of Physiology
. 2016 Jun 27;595(1):193–206. doi: 10.1113/JP271937

Dynamin‐1 deletion enhances post‐tetanic potentiation and quantal size after tetanic stimulation at the calyx of Held

Satyajit Mahapatra 1, Xuelin Lou 1,✉
PMCID: PMC5199734  PMID: 27229184

Abstract

Key points

  • Post‐tetanic potentiation (PTP) is attributed mainly to an increase in release probability (P r) and/or readily‐releasable pool (RRP) in many synapses, but the role of endocytosis in PTP is unknown.

  • Using the calyx of Held synapse from tissue‐specific dynamin‐1 knockout (cKO) mice (P16–20), we report that cKO synapses show enhanced PTP compared to control.

  • We found significant increases in both spontaneous excitatory postsynaptic current (spEPSC) amplitude and RRP size (estimated by a train of 30 APs at 100 Hz) in cKO over control during PTP.

  • Actin depolymerization blocks the increase in spEPSC amplitude in both control and cKO, and it abolishes the enhancement of PTP in cKO. PTP is sensitive to the PKC inhibitor GF109203X in both control and cKO.

  • We conclude that an activity‐dependent quantal size increase contributes to the enhancement of PTP in cKO over control and an altered endocytosis affects short‐term plasticity through quantal size changes.

Abstract

High‐frequency stimulation leads to post‐tetanic potentiation (PTP) at many types of synapses. Previous studies suggest that PTP results primarily from a protein kinase C (PKC)‐dependent increase in release probability (P r) and/or readily‐releasable pool (RRP) of synaptic vesicles (SVs), but the role of SV endocytosis in PTP is unknown. Using the mature calyx of Held (P16–20), we report that tissue‐specific ablation of dynamin‐1 (cKO), an endocytic protein crucial for SV regeneration, enhances PTP in cKO over control. To explore the mechanism of this enhancement, we estimated the changes in paired‐pulse ratios (PPRs) and RRP size during PTP. RRP was estimated by the back‐extrapolation of cumulative EPSC amplitudes during a train of 30 action potentials at 100 Hz (termed RRPtrain). We found an increase in RRPtrain during PTP in both control and cKO, but no significant changes in the PPR. Moreover, the amplitude and frequency of spontaneous excitatory postsynaptic currents (spEPSCs) increased during PTP in both control and cKO; however, the spEPSC amplitude in cKO during PTP was significantly larger than in control. Actin depolymerization reagent latrunculin‐B (Lat‐B) abolished the activity‐dependent increase in spEPSC amplitude in both control and cKO, but selectively blocked the enhancement of PTP in cKO, without affecting PTP in control. PKC inhibitor GF109203X nearly abolished PTP in both control and cKO. These data suggest that the quantal size increase contributes to the enhancement of PTP in dynamin‐1 cKO, and this change depends on strong synaptic activity and actin polymerization.

Key points

  • Post‐tetanic potentiation (PTP) is attributed mainly to an increase in release probability (P r) and/or readily‐releasable pool (RRP) in many synapses, but the role of endocytosis in PTP is unknown.

  • Using the calyx of Held synapse from tissue‐specific dynamin‐1 knockout (cKO) mice (P16–20), we report that cKO synapses show enhanced PTP compared to control.

  • We found significant increases in both spontaneous excitatory postsynaptic current (spEPSC) amplitude and RRP size (estimated by a train of 30 APs at 100 Hz) in cKO over control during PTP.

  • Actin depolymerization blocks the increase in spEPSC amplitude in both control and cKO, and it abolishes the enhancement of PTP in cKO. PTP is sensitive to the PKC inhibitor GF109203X in both control and cKO.

  • We conclude that an activity‐dependent quantal size increase contributes to the enhancement of PTP in cKO over control and an altered endocytosis affects short‐term plasticity through quantal size changes.

Abbreviations

AP

action potential

CH

calyx of Held

cKO

conditional dynamin‐1 knockout

CTZ

cyclothiazide

GF

GF109203X

KYN

kynurenic acid

Lat‐B

latrunculin‐B

PKC

protein kinase C

PPR

paired‐pulse ratio

PTP

post‐tetanic potentiation

Pr

release probability

RRP

readily releasable pool

RRPtrain

RRP estimated by a train of 30 APs at 100 Hz

spEPSCs

spontaneous excitatory postsynaptic currents

SV

synaptic vesicle

Introduction

Post‐tetanic potentiation (PTP) is a common form of short‐term synaptic plasticity involving a transient increase in synaptic transmission induced by a brief high‐frequency stimulation. It relies on elevated residual intracellular Ca2+ in presynaptic terminals after intense synaptic activity (Zucker & Regehr, 2002). The unique structure of the calyx of Held (CH) makes it an ideal model synapse (Forsythe & Barnes‐Davies, 1993; Forsythe, 1994; Borst et al. 1995; Schneggenburger & Forsythe, 2006; Borst & Soria van Hoeve, 2012) for the study of synaptic transmission and plasticity, including the PTP (Habets & Borst, 2005; Korogod et al. 2005). PTP is sensitive to protein kinase C (PKC) inhibition (Alle et al. 2001; Brager et al. 2003; Korogod et al. 2007); genetic studies using the CH suggest that PKCγ/PKCα and PKCβ isoforms are critical for inducing the PTP before and after the hearing onset in mice, respectively (Fioravante et al. 2011, 2014; Chu et al. 2014). PKC‐dependent phosphorylation of transmitter release machinery, such as Munc18‐1 (Genc et al. 2014), plays an important role in potentiating synaptic release.

An increases in release probability (P r) is the dominant factor underlying PTP at the CH before hearing onset (Habets & Borst, 2005, 2007; Korogod et al. 2005; Korogod et al. 2007; Lee et al. 2008; Fioravante et al. 2011), although PTP sometime is accompanied by a small increase in the readily releasable pool (RRP) size. It is noteworthy that the RRPs in these studies were estimated by back‐extrapolating cumulative EPSC amplitudes (termed RRPtrain hereafter) induced by a train of high frequency action potentials (APs) (Schneggenburger et al. 1999). As this approach only detects a fraction of RRP vesicles that are adjacent to Ca2+ channels, its outcome can be affected by other factors irrelevant to the RRP, such as a change in P r (Lou et al. 2008 a; Thanawala & Regehr, 2013; Neher, 2015). It was shown that RRPtrain increase plays a critical role in PTP at the mature CH after hearing onset (Chu et al. 2014). In addition, a quantal size increase from compound vesicle fusion has also been reported during PTP at immature CH (He et al. 2009; Xue & Wu, 2010).

Synaptic vesicles (SVs) are generated through different endocytosis pathways (Saheki & De Camilli, 2012; Kononenko & Haucke, 2015), and SV size can vary greatly depending on synaptic activity (Dickman et al. 2005; Hayashi et al. 2008; He et al. 2009) and on which endocytosis pathways are involved (Hayashi et al. 2008; Watanabe et al. 2013, 2014; Wu et al. 2014). Perturbations of the genes involved in endocytosis, such as clathrin, dynamin, AP‐2 and AP‐180, often lead to formation of large SVs (Zhang et al. 1998; Koh et al. 2004; Ferguson et al. 2007; Lou et al. 2008 b; Gu et al. 2013; Koo et al. 2015); smaller SV formations in AP‐2 mutants (Sato et al. 2009) have also been reported. However, in spite of more than 40 years of research on SV recycling, very little is known about the potential impact of endocytosis‐dependent SV size changes on PTP.

Dynamin is a key protein for clathrin‐mediated endocytosis. Dynamin‐1 knockout (KO) leads to activity‐dependent impairment of clathrin‐mediated endocytosis at central synapses (Ferguson et al. 2007; Lou et al. 2008 b); dynamin‐1 and ‐3 double KO causes severe defects in vesicle recycling and transmitter release (Raimondi et al. 2011; Lou et al. 2012). Here, we report that the tissue‐specific deletion of dynamin‐1 in the auditory brainstem resulted in an enhancement of PTP and a parallel activity‐dependent increase in quantal size. The quantal size increase was sensitive to actin depolymerization, which largely accounted for the enhancement of PTP in dynamin‐1 cKO synapses over controls. This work suggests that altered endocytosis can affect short‐term synaptic plasticity through SV size changes.

Methods

Conditional dynamin‐1 KO mice at the CH

Tissue‐specific dynamin‐1 conditional knockout (cKO) mice were generated by crossing Dnm1f/f (Ferguson et al. 2009) and Krox20Cre mouse lines (Voiculescu et al. 2000). In Dnm1f/f mice, exons 2–4 were flanked with lox‐P sites. Krox20Cre mice carried a Cre‐encoding sequence at the locus of Krox20 (also known as EGR2), a zinc‐finger transcriptional factor, expressed primarily in the auditory brainstem including the CH‐generating globular bushy cells in the ventral cochlear nucleus. In the first generation, we obtained Dnm1f/+ Krox20Cre/+ mice; crossing these mice with Dnm1f/f gave rise to Dnm1f/Δ Krox20Cre/+, Dnm1f/+ Krox20Cre/+, Dnm1f/Δ Krox20+/+ and Dnm1f/+ Krox20+/+ due to germline recombination (Voiculescu et al. 2000). Dnm1f/Δ Krox20Cre/+ mice were further crossed with Dnm1f/f to obtain conditional dynamin‐1 KO (Dnm1f/Δ Krox20Cre/+) mice, in which Cre expression deletes the floxed Dnm1 gene selectively in Krox20Cre‐positive mice from embryonic day (E)9–10 (expression onset of Krox20Cre) onwards (Voiculescu et al. 2000). Dynamin‐1 deletion was confirmed in cKO mice (Dnm1fΔ Krox20Cre/+ or Dnm1f/f Krox20Cre/+) (Mahapatra et al. 2016), and their littermates carrying no Krox20Cre (Dnm1f/Δ or Dnm1f/f) were used as control mice. The cKO mice had no outward phenotype compared to their littermate controls. This differs contrastingly from the much smaller body size (1/3 of control size or less) and shorter lifespan (2 weeks or less) of the conventional dynamin‐1 KO (Lou et al. 2008 b). All animal care and use were carried out in accordance with our institutional guidelines.

Brain slice preparation, patch‐clamp recording and analysis

Mice [postnatal day (P) 16‐20] were decapitated under isoflurane anaesthesia, and acute brain slices (180–200 μm) were prepared using a Leica VT1200 vibrating blade microtome (Leica Biosystem, Nussloch, Germany) as describe previously (Lou et al. 2008 b). Slices were incubated with external solution continuously bubbled with 95% O2 and 5% CO2 (pH 7.4) at 36.0 ± 1.0°C for 30 min. Patch‐clamp recordings were performed with an EPC10‐2 amplifier (HEKA Elektronik, Lambrecht, Germany) as reported previously (Lou et al. 2008 a). Patch pipettes had resistances of 3–5 MΩ. The series resistance (R s) was compensated up to 80% during recordings so that the final uncompensated R S was no more than 3 MΩ. Recordings with R s larger than 15 MΩ were not included for analysis. No offline R s correction was applied. Holding potential (V h) was set at −80 mV (without junction potential correction) and data were acquired at 20 kHz. EPSCs were evoked by a bipolar electrode placed on afferent fibres between the medial nucleus of the trapezoid body (MNTB) and the midline of brain slices. The extracellular solution (ES, in mm) contained: 120 NaCl, 2.5 KCl, 25 NaHCO3, 1.25 NaH2PO4, 2 CaCl2, 1 MgCl2, 25 glucose, 3 myo‐inositol, 2 sodium pyruvate, 0.4 l‐ascorbic acid (pH 7.4 was maintained with continuous bubbling of 95% O2–5% CO2). The patch pipette solution (IS, in mm) for EPSC recordings contained: 137 caesium gluconate, 10 Hepes, 20 TEA‐Cl, 5 sodium phosphocreatinine, 4 Mg‐ATP, 0.3 Na2GTP, 5 Cs‐EGTA, and 2 QX‐314 (pH 7.2). Strychnine‐HCl (2 μm), bicuculline (10 μm) and d‐AP5 (50 μm) were freshly prepared and added to the ES before experiments. d‐AP5 was purchased from Tocris, latrunculin‐B and PKC blocker GF109203X were purchased from Abcam, and all the other chemicals were purchased from Sigma‐Aldrich. Latrunculin‐B (Lat‐B) was perfused at least 10 min prior to and was continuously present during recordings. GF109203X (2 μm) was applied for an hour before recordings and continuously perfused during recordings. All experiments were conducted at room temperature (∼20–22°C).

PTP was induced by a tetanic train of APs at 100 Hz for 4 s, and EPSCs were triggered by a pair of APs (10 ms interval) at 0.1 Hz during PTP. We applied six trials before PTP and 30 trials after PTP induction. The baseline for PTP was calculated from the average EPSC amplitude (the first EPSC in six trials and the induction). PTP was presented as the percentage changes of EPSC peak relative to the baseline in each synapse. We recorded spontaneous excitatory postsynaptic currents (spEPSCs) to analyse quantal size since they were easily distinguishable from AP‐triggered EPSCs. spEPSC events were collected from each sweep (10 s, 0.1 Hz) after skipping the first second that contains evoked EPSCs. We designated the first sweep after the tetanic stimulation as AS‐1. Basal spEPSCs were collected from six traces before the induction in each synapse; average spEPSC in each genotype was calculated from the mean value of each synapse (bar graph, unless otherwise specified). For histogram and probability plots, all individual spEPSCs (at a given time window) were pooled across each synapse in each genotype.

Paired‐pulse ratio (PPR) was calculated as the ratio of the second EPSC amplitude divided by the first. RRPtrain was measured by back‐extrapolating cumulative EPSCs induced by a 100 Hz train (30 APs) to the first peak (Lou et al. 2008 a). Experiments were performed in the presence of 100 μm cyclothiazide and 1 mm kynurenic acid to minimize postsynaptic receptor desensitization and saturation. RRPtrain was monitored at four different time points (20 s before, 15, 40 and 80 s after the induction) as the basal, peak and first and second recovery of RRPtrain. Lat‐B (15 μm, ab144291, Abcam) was perfused during the experiments for 10 min prior to recordings; experiments were completed within 60 min for the same slice. Data were analysed using IgorPro (WaveMetrics, Lake Oswego, OR, USA), Origin (OrginLab, Northampton, MA, USA) and/or Excel (Microsoft). Fiber stimulation artifacts before EPSCs were blanked in some figures for clarity.

Statistics

Values were presented as means ± SEM (standard error of the mean) with n representing the number of synapses unless otherwise indicated. Average values in the bar graphs were calculated from the average of each cell rather than individual events across a group. Each data point for statistics was collected from at least three mice. Statistical analyses were performed using Student's two‐tailed t test or Mann–Whitney U test as specified. The significance level was set at P < 0.05 and denoted with asterisks (* P < 0.05, ** P < 0.01, *** P < 0.001).

Results

Conditional ablation of dynamin‐1 enhances PTP at the mature CH

Dynamin‐1 is the most abundant of the three dynamin isoforms (dynamin‐1, ‐2 and ‐3) expressed in the mammalian brain (Ferguson & De Camilli, 2012). It is crucial for the SV regeneration by promoting vesicle fission. Conventional dynamin‐1 KO mice are perinatally lethal (Ferguson et al. 2007), preventing the study of PTP at mature CH. Therefore, we generated dynamin‐1 conditional KO mice to study PTP after hearing onset (P16–20) (see Methods). We monitored EPSC amplitude changes with repeated paired‐pulses at 0.1 Hz before and after a train of tetanic APs (100 Hz, 4 s). PTP peaked at ∼10 s after the completion of an induction train in controls (121.5 ± 3.8%, n = 8 neurons) and recovered within 1–2 min (Fig. 1 A). With this induction protocol, PTP is smaller at the mature CH (P16–20) than at the immature CH of pre‐hearing animals, consistent with a developmental decrease of PTP under moderate induction (Korogod et al. 2005). The basal EPSC amplitude in cKO (∼6 nA) was not significantly different from control (∼7 nA) (V h = −80 mV, without offline correction) (Mahapatra et al. 2016). Interestingly, PTP is significantly higher in cKO (141.4 ± 6.6%, n = 11 neurons) than control (P = 0.007) (Fig. 1 A and B), and recovered in a similar time (1–2 min). To probe P r changes, we monitored paired‐pulse ratio (PPR) (10 ms interval) throughout the PTP experiment at 10 s intervals (Fig 1 A insets). The PPRs did not significantly change during PTP in both control and cKO synapses, despite a smaller decrease in PTP peak time in cKO. We also found no significant difference in PPR between control and cKO synapses at the peak of PTP (10 s after the induction) (P = 0.52). This is consistent with the previous results at mature CH under normal condition (Chu et al. 2014).

Figure 1. Tissue‐specific dynamin‐1 deletion significantly increases PTP at the mature CH.

Figure 1

Open in a new tab

A, a representative PTP recording from the mature CH (P16–20). A paired pulse (10 ms inter‐interval) was applied at 0.1 Hz before and after the tetanic stimulation (4 s, 100 Hz). Each dashed line shows the average basal EPSC amplitude. B, average PTP for control (121.4 ± 3.8%, n = 8) and KO mice (141.4 ± 6.6%, n = 11; P = 0.0072). C, paired‐pulse ratios remain unaltered [not significant (NS)] at PTP peak for both control (0.69 ± 0.08, n = 8) and KO (0.75 ± 0.05, n = 11, P = 0.52). Arrows indicate starting points of the tetanic stimulation train. [Colour figure can be viewed at wileyonlinelibrary.com]

RRPtrain increases during PTP in both control and dynamin‐1 cKO

We estimated RRPtrain triggered by a train of APs (100 Hz, 30 APs) at different times (Fig. 2 A) in the presence of 100 μm cyclothiazide and 1 mm kynurenic acid to minimize postsynaptic receptor desensitization and saturation (Neher & Sakaba, 2001; Wadiche & Jahr, 2001; Taschenberger et al. 2002; Wong et al. 2003). RRPtrain was slightly smaller in cKO than in control under basal condition, a difference that may be partially explained by the enhanced release rate during the later phase of high‐frequency stimulation in cKO (Fig. 2 C). Direct presynaptic capacitance changes (C m) induced by a 50 ms depolarization did not reveal a significant difference in basal RRP and presynaptic Ca2+ influx between cKO and control (Mahapatra et al. 2016), despite these results diverging from what is observed in conventional dynamin‐1 KO (presumably due to the healthier condition of cKO mice than conventional KOs; see Methods).

Figure 2. RRPtrain increases during PTP at mature CH in both control and cKO.

Figure 2

Open in a new tab

A, the experimental protocol for RRPtrain estimation during PTP. Recordings were performed in the presence of 0.1 mm cyclothiazide and 1 mm kynurenic acid. A train of APs (30 APs, 100 Hz) was applied at different times (arrows); the dashed line represents the EPSC peak level before PTP induction. Tetanic stimulation is a train of high frequency APs at 100 Hz for 4 s. B and C, EPSCs under basal conditions and at the PTP peak; the cumulative EPSC plots are shown on the right. D, RRPtrain under basal conditions and at PTP peak (left panel); their percentage increases (PTP vs. basal) are shown in the right panel (116 ± 4.0% in control, 141.0 ± 6.0% in cKO; P = 0.0076). Average RRPtrain in control (13.8 ± 1.2 nA under basal conditions, 16.1 ± 1.5 nA during PTP, n = 6 synapses) and KO synapses (9.5 ± 1.5 nA under basal conditions, 13.6 ± 2.4 nA during PTP; n = 7). [Colour figure can be viewed at wileyonlinelibrary.com]

During PTP, RRPtrain increased significantly in both control and cKO synapses. Interestingly, the relative increase of RRPtrain in cKO was higher (141 ± 6%, n = 7) than in control synapses (116 ± 4%, n = 6; P = 0.0076) (Fig. 2 B–D). This may be partially explained by the activity‐dependent increase in spEPSC amplitude in cKO (see below). The enhanced release rate late in the train in cKO (Fig. 2) (Mahapatra et al. 2016) may lead to an underestimate of the RRPtrain value, but this factor might have little impact on the relative changes during PTP as it is present before and after the PTP. Thus, we conclude that during PTP at the mature CH, loss of dynamin‐1 leads to a larger RRPtrain increase in cKO than in control.

The increases in RRPtrain have been frequently reported during PTP at the CH (Habets & Borst, 2007; Lee et al. 2008; Chu et al. 2014). It is known that an RRPtrain reflects only a fraction of the RRP (a sub‐pool) that represents vesicles adjacent to calcium channels (Neher, 2015). This poses several limitations in measuring the RRP. The use of RRPtrain in this way depends on two assumptions: release rate must be constant during the train of stimulation, and the vesicle release rate must be equal to the RRP refilling rate late in the train (Neher, 2015). A low P r or weak stimulation can significantly affect the RRPtrain value and underestimate the RRP (Lou et al. 2008 a; Thanawala & Regehr, 2013). In other words, a change in P r could parallel a change in RRPtrain value, without causing a significant change in the RRP estimated by the maximal stimulation (Lou et al. 2008 a).

The loss of dynamin‐1 increases spEPSC amplitude selectively during PTP

We measured spEPSCs in MNTBs under basal condition, and found no significant difference in their amplitude between control (−62.4 ± 4.9 pA, n = 8 synapses) and cKO (−66.1 ± 2.8 pA, n = 11; P = 0.52; unpaired t test) (Fig. 3 A and D). PTP induction in control (4 s, 100 Hz) led to a small increase in spEPSC amplitude at the PTP peak (−70.1 ± 4.8 pA; P = 0.0013, paired t test; Fig. 3 A and D), similar to some recent studies (He et al. 2009; Fioravante et al. 2011). This increase peaked within ∼10 s after PTP induction (Fig. 3 D) and recovered faster than PTP itself (Fig. 1 B). Interestingly, this increase was more prominent in cKO (from −66.1 ± 2.8 to −76.8 ± 3.4 pA before and during PTP, respectively; P = 0.0008, paired t test) than control synapses (Fig. 3 A–C), and it lasted longer (Fig. 3 D). The histogram of individual spEPSC amplitudes at the PTP peak (the first trace after tetanic stimulation, AS‐1, see Methods) revealed an addition of subpopulation of spEPSCs with larger amplitudes in cKO (Fig. 3 B); accordingly, the cumulative probability plots in cKO were clearly right‐skewed (Fig. 3 C). The cumulative probability curve of spEPSC amplitudes at PTP peak in cKO was largely right‐skewed (basal vs. PTP peak at AS‐1), in contrast to a marginal, parallel right‐shift in control. The average kinetics between the large spEPSCs and normal spEPSCs were comparable. These data demonstrate a more pronounced increase in spEPSC amplitude in cKO than control during PTP, due to the increased population of large spEPSCs induced by PTP induction. spEPSC frequencies in control and cKO were not different at rest, and both showed a ∼3‐fold increase during PTP (Fig. 3 E).

Figure 3. Increase in spEPSC amplitude during PTP in the absence of dynamin‐1.

Figure 3

Open in a new tab

A, representative spEPSC traces at rest and at PTP peak (AS‐1). Inset shows the enlarged view of a large spEPSC. Note the pronounced increases in both spEPSC amplitude and frequency, particularly in cKO. B, histogram of spEPSC amplitudes at the PTP peak (AS‐1) from both control (average amplitude: −70.1 ± 4.8 pA, n = 1259 events from 8 synapses) and cKO (average amplitude: −76.8 ± 3.4 pA, n = 1647 events from 11 synapses; P < 0.001, Mann–Whitney U test). Inset shows an expanded view between −70 and −300 pA. C, cumulative probability of spEPSC amplitudes shown in B. Note the right‐skewed curve in cKO compared to control. D, average increases in spEPSC amplitude (from cell averages) during PTP. Note the basal spEPSC amplitude was similar between control (−62.4 ± 4.9 pA, n = 8 synapses) and KO (−66.1 ± 2.8 pA, n = 11 synapses; P = 0.52). During PTP, AS‐1 control: −70.14 ± 4.8 pA, P = 0.0014 (paired t test, for AS‐1 vs. basal); cKO: −76.8 ± 3.4 pA, P = 0.0008 (for AS‐1 vs. basal). During PTP AS‐2, control: 66.0 ± 5.9 pA, P = 0.12 (for AS‐2 vs. basal); cKO: −76.6 ± 3.5 pA, P = 0.0022 (for AS‐2 vs. basal). E, average spEPSC frequency (from cell averages) changes during PTP. Basal frequency in control (5.5 ± 0.7 Hz, n = 8) was not different from cKO (3.72 ± 0.6 Hz, n = 11 synapses, P = 0.08). During PTP, AS‐1 control: 16.7 ± 2.4 Hz, P = 0.0004 (for AS‐1 vs. basal); AS‐1 KO: 15.9 ± 4.0, P = 0.009 (for AS‐1 vs. basal); AS‐2 control: 11.2 ± 1.5 Hz, P = 0.001 (for AS‐2 vs. basal); cKO: 10.3 ± 2.6 Hz; P = 0.02 (for AS‐2 vs. basal). Paired t tests were applied in D and E. [Colour figure can be viewed at wileyonlinelibrary.com]

In principle, the spEPSC amplitude increase during PTP can arise from both pre‐ and postsynaptic factors. However, the comparable spEPSC amplitude between control and cKO at rest (Fig. 3 D) does not support a postsynaptic effect. Moreover, given that the activity‐dependent formation of large vesicles and vacuoles in nerve terminals has been observed with EM in cultured neurons (Ferguson et al. 2007) and in the CH (Lou et al. 2008 b), these data further support the view that the increase in large spEPSC amplitude is caused by an increase in vesicle size.

Actin depolymerization preferentially abolishes the spEPSC amplitude increase and PTP enhancement in cKO

It is well established that intense stimulation induces bulk endocytosis at nerve terminals (Clayton et al. 2008; Hayashi et al. 2008; Saheki & De Camilli, 2012; Kononenko et al. 2014; Wu et al. 2014; Kononenko & Haucke, 2015), and this form of endocytosis requires actin polymerization (Holt et al. 2003; Nguyen et al. 2012; Gormal et al. 2015). Interestingly, bulk endocytosis is enhanced in dynamin‐1 KO synapses under intense stimulation (Hayashi et al. 2008; Wu et al. 2014). In addition, dynamin gene deletion leads to increased F‐actin puncta in fibroblasts (Ferguson et al. 2009) and enhanced actin polymerization in pancreatic β‐cells (Fan et al. 2015). These data raise the possibility that the loss of dynamin enhances actin polymerization at nerve terminals, which in turn promotes bulk endocytosis and the formation of larger vesicles in cKO during intense stimulation. This might explain the prominent activity‐dependent increase of spEPSCs in cKO during PTP.

To test this idea, we applied Lat‐B (15 μm), an inhibitor of actin polymerization, to interrupt actin‐dependent processes, such as bulk and/or ultrafast endocytosis. Figure 4 A and B shows representative recordings from different groups. Lat‐B had no effect on PTP in control synapses (121.6 ± 5.6, n = 8), but blocked the enhancement of PTP in cKO (128.72 ± 4.5, n = 7; P = 0.34) resulting in a decrease of PTP in cKO to the level of control synapses (Fig. 4 C). We also observed transmission failures in 1 out of 7 cKO cells during 4 s tetanic stimulation (Fig. 4 B, insert), apparently without affecting the PTP amplitude as compared to other recordings. An example without failure in a cKO synapse is also shown (Fig. 4 B, bottom). Transmission and AP failures in cKO have been characterized in a different study (Mahapatra et al. 2016).

Figure 4. Actin depolymerization abolishes the PTP enhancement in cKO synapses.

Figure 4

Open in a new tab

A, intact PTP in the presence of Lat‐B (15 μm) in a control synapse. B, two representative PTP recordings in cKO synapses. The top panel shows transmission failures during 4 s tetanic stimulation (in 1 out of 7 synapses) and the bottom panel shows an example without failure. Inset shows the tetanic stimulation on an expanded time scale. C, Lat‐B reduced PTP in cKO (128.72 ± 4.5%, n = 7 synapses), without affecting PTP in control (121.64 ± 5.6%, n = 8 synapses). PTPs between cKO and control were statistically indistinguishable in the presence of Lat‐B (P = 0.34). [Colour figure can be viewed at wileyonlinelibrary.com]

Interestingly, the increase in spEPSC amplitude during PTP was fully blocked by Lat‐B in both controls (70.7 ± 4.9 and 66.3 ± 4.0 pA, before and during PTP, respectively; n = 5; P = 0.28) and cKO (71.3 ± 6.5 and 72.6 ± 5.4 pA, before and during PTP, respectively; n = 7; P = 0.73) (Fig. 5). The spEPSC frequency increase was not affected during PTP in both control (4.7 ± 0.5 and 13.1 ± 2.8 Hz, for basal and PTP, respectively; P = 0.03) and cKO (3.7 ± 0.7 and 11.8 ± 2.9 Hz, for basal and PTP, respectively; P = 0.014) (Fig. 5). This is consistent with the increased residual intracellular Ca2+ right after PTP induction (Korogod et al. 2005). The parallel effects of Lat‐B on reducing the amplitudes of both spEPSC and PTP in cKO suggest that the increase in a number of large spEPSCs contributes to PTP enhancement in cKO. This blocking effect of Lat‐B may reflect a role of actin in large vesicle regeneration in both bulk and ultrafast endocytosis (Hayashi et al. 2008; Watanabe et al. 2014; Wu et al. 2014). It was reported that ultrafast endocytosis occurs only at physiological temperature but not at room temperature (Watanabe et al. 2014), while bulk endocytosis still operates robustly at room temperature (Hayashi et al. 2008; Wu et al. 2014). Therefore, the actin‐dependent formation of large vesicles most likely results from bulk endocytosis rather than ultrafast endocytosis since our experiments were performed at room temperature. Although we noticed a marginal increase in basal spEPSC amplitudes under Lat‐B (compared to normal condition), they were not significantly different both in control (in normal condition: −62.4 ± 4.9 pA; under Lat‐B: −70.7 ± 4.9 pA; P = 0.25) and in cKO (in normal condition: −66.1 ± 2.8 pA; under Lat‐B: −71.3 ± 6.5 pA; P = 0.48).

Figure 5. Actin depolymerization abolished spEPSC amplitude increase during PTP in cKO synapses.

Figure 5

Open in a new tab

A, representative spEPSC traces at basal condition and the PTP peak (AS‐1) in the presence of Lat‐B (15 μm). B, Lat‐B blocked the increase of spEPSC amplitude during PTP in both control and cKO synapses. Note the nearly overlapping cumulative probability curves before and after PTP induction (AS‐1) for both control and cKO. All spEPSC events from each genotype were pooled under basal conditions (n = 213 events from 5 control neurons; n = 231 events from 7 cKO neurons; P = 0.45, Mann–Whitney U test) and at PTP peak (AS‐1) (n = 633, 5 control neurons; n = 745 events, 7 cKO; P = 0.11, Mann–Whitney U test). C, average spEPSC amplitude and frequency from each cell mean in control (n = 5 synapses) and cKO (n = 7 synapses) (paired t test). Note that three recordings from MNTBs in control showed no spEPSCs at rest although these neurons showed normal evoked EPSCs, and they were excluded from spEPSC analysis. [Colour figure can be viewed at wileyonlinelibrary.com]

PKC blocker GF109203X inhibits PTP and spEPSC amplitude in both control and cKO

Although Lat‐B abolished the enhancement of PTP in cKO over control, PTP was still prominent in cKO synapses. Given the important role of PKC‐dependent signalling at the CH (Korogod et al. 2007; Fioravante et al. 2011), we evaluated the effect of a conventional pan‐PKC blocker GF109203X (2 μm) on PTP in cKO. Brain slices were pre‐incubated for a minimum of 60 min in GF109203X before recordings (Chu et al. 2014). Under normal conditions, PTP peaked at the second test pulse (10 s) after induction and then declined. GF109203X almost completely abolished PTP at this time point in both control (107.3 ± 2.9%; n = 8) and cKO (107.1 ± 5.1%; n = 8) synapses (Fig. 6 A and B), in agreement with previous studies (Fioravante et al. 2011; Chu et al. 2014). In addition, we also noticed a delayed increase in EPSC amplitude in both control and cKO synapses in the presence of GF109203X, which recovered very slowly (Fig. 6 B).

Figure 6. PTP is sensitive to the PKC blocker GF109203X in both control and cKO synapses.

Figure 6

Open in a new tab

A, PTPs from a control and a cKO synapse in the presence of 2 μm GF109203X. After tetanic stimulation (4 s, 100 Hz), the EPSC potentiation at the second test pulse was almost abolished, but EPSCs became larger at a later stage and remained elevated for a longer time than in the absence of the drug. B, the average PTP in the presence of GF109203X. PTPs at 10 s after the induction in control (107.3 ± 2.9%; n = 8) and cKO (107.1 ± 5.1%; n = 8) synapses were blocked as compared to normal conditions (without GF109203X). C, the average spEPSC amplitude (left) and frequency (right) from both control and cKO synapses at basal and AS‐1. Note that the average frequency, but not the spEPSC amplitude, in AS‐1 was still significantly higher than basal levels. [Colour figure can be viewed at wileyonlinelibrary.com]

The basal amplitude of spEPSCs in the presence of GF109203X was slightly larger than in the normal condition; however, the difference was not statistically significant both in control (P = 0.09) and in cKO (P = 0.14). The spEPSC amplitude increase during PTP (AS‐1) was blocked by GF109203X (Fig. 6 C, left panel) in control (basal: −80.3 ± 7.4 pA; AS‐1: −81.3 ± 6.0 pA; P = 0.93), consistent with a previous report under a similar stimulation condition (4 s, 100 Hz) in young rats (Xue & Wu, 2010). Similar results were observed in cKO (basal: −75.9 ± 5.4 pA; AS‐1: 72.2 ± 2.7 pA; P = 0.38). The spEPSC frequency remained much higher during PTP than basal (Fig. 6 C, right panel) in control (basal: 4.4 ± 0.96 Hz; AS‐1: 10.8 ± 1.8 Hz, P = 0.0018) and cKO (basal: 3.04 ± 0.7 Hz; AS‐1: 6.5 ± 1.5, P = 0.017). These data suggest that PKC is essential for PTP in both control and cKO. In addition, PKC also influences spEPSC amplitude increase during PTP, particularly in cKO synapses.

Discussion

Using the mature CH, we have shown that tissue‐specific dynamin‐1 deletion enhances PTP. This effect results from increases in both RRPtrain and spEPSC amplitude in cKO synapses after induction of PTP. Actin depolymerization induced by Lat‐B blocked the increase in spEPSC amplitude in both control and cKO synapses; Lat‐B accordingly abolished the enhancement of PTP in cKO synapses, without affecting PTP in control. The increases in spEPSC amplitude and PTP were sensitive to PKC inhibition in both control and cKO. These data suggest that endocytic control of quantal size also affects short‐term synaptic plasticity. This may provide a new way for synapses to adapt to different environments under physiological and pathophysiological conditions. As an important protein for SV regeneration, dynamin‐1 improves the quality control of vesicle size, thus helping to maintain the stability of PTP in addition to supplying new SVs.

The contribution of P r, RRP and quantal size in PTP at the CH

PTP changes in the CH during development (Korogod et al. 2005). Previous studies suggest that a P r increase plays a dominant role in PTP before hearing onset (Habets & Borst, 2005; Korogod et al. 2005). This effect mainly results from an increase in apparent Ca2+ sensitivity of vesicle fusion (Lou et al. 2005; Korogod et al. 2007) mediated by conventional PKC activation (Korogod et al. 2007; Fioravante et al. 2011) and subsequent Munc18‐1 phosphorylation (Genc et al. 2014). In these studies, PPR decreases significantly during PTP, with or without a moderate RRPtrain increase. However, after hearing onset, the RRPtrain increase appears to play an important role in PTP at CH (Chu et al. 2014). In agreement with these observations, we observed a large RRPtrain increase during PTP, without significant PPR changes. Interestingly, the relative increase in RRPtrain during PTP in cKO is significantly larger than in control. This may be partially explained by the increased spEPSC amplitude during PTP in cKO.

Most, if not all, RRP measurements in PTP studies are performed by using the RRPtrain, a method that is based on back‐extrapolation of the cumulative EPSC amplitudes evoked by a high‐frequency train of APs (Schneggenburger et al. 1999). This approach relies on two assumptions for the proper estimation of RRP (Neher, 2015): release rate must remain constant during the train and be equal to the vesicle refilling rate at steady‐state. In many cases, RRPtrain only reflects a fraction of RRP vesicles adjacent to calcium channels (Neher, 2015). This may explain why RRP size estimated from RRPtrain is consistently smaller than that estimated from maximal stimulation through direct presynaptic depolarization and Ca2+ uncaging (Schneggenburger et al. 2002). Low P r and insufficient stimulation may lead to a small RRPtrain due to incomplete RRP depletion (Lou et al. 2008 a; Thanawala & Regehr, 2013), without affecting the RRP measured by maximal stimulation (Lou et al. 2008 a). Therefore, despite RRPtrain providing valuable information for the relative changes of effective releasable pool during PTP, an RRPtrain change may not necessarily reflect a change in the RRP (the total number of release‐competent SVs), and it can be influenced by a P r change. A significant PPR decrease was reported when KYN and CTZ were present during PTP (Chu et al. 2014), suggesting a potential P r increase during PTP at mature CH. This raises the possibility that PPR may not always be sensitive to the P r change at mature CH under normal conditions. It is likely that the uniform PKC‐dependent pathway contributes to PTP through a P r increase at different developmental stages. Future experiments are required to test this idea.

We observed no significant difference in spEPSC amplitude between control and cKO under basal condition, consistent with our previous study (Lou et al. 2008 b). PTP induction led to a significant increase in spEPSC frequency and a very small increase in spEPSC amplitude during PTP in control, in agreement with previous studies (He et al. 2009; Xue & Wu, 2010; Fioravante et al. 2011). Interestingly, the increase of the spEPSC amplitude was much larger in cKO than in control synapses, and more events with a large spEPSC amplitude appeared during PTP in cKO (Fig. 3 C). Lat‐B blocked this change in spEPSC amplitude during PTP (Fig. 5 B) and selectively abolished the enhancement of PTP in cKO over control (Fig. 4), suggesting a role of spEPSC amplitude increase in the enhancement of PTP in cKO. This may relate to a potential alteration of actin in cKO synapses since the loss of dynamin influences actin polymerization and function (Ferguson et al. 2009; Fan et al. 2015). The finding of an activity‐dependent increase in quantal size during PTP in cKO agrees well with the synaptic ultrastructural alterations that are observed in the absence of dynamin‐1 (Hayashi et al. 2008; Lou et al. 2008 b; Wu et al. 2014).

Synaptic transmission and vesicle recycling are adapted to physiological temperature (37–38°C) to optimize normal brain function, and their temperature dependence has been characterized (Pyott & Rosenmund, 2002; Kushmerick et al. 2006; Postlethwaite et al. 2007; Renden & von Gersdorff, 2007; Watanabe et al. 2013). At the CH, physiological temperature decreases presynaptic AP amplitude and duration (Borst & Sakmann, 1998; Kushmerick et al. 2006), reduces AP‐triggered calcium influx (Borst & Sakmann, 1998), increases vesicle endocytosis efficiency by the addition of a fast retrieval component (Renden & von Gersdorff, 2007), and accelerates vesicle recruitment rate to reduce short‐term depression (Kushmerick et al. 2006). On the postsynaptic side, raised temperature increases the EPSC amplitude and accelerates its kinetics (Kushmerick et al. 2006; Postlethwaite et al. 2007) through the opening of AMPA receptor channels to higher conducting states (Postlethwaite et al. 2007). On the other hand, the basic molecular processes of neurotransmission remain unchanged at different temperatures. Raised temperature does not change P r and RRP at the CH (Kushmerick et al. 2006); it only marginally accelerates PTP decay in P4–11 rat CH (Habets & Borst, 2007). Thus, the results obtained at room temperature here provide new insight into PTP modulation in the mammalian brain.

Endocytosis‐mediated large SV formation contributes to the spEPSC amplitude increase and PTP enhancement in the absence of dynamin‐1

Given the quantal nature of synaptic transmission, any changes in RRP, P r and quantal size during PTP could, in principle, affect transmitter release. As discussed above, the increases in P r and/or RRPtrain are primary factors that contribute to PTP through the PKC‐dependent pathway; however, the potential role of endocytosis‐mediated quantal size changes in PTP remains poorly understood. On the other hand, quantal size changes are very common during vigorous synaptic activity (Heuser, 1974; Llano et al. 2000; Matthews & Sterling, 2008; He et al. 2009; Li et al. 2009) and in many gene mutants related to synaptic endocytosis. Multiple mechanisms have been proposed to account for the quantal size increases, including endocytic reformation of large vesicles, multiple quantal release (Llano et al. 2000; Wadiche & Jahr, 2001; Singer et al. 2004; Li et al. 2009), compound vesicle formation from homotypic vesicle fusion (He et al. 2009; Xue & Wu, 2010), and vesicle fusion pore changes (Chapochnikov et al. 2014). Among these mechanisms, large vesicle formation due to defective endocytosis has been well documented in synaptic terminals after genetic perturbations of endocytosis regulatory proteins (Zhang et al. 1998; Deak et al. 2004; Dickman et al. 2005; Ferguson et al. 2007; Lou et al. 2008 b). The exact mechanism of vesicle size control is unclear, but different SV regeneration pathways and vesicle coating or fission proteins (Wu et al. 2014; Kononenko & Haucke, 2015) may regulate SV size control.

Here we provide evidence that perturbing endocytosis by dynamin‐1 deletion significantly enhances PTP at the CH, and that an spEPSC amplitude increase partly contributes to the enhancement of PTP. Two sets of data suggest that endocytosis‐mediated large vesicle formation in cKO contributes to the increase in spEPSC amplitude during PTP. First, dynamin‐1 deletion causes a pronounced increase in the number of large spEPSCs in cKO after tetanic stimulation (Fig. 3). The large SV formation induced by intense stimulation is enhanced in dynamin‐1 KO synapses (Ferguson et al. 2007; Hayashi et al. 2008; Lou et al. 2008 b; Wu et al. 2014), as well as in terminals after endocytic gene perturbations (Zhang et al. 1998; Koh et al. 2004; Gu et al. 2013; Koo et al. 2015). Second, the spEPSC size increase during PTP can be blocked by Lat‐B, a reagent that disrupts F‐actin polymerization and bulk endocytosis (Holt et al. 2003; Nguyen et al. 2012; Gormal et al. 2015). A potential explanation for this effect is that actin depolymerization alters bulk endocytosis and SV formation (Shupliakov et al. 2002; Richards et al. 2004; Watanabe et al. 2013).

In conjunction with our recent work in cKO demonstrating a reduction of synaptic depression (Mahapatra et al. 2016), our current findings suggest a role of endocytosis in the control of vesicle size during PTP. The loss of dynamin‐1 affects multiple aspects of SV recycling, including the efficiency and pathways of membrane retrieval and subsequent vesicle reformation. These effects appear to be able to modulate multiple forms of short‐term synaptic plasticity under various synaptic activities and on different time scales at central synapses.

Additional information

Competing interests

None declared.

Author contributions

S.M.: conception and design; collection and assembly of data; data analysis and interpretation; manuscript writing. X.L.: conception and design; financial support; data analysis and interpretation; manuscript writing. Both authors have approved the final version of the manuscript and agree to be accountable for all aspects of the work. All persons designated as authors qualify for authorship, and all those who qualify for authorship are listed.

Funding

This work is supported by the National Institutes of Health (NIH) grants R01DK093953 (X.L.) and P30NS069271, brain research foundation BRFSG201407.

Acknowledgements

We thank Pietro DeCamilli and Shawn Ferguson for providing DNM1f/f mice, Patrick Charnay and Ralf Schneggenburger for providing Krox20Cre mice. We thank Chen Ji and Fan Fan for very helpful discussions; Charles Wollitz, Jaffna Mathiaparanam and Elizabeth Westrick for technical support; Bill Chiu and Meyer Jackson for reading the manuscript and very helpful suggestions.

References

  1. Alle H, Jonas P & Geiger JR (2001). PTP and LTP at a hippocampal mossy fibre‐interneuron synapse. Proc Natl Acad Sci USA 98, 14708–14713. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Borst JG, Helmchen F & Sakmann B (1995). Pre‐ and postsynaptic whole‐cell recordings in the medial nucleus of the trapezoid body of the rat. J Physiol 489, 825–840. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Borst JG & Sakmann BCP (1998). Calcium current during a single action potential in a large presynaptic terminal of the rat brainstem. J Physiol 506, 143–157. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Borst JG & Soria van Hoeve J (2012). The calyx of Held synapse: from model synapse to auditory relay. Annu Rev Physiol 74, 199–224. [DOI] [PubMed] [Google Scholar]
  5. Brager DH, Cai X & Thompson SM (2003). Activity‐dependent activation of presynaptic protein kinase C mediates post‐tetanic potentiation. Nat Neurosci 6, 551–552. [DOI] [PubMed] [Google Scholar]
  6. Chapochnikov NM, Takago H, Huang CH, Pangrsic T, Khimich D, Neef J, Auge E, Gottfert F, Hell SW, Wichmann C, Wolf F & Moser T (2014). Uniquantal release through a dynamic fusion pore is a candidate mechanism of hair cell exocytosis. Neuron 83, 1389–1403. [DOI] [PubMed] [Google Scholar]
  7. Chu Y, Fioravante D, Leitges M & Regehr WG (2014). Calcium‐dependent PKC isoforms have specialized roles in short‐term synaptic plasticity. Neuron 82, 859–871. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Clayton EL, Evans GJ & Cousin MA (2008). Bulk synaptic vesicle endocytosis is rapidly triggered during strong stimulation. J Neurosci 28, 6627–6632. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Deak F, Schoch S, Liu X, Sudhof TC & Kavalali ET (2004). Synaptobrevin is essential for fast synaptic‐vesicle endocytosis. Nat Cell Biol 6, 1102–1108. [DOI] [PubMed] [Google Scholar]
  10. Dickman DK, Horne JA, Meinertzhagen IA & Schwarz TL (2005). A slowed classical pathway rather than kiss‐and‐run mediates endocytosis at synapses lacking synaptojanin and endophilin. Cell 123, 521–533. [DOI] [PubMed] [Google Scholar]
  11. Fan F, Ji C, Wu Y, Ferguson SM, Tamarina N, Philipson LH & Lou X (2015). Dynamin 2 regulates biphasic insulin secretion and plasma glucose homeostasis. J Clin Invest 125, 4026–4041. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Ferguson SM, Brasnjo G, Hayashi M, Wolfel M, Collesi C, Giovedi S, Raimondi A, Gong LW, Ariel P, Paradise S, O'Toole E, Flavell R, Cremona O, Miesenbock G, Ryan TA & De Camilli P (2007). A selective activity‐dependent requirement for dynamin 1 in synaptic vesicle endocytosis. Science 316, 570–574. [DOI] [PubMed] [Google Scholar]
  13. Ferguson SM & De Camilli P (2012). Dynamin, a membrane‐remodelling GTPase. Nat Rev Mol Cell Biol 13, 75–88. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Ferguson SM, Raimondi A, Paradise S, Shen H, Mesaki K, Ferguson A, Destaing O, Ko G, Takasaki J, Cremona O, O2 Toole E & De Camilli P (2009). Coordinated actions of actin and BAR proteins upstream of dynamin at endocytic clathrin‐coated pits. Dev Cell 17, 811–822. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Fioravante D, Chu Y, de Jong AP, Leitges M, Kaeser PS & Regehr WG (2014). Protein kinase C is a calcium sensor for presynaptic short‐term plasticity. Elife 3, e03011. [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
  16. Fioravante D, Chu Y, Myoga MH, Leitges M & Regehr WG (2011). Calcium‐dependent isoforms of protein kinase C mediate posttetanic potentiation at the calyx of Held. Neuron 70, 1005–1019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Forsythe ID ( 1994). Direct patch recording from identified presynaptic terminals mediating glutamatergic EPSCs in the rat CNS, in vitro. J Physiol 479, 381–387. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Forsythe ID & Barnes‐Davies M (1993). The binaural auditory pathway: excitatory amino acid receptors mediate dual timecourse excitatory postsynaptic currents in the rat medial nucleus of the trapezoid body. Proc Biol Sci 251, 151–157. [DOI] [PubMed] [Google Scholar]
  19. Genc O, Kochubey O, Toonen RF, Verhage M & Schneggenburger R (2014). Munc18‐1 is a dynamically regulated PKC target during short‐term enhancement of transmitter release. Elife 3, e01715. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Gormal RS, Nguyen TH, Martin S, Papadopulos A & Meunier FA (2015). An acto‐myosin II constricting ring initiates the fission of activity‐dependent bulk endosomes in neurosecretory cells. J Neurosci 35, 1380–1389. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Gu M, Liu Q, Watanabe S, Sun L, Hollopeter G, Grant BD & Jorgensen EM (2013). AP2 hemicomplexes contribute independently to synaptic vesicle endocytosis. Elife 2, e00190. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Habets RL & Borst JG (2005). Post‐tetanic potentiation in the rat calyx of Held synapse. J Physiol 564, 173–187. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Habets RL & Borst JG (2007). Dynamics of the readily releasable pool during post‐tetanic potentiation in the rat calyx of Held synapse. J Physiol 581, 467–478. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Hayashi M, Raimondi A, O'Toole E, Paradise S, Collesi C, Cremona O, Ferguson SM & De Camilli P (2008). Cell‐ and stimulus‐dependent heterogeneity of synaptic vesicle endocytic recycling mechanisms revealed by studies of dynamin 1‐null neurons. Proc Natl Acad Sci USA 105, 2175–2180. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. He L, Xue L, Xu J, McNeil BD, Bai L, Melicoff E, Adachi R & Wu LG (2009). Compound vesicle fusion increases quantal size and potentiates synaptic transmission. Nature 459, 93–97. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Heuser JE(1974). A possible origin of the ‘giant’ spontaneous potentials that occur after prolonged transmitter release at frog neuromuscular junctions. J Physiol 239, 106p–108p. [DOI] [PubMed] [Google Scholar]
  27. Holt M, Cooke A, Wu MM & Lagnado L (2003). Bulk membrane retrieval in the synaptic terminal of retinal bipolar cells. J Neurosci 23, 1329–1339. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Koh TW, Verstreken P & Bellen HJ (2004). Dap160/intersectin acts as a stabilizing scaffold required for synaptic development and vesicle endocytosis. Neuron 43, 193–205. [DOI] [PubMed] [Google Scholar]
  29. Kononenko NL & Haucke V (2015). Molecular mechanisms of presynaptic membrane retrieval and synaptic vesicle reformation. Neuron 85, 484–496. [DOI] [PubMed] [Google Scholar]
  30. Kononenko NL, Puchkov D, Classen GA, Walter AM, Pechstein A, Sawade L, Kaempf N, Trimbuch T, Lorenz D, Rosenmund C, Maritzen T & Haucke V (2014). Clathrin/AP‐2 mediate synaptic vesicle reformation from endosome‐like vacuoles but are not essential for membrane retrieval at central synapses. Neuron 82, 981–988. [DOI] [PubMed] [Google Scholar]
  31. Koo SJ, Kochlamazashvili G, Rost B, Puchkov D, Gimber N, Lehmann M, Tadeus G, Schmoranzer J, Rosenmund C, Haucke V & Maritzen T (2015). Vesicular synaptobrevin/VAMP2 levels guarded by AP180 control efficient neurotransmission. Neuron 88, 330–344. [DOI] [PubMed] [Google Scholar]
  32. Korogod N, Lou X & Schneggenburger R (2005). Presynaptic Ca2+ requirements and developmental regulation of posttetanic potentiation at the calyx of Held. J Neurosci 25, 5127–5137. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Korogod N, Lou X & Schneggenburger R (2007). Posttetanic potentiation critically depends on an enhanced Ca2+ sensitivity of vesicle fusion mediated by presynaptic PKC. Proc Natl Acad Sci USA 104, 15923–15928. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Kushmerick C, Renden R & von Gersdorff H (2006). Physiological temperatures reduce the rate of vesicle pool depletion and short‐term depression via an acceleration of vesicle recruitment. J Neurosci 26, 1366–1377. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Lee JS, Kim MH, Ho WK & Lee SH (2008). Presynaptic release probability and readily releasable pool size are regulated by two independent mechanisms during posttetanic potentiation at the calyx of Held synapse. J Neurosci 28, 7945–7953. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Li GL, Keen E, Andor‐Ardo D, Hudspeth AJ & von Gersdorff H (2009). The unitary event underlying multiquantal EPSCs at a hair cell's ribbon synapse. J Neurosci 29, 7558–7568. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Llano I, Gonzalez J, Caputo C, Lai FA, Blayney LM, Tan YP & Marty A (2000). Presynaptic calcium stores underlie large‐amplitude miniature IPSCs and spontaneous calcium transients. Nat Neurosci 3, 1256–1265. [DOI] [PubMed] [Google Scholar]
  38. Lou X, Fan F, Messa M, Raimondi A, Wu Y, Looger LL, Ferguson SM & De Camilli P (2012). Reduced release probability prevents vesicle depletion and transmission failure at dynamin mutant synapses. Proc Natl Acad Sci USA 109, E515–E523. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Lou X, Korogod N, Brose N & Schneggenburger R (2008. a). Phorbol esters modulate spontaneous and Ca2+‐evoked transmitter release via acting on both Munc13 and protein kinase C. J Neurosci 28, 8257–8267. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Lou X, Paradise S, Ferguson SM & De Camilli P (2008. b). Selective saturation of slow endocytosis at a giant glutamatergic central synapse lacking dynamin 1. Proc Natl Acad Sci USA 105, 17555–17560. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Lou X, Scheuss V & Schneggenburger R (2005). Allosteric modulation of the presynaptic Ca2+ sensor for vesicle fusion. Nature 435, 497–501. [DOI] [PubMed] [Google Scholar]
  42. Mahapatra S, Fan F & Lou X (2016). Tissue‐specific dynamin‐1 deletion at the calyx of Held decreases short‐term synaptic depression through a mechanism distinct from vesicle resupply. Proc Natl Acad Sci USA 113, E3150–3158. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Matthews G & Sterling PC (2008). Evidence that vesicles undergo compound fusion on the synaptic ribbon. J Neurosci 28, 5403–5411. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Neher E ( 2015). Merits and limitations of vesicle pool models in view of heterogeneous populations of synaptic vesicles. Neuron 87, 1131–1142. [DOI] [PubMed] [Google Scholar]
  45. Neher E & Sakaba T (2001). Estimating transmitter release rates from postsynaptic current fluctuations. J Neurosci 21, 9638–9654. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Nguyen TH, Maucort G, Sullivan RK, Schenning M, Lavidis NA, McCluskey A, Robinson PJ & Meunier FA (2012). Actin‐ and dynamin‐dependent maturation of bulk endocytosis restores neurotransmission following synaptic depletion. PLoS One 7, e36913. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Postlethwaite M, Hennig MH, Steinert JR, Graham BP & Forsythe IDCP (2007). Acceleration of AMPA receptor kinetics underlies temperature‐dependent changes in synaptic strength at the rat calyx of Held. J Physiol 579, 69–84. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Pyott SJ & Rosenmund C (2002). The effects of temperature on vesicular supply and release in autaptic cultures of rat and mouse hippocampal neurons. J Physiol 539, 523–535. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Raimondi A, Ferguson SM, Lou X, Armbruster M, Paradise S, Giovedi S, Messa M, Kono N, Takasaki J, Cappello V, O'Toole E, Ryan TA & De Camilli P (2011). Overlapping role of dynamin isoforms in synaptic vesicle endocytosis. Neuron 70, 1100–1114. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Renden R & von Gersdorff H (2007). Synaptic vesicle endocytosis at a CNS nerve terminal: faster kinetics at physiological temperatures and increased endocytotic capacity during maturation. J Neurophysiol 98, 3349–3359. [DOI] [PubMed] [Google Scholar]
  51. Richards DA, Rizzoli SO & Betz WJ (2004). Effects of wortmannin and latrunculin A on slow endocytosis at the frog neuromuscular junction. J Physiol 557, 77–91. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Saheki Y & De Camilli P (2012). Synaptic vesicle endocytosis. Cold Spring Harb Perspect Biol 4, a005645. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Sato K, Ernstrom GG, Watanabe S, Weimer RM, Chen CH, Sato M, Siddiqui A, Jorgensen EM & Grant BD (2009). Differential requirements for clathrin in receptor‐mediated endocytosis and maintenance of synaptic vesicle pools. Proc Natl Acad Sci USA 106, 1139–1144. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Schneggenburger R & Forsythe ID (2006). The calyx of Held. Cell Tissue Res 326, 311–337. [DOI] [PubMed] [Google Scholar]
  55. Schneggenburger R, Meyer AC & Neher E (1999). Released fraction and total size of a pool of immediately available transmitter quanta at a calyx synapse. Neuron 23, 399–409. [DOI] [PubMed] [Google Scholar]
  56. Schneggenburger R, Sakaba T & Neher E (2002). Vesicle pools and short‐term synaptic depression: lessons from a large synapse. Trends Neurosci 25, 206–212. [DOI] [PubMed] [Google Scholar]
  57. Shupliakov O, Bloom O, Gustafsson JS, Kjaerulff O, Low P, Tomilin N, Pieribone VA, Greengard P & Brodin L (2002). Impaired recycling of synaptic vesicles after acute perturbation of the presynaptic actin cytoskeleton. Proc Natl Acad Sci USA 99, 14476–14481. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Singer JH, Lassova L, Vardi N & Diamond JS (2004). Coordinated multivesicular release at a mammalian ribbon synapse. Nat Neurosci 7, 826–833. [DOI] [PubMed] [Google Scholar]
  59. Taschenberger H, Leao RM, Rowland KC, Spirou GA & von Gersdorff H (2002). Optimizing synaptic architecture and efficiency for high‐frequency transmission. Neuron 36, 1127–1143. [DOI] [PubMed] [Google Scholar]
  60. Thanawala MS & Regehr WG (2013). Presynaptic calcium influx controls neurotransmitter release in part by regulating the effective size of the readily releasable pool. J Neurosci 33, 4625–4633. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Voiculescu O, Charnay P & Schneider‐Maunoury S (2000). Expression pattern of a Krox‐20/Cre knock‐in allele in the developing hindbrain, bones, and peripheral nervous system. Genesis 26, 123–126. [DOI] [PubMed] [Google Scholar]
  62. Wadiche JI & Jahr CE (2001). Multivesicular release at climbing fibre‐Purkinje cell synapses. Neuron 32, 301–313. [DOI] [PubMed] [Google Scholar]
  63. Watanabe S, Rost BR, Camacho‐Perez M, Davis MW, Sohl‐Kielczynski B, Rosenmund C & Jorgensen EM (2013). Ultrafast endocytosis at mouse hippocampal synapses. Nature 504, 242–247. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Watanabe S, Trimbuch T, Camacho‐Perez M, Rost BR, Brokowski B, Sohl‐Kielczynski B, Felies A, Davis MW, Rosenmund C & Jorgensen EM (2014). Clathrin regenerates synaptic vesicles from endosomes. Nature 515, 228–233. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Wong AY, Graham BP, Billups B & Forsythe ID (2003). Distinguishing between presynaptic and postsynaptic mechanisms of short‐term depression during action potential trains. J Neurosci 23, 4868–4877. [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Wu Y, O'Toole ET, Girard M, Ritter B, Messa M, Liu X, McPherson PS, Ferguson SM & De Camilli P (2014). A dynamin 1‐, dynamin 3‐ and clathrin‐independent pathway of synaptic vesicle recycling mediated by bulk endocytosis. Elife 3, e01621. [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Xue L & Wu LG (2010). Post‐tetanic potentiation is caused by two signalling mechanisms affecting quantal size and quantal content. J Physiol 588, 4987–4994. [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Zhang B, Koh YH, Beckstead RB, Budnik V, Ganetzky B & Bellen HJ (1998). Synaptic vesicle size and number are regulated by a clathrin adaptor protein required for endocytosis. Neuron 21, 1465–1475. [DOI] [PubMed] [Google Scholar]
  69. Zucker RS & Regehr WG (2002). Short‐term synaptic plasticity. Annu Rev Physiol 64, 355–405. [DOI] [PubMed] [Google Scholar]

Articles from The Journal of Physiology are provided here courtesy of The Physiological Society

RESOURCES