Skip to main content
NCBI home page
Proceedings of the National Academy of Sciences of the United States of America logoLink to Proceedings of the National Academy of Sciences of the United States of America
. 2006 Nov 16;103(48):18332–18337. doi: 10.1073/pnas.0608507103

Munc18-1 expression levels control synapse recovery by regulating readily releasable pool size

Ruud F G Toonen *, Keimpe Wierda *,†, Michèle S Sons ‡, Heidi de Wit *, L Niels Cornelisse *, Arjen Brussaard §, Jaap J Plomp ‡, Matthijs Verhage *,†,
PMCID: PMC1838751  PMID: 17110441

Abstract

Prompt recovery after intense activity is an essential feature of most mammalian synapses. Here we show that synapses with reduced expression of the presynaptic gene munc18-1 suffer from increased depression during intense stimulation at glutamatergic, GABAergic, and neuromuscular synapses. Conversely, munc18-1 overexpression makes these synapses recover faster. Concomitant changes in the readily releasable vesicle pool and its refill kinetics were found. The number of vesicles docked at the active zone and the total number of vesicles per terminal correlated with both munc18-1 expression levels and the size of the releasable vesicle pool. These data show that varying expression of a single gene controls synaptic recovery by modulating the number of docked, release-ready vesicles and thereby replenishment of the secretion capacity.

Keywords: autapse, docking, exocytosis, secretion, synaptic transmission

Reliable and sustainable neurotransmitter release is essential for effective neuronal communication. However, neurons only have a limited number of fusion-ready vesicles that reside in a vesicle pool at the membrane of the presynaptic terminal (1). During periods of increased activity, this vesicle pool is depleted, resulting in a decreased reliability of neurotransmission. To ensure efficient neurotransmission, neurons need to be able to increase the initial number of fusion-ready vesicles [the so-called readily releasable pool (RRP)] and/or the rate at which this pool is replenished during activity. However, surprisingly little is known about the molecular mechanisms that control the size of the RRP and the way vesicles are recruited to this pool.

The Sec1/Munc18-like (SM) protein Munc18-1 has emerged as a key component for calcium-dependent neurotransmitter release (2). SM proteins function in all intracellular membrane trafficking pathways across species. Genetic deletion of Munc18-1 and most other SM genes involved in synaptic-vesicle release across species results in a complete block of neurotransmitter release (3–5), which shows that Munc18-1 and probably all SM proteins are indispensable factors that promote vesicle secretion (2, 6, 7). However, identifying where SM proteins act in the cascade of events leading to the release of neurotransmitter has proven to be difficult and has generated apparently conflicting data (8–10).

Here, we analyzed the effect of different Munc18-1 expression levels on synaptic function in autaptic synapses of GABAergic and glutamatergic central neurons, as well as in the peripheral neuromuscular junction (NMJ). We combined electrophysiological and optical measurements to show that Munc18-1 controls synapse efficacy in a bidirectional way via the control of the size and replenishment rate of the RRP.

Results

In homozygous munc18-1-null mutant mice, synapses are silent (3), identifying munc18-1 as an essential gene but providing little information on its molecular function. Heterozygous mice (munc18-1+/−) had a 50% reduction of Munc18-1 protein expression but no reduction in the levels of any of its known binding partners or 22 other synaptic proteins (Table 1, which is published as supporting information on the PNAS web site) (11). Autaptic cultures from these mice had similar dendrite length and number of synapses as cultures from WT littermates (Fig. 5, which is published as supporting information on the PNAS web site).

Munc18-1 Heterozygous Autapses Contain a Smaller Pool of Readily Releasable Vesicles.

Whole-cell recordings of autaptic glutamatergic or GABAergic munc18+/− and WT littermate neurons showed similar excitatory and inhibitory postsynaptic currents (EPSCs and IPSCs) upon single depolarizations (Figs. 5B and 6C, which are published as supporting information on the PNAS web site). The characteristics of spontaneous miniature (m) excitatory and inhibitory postsynaptic events (amplitude, frequency, and decay time) also did not differ between munc18+/− and WT neurons (Figs. 5 C and D and 6 A and B). Thus, a reduction of Munc18-1 protein level does not affect synaptic physiology under basal conditions nor does it appear to influence postsynaptic receptor number or sensitivity.

However, repeated stimulation produced a more pronounced rundown of evoked responses (synaptic depression) in munc18+/− neurons compared with WT neurons. This increased synaptic depression was observed both in glutamatergic and in GABAergic synapses and was most pronounced at 10-Hz stimulation (Fig. 1A and Fig. 6D). The rundown kinetics of glutamatergic synapses at 10 Hz were best characterized by biexponential curve fits (12), which revealed an increased rundown especially of the slow phase (τslow) in munc18-1+/− neurons. At 40-Hz stimulation, 80% depression was reached within 1 s in all groups, and a significant increase in depression in munc18+/− neurons was observed in GABAergic (Fig. 6E) but not in glutamatergic neurons (Fig. 7, which is published as supporting information on the PNAS web site). Differences in synaptic release probability, RRP size, and replenishment rate all may contribute to the observed increase in synaptic depression. To test RRP size, we applied hypertonic sucrose solution to empty the RRP via a Ca2+-independent mechanism (13, 14). The sucrose response in munc18-1+/− neurons was significantly smaller (0.67 ± 0.1 nC, n = 30) compared with WT neurons (1.06 ± 0.1 nC, n = 30, P < 0.01; Fig. 1B). To test whether a reduction of Munc18-1 also affected the RRP refilling, we used two approaches. First, we depleted the RRP with 40-Hz stimulation and measured the recovery by using 0.5-Hz stimulations (15). The 40-Hz stimulation resulted in similar depletion of the RRP in both genotypes (see also Fig. 7), and recovery from RRP depletion was not significantly slower in munc18-1+/− neurons (Fig. 1C). Second, we applied paired pulses of hypertonic sucrose with different time intervals between pulses (16). Again, no difference in RRP recovery was observed between WT and munc18-1+/− neurons (Fig. 1D).

Fig. 1.

Fig. 1.

Open in a new tab

Synaptic transmission in munc18-1 heterozygous autaptic neurons. (A) Synaptic rundown of glutamatergic EPSCs during 10-Hz stimulation is faster in munc18+/− compared with WT. Rundown kinetics were best characterized with biexponential fits and revealed that the slow component of the rundown was decreased in munc18+/− neurons. EPSC τfast = 0.12 ± 0.03, τslow = 3.93 ± 0.87 for munc18+/− neurons, and τfast = 0.12 ± 0.01, τslow = 6.86 ± 1.11 for WT (n = 11 and n = 20, P < 0.05 for τslow). Averaged weights of τfast and τslow were not different between WT and munc18+/−. (Insets) The first and last EPSC of the 10-Hz stimulation. For clarity, the stimulus artifact (see B Inset) was blanked from the traces. (B) Hypertonic sucrose (500 mM) application shows a 36% decrease in RRP size in munc18+/− neurons compared with WT neurons (WT: 1.06 ± 0.1 nC, n = 30; munc18+/−: 0.67 ± 0.09 nC, n = 30, P < 0.01). (Inset) Example traces during 500 mM sucrose application. (C) Activity- and calcium-dependent refill kinetics of the RRP after depletion of the pool by 2.5-s stimulation at 40 Hz are similar between WT and munc18+/− neurons (WT recovery: τ = 2.9 ± 0.4 s, n = 20; munc18+/− recovery: τ = 2.6 ± 0.3 s, n = 21, P > 0.05). (Insets) Individual WT traces during the paradigm. (D) Activity- and calcium-independent refill kinetics of the RRP tested by paired sucrose application with different interstimulus intervals are not different between WT and munc18+/− neurons (number of cells is in brackets, no significant difference at each of the different time points tested). The response of the second stimulus is plotted as a percentage of the first stimulus. (Inset) Typical responses to two sucrose applications with 4-s interval for WT and munc18+/− neurons.

Thus, Munc18-1 levels are rate-limiting during high-frequency neurotransmission. A 50% reduction in protein levels results in a reduction of the RRP size without affecting the rate by which this (smaller) pool is replenished.

Munc18-1 Heterozygous Mice Have Impaired Neuromuscular Synaptic Function.

Studies on SM proteins in Drosophila and Caenorhabditis elegans have been conducted on neuromuscular synapses (5, 17). To be able to directly compare our results, we performed electrophysiological recordings on diaphragm NMJs of munc18+/− mice and WT littermates. To test whether munc18+/− NMJs, like the autaptic cultures, were impaired in sustaining vesicle release during periods of high activity, we applied a high-rate (40-Hz) nerve-stimulation protocol. Munc18+/− NMJs displayed a significantly larger rundown of evoked endplate potential (EPP) amplitudes (to 67% of the first EPP compared with 78% for WT, P < 0.01; Fig. 2A).

Fig. 2.

Fig. 2.

Open in a new tab

Synaptic transmission at NMJs of munc18-1 heterozygote, WT, and Munc18-1-overexpressing littermates. (A) Munc18+/− mice are less able to sustain high-frequency evoked transmitter release at neuromuscular synapses. Indicated are the amplitudes of synaptic responses (EPPs) to each individual stimulus for 40 stimuli delivered to the phrenic nerve at 40 Hz, expressed as percentage of the first response. Data represent means ± SEM of five animals per group and 15 NMJs sampled per animal. (Inset) A typical example of the 40-Hz EPP rundown. No gross morphological differences were observed between NMJs of the two genotypes (Fig. 10 E and D). (B) Typical examples of MEPP frequency recordings in WT and munc18+/− NMJs. (C) Several physiological parameters in munc18+/− and WT NMJs. WT value was set at 100%. Where applicable, absolute values are indicated. Data represent means ± SEM of eight to nine animals per group and 10–15 NMJs sampled per animal. Differences between the groups were statistically significant for quantal content (P < 0.01), MEPP frequency (P < 0.05), sucrose response (P < 0.05), and statistical releasable pool n (P < 0.01). (D) Spontaneous MEPPs in munc18OE and WT mice (Upper, 10 s). The amplitude and kinetics of MEPPs were similar at WT and munc18OE NMJs (Lower). (E) Physiological parameters at NMJs of munc18OE and WT mice. WT value was set at 100%. Where applicable, absolute values are indicated. Data represent means ± SEM of 10–11 animals per group and 10–15 NMJs sampled per animal. Differences between groups were significant for quantal content (P < 0.05), MEPP frequency (P < 0.001), and statistical releasable pool n (P < 0.05).

As in autapses, we tested whether the EPP rundown could be explained by a reduction in RRP size by applying 500 mM sucrose. The response, measured as miniature endplate potential (MEPP) frequency, was 31% lower at munc18-1+/− endplates (P < 0.05; Fig. 2C). As an alternative approach, we used the calculation method of Miyamoto (18) to estimate RRP size and release probability from our EPP data. This method showed that the release probability was unchanged at munc18-1+/− NMJs, whereas the RRP size was reduced by 21% (P < 0.01; Fig. 2C and Table 2, which is published as supporting information on the PNAS web site). In addition, in munc18-1+/− NMJs, the frequency of spontaneous uniquantal acetylcholine release events (measured as MEPP frequency) and the 0.3-Hz evoked release (quantal content) were reduced by 20–25% (P < 0.05 and < 0.01, respectively; Fig. 2C). These single synapse recordings reveal a reduction in the RRP size as well as a concomitant decrease in evoked release (quantal content). Given that the quantal content is the product of the size of the RRP and the probability that a vesicle is released upon stimulation (19), these data suggest that Munc18-1 does not substantially influence vesicular release probability.

Munc18-1 Overexpression Results in a Larger RRP and Enhances Activity-Dependent RRP Replenishment.

To investigate the effect of Munc18-1 overexpression on synaptic-vesicle release, we applied two viral-expression systems in autaptic cultures. We tested the effect of acute, high overexpression of Munc18-1 with the Semliki Forest virus system, 6 to 8 h postinfection (20). In addition, we used a Lenti viral system to investigate the effect of long-term, moderate overexpression of Munc18-1, 10 to 14 days postinfection (21). Munc18-1 overexpression with either Semliki (SFVM18) or Lenti virus did not affect neuronal morphology or total synapse number in glutamatergic autaptic neurons (Fig. 5E). Also, evoked postsynaptic responses in SFVM18-overexpressing glutamatergic neurons did not differ from WT responses (Fig. 5F, EPSC). Effects of Munc18-1 overexpression on evoked responses may be masked by postsynaptic receptor saturation. Therefore, evoked responses were measured in the presence of the competitive NMDA/α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptor blocker kynurenic acid (22). In the presence of 200 μM kynurenic acid, the EPSC amplitude in WT and SFVM18 neurons was reduced to the same extent, indicating that saturation of postsynaptic receptors did not mask an effect in SFVM18 neurons (Fig. 8 A and B, which is published as supporting information on the PNAS web site). Evoked responses after SFVM18- or Lenti virus-mediated Munc18-1 overexpression in GABAergic neurons also were similar to WT (IPSC, Fig. 9A, which is published as supporting information on the PNAS web site). Thus, acute as well as chronic overexpression of Munc18-1 does not affect basal synaptic strength or synapse formation in autaptic cultures.

To test whether overexpression of Munc18-1 increases RRP size, we applied hypertonic sucrose (500 mM). The response from SFVM18-overexpressing neurons was not significantly different from WT neurons (SFVM18: 1.3 ± 0.2 nC, n = 26; WT: 1.1 ± 0.1 nC, n = 30, P = 0.4; Fig. 3A). Again, postsynaptic receptor saturation could mask potential effects of Munc18-1 overexpression. Therefore, we conducted three additional experiments. First, we applied a milder osmotic stimulus (200 mM sucrose) known to result in submaximal RRP depletion (14) to WT, munc18+/−-, and SFVM18-overexpressing neurons. This stimulus resulted in a >2-fold higher response in Munc18-overexpressing neurons compared with WT, and, consistent with the data in Fig. 1, Munc18-1+/− neurons showed a significantly smaller response (SFVM18: 0.17 ± 0.041 nC, n = 16; munc18+/−: 0.035 ± 0.011 nC, n = 11; and WT: 0.074 ± 0.011 nC, n = 11, P < 0.05 between WT and SFVM18; Fig. 3B). This result suggests that application of 500 mM sucrose indeed saturated postsynaptic receptors, masking an increase in RRP size. Second, we applied 500 mM sucrose in the presence of 200 μM kynurenic acid (see above). Consistent with the first experiment, an almost 2-fold difference in RRP size now became evident (WT: 0.95 ± 0.14 nC; and WT plus kynurenic acid: 0.63 ± 0.1 nC, P < 0.05; SFVM18: 1.04 ± 0.10 nC; and SFVM18 plus kynurenic acid: 0.96 ± 0.08, P = 0.5; Fig. 3C). Third, we directly labeled the RRP by using the styryl dye FM4-64 in combination with 500 mM sucrose application (23, 24). This direct presynaptic assessment also showed an ≈2-fold higher fluorescence intensity on Munc18-1 overexpression (Fig. 3D) Together, these independent lines of evidence confirm that Munc18-1 overexpression indeed leads to a larger initial RRP size.

Fig. 3.

Fig. 3.

Open in a new tab

Synaptic transmission in Munc18-1-overexpressing autaptic neurons. (A) Single 500 mM sucrose response is not significantly different between WT and SFVM18 overexpression (WT: 1.1 ± 0.1 nC, n = 39; SFVM18: 1.3 ± 0.2 nC, n = 35, P = 0.4). (B) Single 200 mM sucrose application reveals an increased RRP size in SFVM18-overexpressing neurons (WT: 0.074 ± 0.011 nC, n = 11; munc18+/−: 0.035 ± 0.011 nC, n = 11; SFVM18: 0.17 ± 0.041 nC, n = 16, P < 0.05 between WT and SFVM18 and P < 0.01 between SFVM18 and munc18+/− neurons). (Inset) Typical responses to 200 mM sucrose application for the three genotypes tested. (C) Single 500 mM sucrose application in the presence of 200 μM of the NMDA/α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptor blocker kynurenic acid results in an expected decreased response in WT neurons, whereas the response of SFVM18-overexpressing neurons is unaffected (WT minus kynurenic acid: 0.95 ± 0.14 nC, n = 15 and WT plus kynurenic acid: 0.63 ± 0.1 nC, n = 15, P < 0.05; SFVM18 minus kynurenic acid: 1.04 ± 0.1 nC, n = 15 and SFVM18 plus kynurenic acid: 0.96 ± 0.08, n = 15, P = 0.5). This finding shows that the increased release of glutamate in SFVM18-overexpressing neurons leads to receptor saturation and indicates that the RRP is increased on Munc18-1 overexpression. (D) Direct labeling of the RRP by using 500 mM sucrose solution containing 16 μM FM4-64 reveals a 2-fold larger RRP in SFVM18-overexpressing neurons compared with WT. Shown are average arbitrary fluorescent units (a.u.) from 769 synapses on 18 neurons for WT and 960 synapses on 18 neurons for SFVM18 from four independent experiments (WT: 91 ± 7.6 a.u., SFVM18: 191 ± 10.8 a.u., P < 0.01 with n = 4). (Insets) Examples of WT and SFVM18-overexpressing presynaptic terminals labeled with FM4-64 (red) by using 500 mM sucrose on EGFP (green)-filled dendrites. (Bar: 5 μm.) (E) Munc18-1 overexpression increases the recovery rate after activity-dependent RRP depletion. EPSC amplitude was sampled at a frequency of 0.5 Hz after depletion of the RRP with a 40-Hz stimulation train for 2.5 s. (Insets) Individual WT and SFVM18 traces during the paradigm. (F) Single exponential fits of RRP recovery show a significant faster replenishment in neurons overexpressing Munc18-1 (SFV Munc18-1: τ = 1.8 ± 0.2 s, n = 15; WT: τ = 2.9 ± 0.4 s, n = 20, P < 0.05). (G) Munc18-1 overexpression decreases synaptic rundown during high-frequency stimulation (40-Hz, 2.5 s). (H) Single exponential fits of the synaptic rundown show a significant decrease in synaptic rundown in neurons overexpressing Munc18-1 (SFV Munc18-1: τ = 0.57 ± 0.1 s, n = 15, WT: τ = 0.27 ± 0.04 s, n = 20, P = 0.003). (I) mEPSC frequency is significantly increased on Munc18-1 overexpression (WT: 20.3 ± 3.9 Hz, n = 11 cells, 2,300 events; SFVM18: 51.6 ± 7.0 Hz, n = 12 cells, 2,000 events, P < 0.001). Miniature amplitude and decay time are not affected by Munc18-1 overexpression (WT: 28.5 ± 3.4 pA, 2.28 ± 0.12 ms; SFVM18: 35.7 ± 3.5 pA, 2.40 ± 0.15 ms, P > 0.05 for both parameters). (J) Example traces of spontaneous glutamatergic release in WT and SFVM18-overexpressing neurons.

Next, we examined whether Munc18-1 overexpression also resulted in a smaller rundown and faster RRP replenishment. Indeed, synapses overexpressing Munc18-1 showed less synaptic depression during high-frequency stimulation (Fig. 3 G and H, EPSC rundown; 40 Hz) and an accelerated refilling of the RRP after high-rate (40-Hz) electrical stimulation (Fig. 3E; SFV Munc18-1: τ = 1.8 ± 0.2 s and WT: τ = 2.9 ± 0.4 s, P < 0.05). The same effect was observed in GABAergic neurons infected with either SFVM18 or Lenti virus Munc18-1 (Fig. 9 B and C). We ruled out any contribution of postsynaptic receptor desensitization by repeating the experiment in the presence of 50 μM cyclothiazide, a drug that blocks glutamate receptor desensitization (Fig. 10, which is published as supporting information on the PNAS web site). In contrast to the faster RRP replenishment after activity-dependent depletion, paired sucrose application did not reveal an effect of Munc18-1 overexpression on the rate of RRP replenishment when tested in the presence of 200 μM kynurenic acid (Fig. 8 C and D). Thus, Munc18-1 overexpression increases the RRP size and its replenishment, probably by activity- and/or Ca2+-dependent mechanisms. As a result, synapses recover faster. Consistent with an increase in RRP size, Munc18-1 overexpression also led to a >2-fold increase in the frequency of spontaneous vesicle release in both glutamatergic and GABAergic neurons, without affecting postsynaptic receptor sensitivity and/or numbers (Fig. 3 I and J, mEPSCs, and Fig. 9 D and L, mIPSCs).

Munc18-1-Overexpressing Mice Have Enhanced Synaptic Function in NMJs.

We analyzed diaphragm NMJs of Munc18-1-overexpressing mice (munc18OE, Fig. 11, which is published as supporting information on the PNAS web site). We first assessed RRP size with 500 mM sucrose. The sucrose-induced MEPP frequency was not significantly different between munc18OE and WT NMJs (P = 0.16; Fig. 2E). Both spontaneous uniquantal acetylcholine release (MEPP frequency) and evoked release (quantal content at 0.3-Hz stimulation) were increased compared with WT controls, by 53% and 14%, respectively (P < 0.001 and P < 0.05; Fig. 2 D and E). As in munc18+/− mice, no differences were seen in resting membrane potential, quantal size (MEPP amplitude), upward or downward slopes of spontaneous events, and evoked response latency (Fig. 2D and data not shown). Hence, in accordance with autaptic neurons and in contrast to the robust decrease of EPP responses in Drosophila NMJs on overexpression of Rop (9), increased levels of Munc18-1 lead to an increase in synaptic efficacy of the NMJ without affecting the fusion process.

Munc18-1 Proteins Levels Control the Number of Docked Synaptic Vesicles.

In autaptic neurons, the RRP size correlates with the number of docked vesicles (25). Therefore, we performed quantitative electron microscopy on glutamatergic synapses at 14 days in vitro (DIV). At low magnification, the general appearance of autaptic cultures from munc18+/−, WT littermates, and WT neurons infected with SFV munc18-1 was similar, and asymmetrical synapses with a clear active zone and postsynaptic density were present in comparable numbers (Fig. 4A). The size of the active zone, postsynaptic density, and synaptic vesicle cluster perimeter did not differ between genotypes (Fig. 4B), indicating that Munc18-1 levels do not influence general synapse morphology or size. However, the number of vesicles in immediate contact with the presynaptic active zone membrane increased significantly with increasing munc18-1 expression levels (munc18+/−: 3.1 ± 0.6, n = 26; WT: 4.8 ± 0.3, n = 62; and SFVM18: 7.0 ± 0.4, n = 45; P < 0.05 for munc18+/− versus WT and P < 0.001 for WT versus SFVM18). Hence, these morphometric data show that Munc18-1 expression levels correlate with the number of docked synaptic vesicles per active zone area, which parallels the observed physiological changes in RRP size. We also quantified the number of vesicles within 150 nm (approximately three times the synaptic vesicle size) from the active zone. Again, a positive correlation between the number of vesicles present in this pool and Munc18-1 expression levels was found (Fig. 4B), which may explain the faster RRP replenishment. Finally, the total number of vesicles present in the synapse also increases when more Munc18-1 is present (Fig. 4B).

Fig. 4.

Fig. 4.

Open in a new tab

Munc18 protein levels control the number of docked synaptic vesicles. (A) Electron micrographs of typical asymmetrical glutamatergic synapses from munc18+/, WT, and SFVM18-overexpressing autaptic neurons. Hippocampal autaptic neurons were analyzed after 14 days in culture and 6 h after infection with SFV. (Scale bar: 200 nm.) (B) The number of vesicles docked at the active zone increases with increasing amounts of munc18-1 (munc18+/−: 3.1 ± 0.6, n = 26; WT: 4.8 ± 0.3, n = 62; and SFVM18: 7.0 ± 0.4, n = 45; P < 0.05 for munc18+/− vs. WT and P < 0.001 for WT vs. SFVM18), whereas the size of the active zone and the vesicle cluster perimeter do not significantly change (ANOVA, P > 0.05). The number of vesicles within 150 nm from the active zone (WT: 24.4 ± 0.9; SFVM18: 41.9 ± 2.6, P < 0.001) and total number of vesicles per synapse is higher in SFV Munc18-1-overexpressing synapses compared with WT (WT: 62.6 ± 3.8; SFVM18: 90.1 ± 7.2, P < 0.001) but not in munc18+/− neurons.

Discussion

In the present study we show that the efficacy of synaptic transmission at both excitatory and inhibitory synapses depends on Munc18-1 levels, and we identify changes in RRP size and replenishment as the primary underlying mechanism. This mechanism may be related to the way in which protein kinase C (26, 27) and ras GTPases (28) modulate synaptic transmission, but it is distinct from the two other major mechanisms to control synaptic efficacy in nerve terminals, which rely on changes in vesicle-release probability (29–31) or connectivity changes [changes in the number of synapses that connect two neurons and/or the size of the terminals (32)]. Although all three mechanisms can increase presynaptic output, only an increase in RRP size and replenishment warrants a sustained enhancement in synaptic efficacy.

Munc18-1-dependent changes in RRP size can be explained by the observed bidirectional changes in docked vesicles (fewer in munc18+/− neurons and more on Munc18-1 overexpression). These observations are in line with our previous observations in munc18-1 null mutant chromaffin cells (33) and neocortex at late embryonic stages (34) as well as with studies in C. elegans (5). Earlier in development, a docking defect was not observed in mouse neocortex (3). The docking phenotype suggests a role for Munc18-1 that is distinct from other presynaptic proteins like Munc13–1, rab3s, complexins, and Rim1α, which all influence synaptic efficacy through modulation of the release probability without affecting vesicle docking and number or their localization (29, 31, 35). The increase in docked vesicles on Munc18 overexpression may be a consequence of the increased total number of vesicles in the terminal. However, in previous studies, changes in total vesicle number did not lead to concomitant changes in the number of docked vesicles (5, 36), and Munc18 overexpression in chromaffin cells also increased docking but without affecting total vesicle numbers (37). Hence, often docking does not correlate with the total number of available vesicles, presumably because of the limited availability of docking sites. Therefore, we conclude that Munc18-1 specifically regulates the docking step, maybe by establishing such sites. In addition, Munc18-1 overexpression apparently influences the total number of vesicles in the synapse by an unknown mechanism. The increased total number of vesicles may contribute to the faster RRP replenishment, especially the increased number of vesicles within 150 nm from the membrane (Fig. 4B). Because the cluster perimeter and active zone area were unchanged (Fig. 4B), the vesicle concentration within the cluster must be higher, which may promote faster reloading and greater occupancy of docking sites. The unaltered responses to paired hypertonic shocks suggest that Munc18-1 overexpression increases replenishment in a calcium-dependent manner.

The conclusion that Munc18-1 regulates the docking step, together with the unaltered kinetics of spontaneous and evoked fusion events, both in autapses and NMJs, argue for a role of Munc18-1 upstream of the synaptic vesicle fusion process itself. A large body of evidence suggests that the interaction of soluble N-ethylmaleimide-sensitive factor attachment protein receptors (SNARE) proteins from vesicular and target membranes drives this fusion reaction (38–40). Deletion of SNAP-25 or synaptobrevin, two members of the SNARE complex involved in synaptic-vesicle release, results in a severe impairment of neurotransmission without affecting vesicle docking (40, 41). Hence, our data suggest that Munc18-1 functions upstream of SNARE complex formation. In vitro, Munc18-1 interacts tightly with the SNARE protein syntaxin 1 (42, 43). As Munc18-1 has no membrane interacting domains, the interaction with syntaxin 1 at the plasma membrane may be important for vesicle docking. Therefore, it seems plausible that the Munc18-1/syntaxin 1 dimer is involved in initial steps preceding vesicle fusion, the docking step, and that Munc18-1 subsequently is displaced to allow syntaxin to initiate SNARE complex formation and vesicle fusion.

Most of the effects of altered Munc18-1 expression described here can be explained by changes in the RRP size only (sucrose response, FM-dye loading, and vesicle distribution in electron micrographs). The increase in miniature frequency in Munc18-1-overexpressing neurons and the decrease in the munc18+/− NMJ are consistent with this conclusion. However, miniature frequency was not significantly different in munc18+/− autaptic neurons, as expected given the smaller RRP size in these cells, which is probably because of the fact that this frequency is rather variable in autapses. There is a trend between the groups (27% difference in group average; Fig. 5D); in fact, this is rather similar to the NMJ (29% difference; Fig. 2C). Increasing the release rate by using 200 mM sucrose (Fig. 3B) did result in the expected Munc18-1-dependent changes. Secondly, single evoked responses were normal in munc18+/− and Munc18-1-overexpressing neurons (Figs. 5B and F, 6C, and 9A) despite differences in RRP size. This finding suggests that the release probability of the vesicles released during these single stimuli is altered. The observed differences in synaptic depression upon repetitive stimulation (Figs. 1A, 2A, 3G, 6 D and E, and 9 F and G) are consistent with such an explanation, although in this case changes in RRP size may still contribute. On the other hand, the fact that, at the monosynaptic NMJ, changes in RRP size do correlate with changes in EPP amplitude and miniature frequency strengthens the explanation that Munc18-1 does not directly control release probability but primarily RRP size. It is becoming evident that synaptic transmission in central synapses cannot be described accurately unless heterogeneity of vesicular release probability is taken into account. The fact that single evoked responses are unaltered in autapses despite differences in RRP size also may be attributable to such heterogeneous release probability. The initial release probability per synapse may be adjusted to the optimal dynamic range (44) despite differences in RRP size and the release probability of the first vesicle, released on a single stimulus, may be different from other releasable vesicles.

Materials and Methods

Transgenic Mice.

Two independent null-mutant mouse lines were produced for the munc18-1 gene as described (3). Mice were bred as heterozygotes by using standard mouse husbandry and backcrossed for at least six generations to a C57BL/6 background.

Neuromuscular Synapse Electrophysiology.

Measurements were performed on nerve/muscle preparations from the diaphragm of 2- to 5-month-old WT, munc18+/−, and munc18OE mice as described (45).

Cell Culture, Electrophysiology, and Viral Transduction.

Microisland cultures were prepared from munc18+/− and WT littermate embryos at embryonic day 18 according to ref. 14. Lenti viral particles containing Munc18-1 cDNA coupled to enhanced GFP (EGFP) via an internal ribosomal entry site (IRES), and particles containing IRES-EGFP as control were prepared according to ref. 21 and Semliki Forest particles were prepared as described in ref. 33. Neurons were infected at DIV 1 with Lenti virus or 6–8 h before electrophysiological recordings with Semliki Forest virus. Whole-cell voltage-clamp recordings (holding potential −70 mV) were performed on cultured neurons between DIV 12 and 15. All experiments were conducted at 32°C.

Microscopy and Fluorescence Imaging.

To selectively label RRP in WT and SFVM18-overexpressing neurons, cells were loaded with FM4-64 in calcium-free Tyrode's containing 500 mM sucrose for 3–4 s (16 μM FM4-64/500 mM sucrose/0 mM CaCl2/2.5 mM KCl/119 mM NaCl/3 mM MgCl2/30 mM glucose/25 mM Hepes, pH 7.4). The 500 mM sucrose-containing solution was replaced by calcium-free Tyrode's containing 16 μM FM4-64 for an additional 60 s to ensure labeling of all exocytosed vesicles. Cells were washed for 10 min with calcium-free Tyrode's. Both loading and washing were performed in calcium-free solution to reduce spontaneous release of vesicles and subsequent loss of fluorescence. All solutions contained 50 μM 2-amino-5-phosphonopentoic acid and 10 μM 6-cyano-7-nitroquinosaline-2,3-dione to prevent recurrent activity.

Electron Microscopy.

Hippocampal islands cultures of munc18+/− or littermate WT mice (embryonic day 18) obtained from four different litters were grown on BELLCO photo-etched grid coverslips (BELLCO Glass Inc., Vineland, NJ). WT hippocampal neurons were infected (DIV 14) with SFV munc18-1-IRES-EGFP or SFV IRES-EGFP as control and observed under a fluorescence microscope 6 h after infection to map the location of infected cells. As for electrophysiology, only glia islands containing a single neuron were used for analysis. Autaptic synapses were selected in serial ultrathin sections at low magnification by using a JEOL (Tokyo, Japan) 1010 electron microscope, and high-resolution images were acquired at ×100,000 magnification.

Statistical Analysis.

Data shown are mean values ± SEM. Statistical significance was determined by using Student's t test, and overall group differences were analyzed by using ANOVA.

Supporting Information.

For further information on these methods as well as detailed descriptions of autaptic electrophysiology, Northern blotting, in situ hybridization, and protein analyses, see Supporting Materials and Methods, which is published as supporting information on the PNAS web site.

Supplementary Material

Supporting Information
pnas_0608507103_index.html (36.5KB, html)

Acknowledgments

We thank Robbert Zalm, Desiree Schut, Hans Lodder, and Ineke Lavrijsen for invaluable technical assistance. This work was supported by Dutch Organization for Scientific Research Grants NWO-GBMW 903-42-073 (to J.J.P.) and NWO-GBMW 903-42-023; ZonMW Veni Grants 016-066-101 (to R.F.G.T.), GpD 970-10-036 (to M.V. and H.d.W.), and 916-36-043 (to H.d.W.); Zon-MW Pionier Grant MW-PIO900-01-001 (to M.V.); and NeuroBsik Mouse Phenomics Consortium (Grant BSIK03053).

Abbreviations

RRP

readily releasable pool

NMJ

neuromuscular junction

SM

Sec1/Munc18-like

EPSC

excitatory postsynaptic current

IPSC

inhibitory postsynaptic current

mEPSC

miniature EPSC

mIPSC

miniature IPSC

EPP

evoked endplate potential

MEPP

miniature endplate potential

DIV

days in vitro

SNARE

soluble N-ethylmaleimide-sensitive factor attachment protein receptors

EGFP

enhanced GFP

IRES

internal ribosomal entry site.

Footnotes

The authors declare no conflict of interest.

References

  • 1.Rizzoli SO, Betz WJ. Nat Rev Neurosci. 2005;6:57–69. doi: 10.1038/nrn1583. [DOI] [PubMed] [Google Scholar]
  • 2.Rizo J, Südhof TC. Nat Rev Neurosci. 2002;3:641–653. doi: 10.1038/nrn898. [DOI] [PubMed] [Google Scholar]
  • 3.Verhage M, Maia AS, Plomp JJ, Brussaard AB, Heeroma JH, Vermeer H, Toonen RF, Hammer RE, van den Berg TK, Missler M, et al. Science. 2000;287:864–869. doi: 10.1126/science.287.5454.864. [DOI] [PubMed] [Google Scholar]
  • 4.Harrison SD, Broadie K, van de Goor J, Rubin GM. Neuron. 1994;13:555–566. doi: 10.1016/0896-6273(94)90025-6. [DOI] [PubMed] [Google Scholar]
  • 5.Weimer RM, Richmond JE, Davis WS, Hadwiger G, Nonet ML, Jorgensen EM. Nat Neurosci. 2003;6:1023–1030. doi: 10.1038/nn1118. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Toonen RF, Verhage M. Trends Cell Biol. 2003;13:177–186. doi: 10.1016/s0962-8924(03)00031-x. [DOI] [PubMed] [Google Scholar]
  • 7.Südhof TC. Annu Rev Neurosci. 2004;27:509–547. doi: 10.1146/annurev.neuro.26.041002.131412. [DOI] [PubMed] [Google Scholar]
  • 8.Dresbach T, Burns ME, O'Connor V, DeBello WM, Betz H, Augustine GJ. J Neurosci. 1998;18:2923–2932. doi: 10.1523/JNEUROSCI.18-08-02923.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Schulze KL, Littleton JT, Salzberg A, Halachmi N, Stern M, Lev Z, Bellen HJ. Neuron. 1994;13:1099–1108. doi: 10.1016/0896-6273(94)90048-5. [DOI] [PubMed] [Google Scholar]
  • 10.Scott BL, Van Komen JS, Irshad H, Liu S, Wilson KA, McNew JA. J Cell Biol. 2004;167:75–85. doi: 10.1083/jcb.200405018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Toonen RF, de Vries KJ, Zalm R, Südhof TC, Verhage M. J Neurochem. 2005;93:1393–1400. doi: 10.1111/j.1471-4159.2005.03128.x. [DOI] [PubMed] [Google Scholar]
  • 12.Pyott SJ, Rosenmund C. J Physiol (London) 2002;539:523–535. doi: 10.1113/jphysiol.2001.013277. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Stevens CF, Tsujimoto T. Proc Natl Acad Sci USA. 1995;92:846–849. doi: 10.1073/pnas.92.3.846. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Rosenmund C, Stevens CF. Neuron. 1996;16:1197–1207. doi: 10.1016/s0896-6273(00)80146-4. [DOI] [PubMed] [Google Scholar]
  • 15.Rhee JS, Betz A, Pyott S, Reim K, Varoqueaux F, Augustin I, Hesse D, Südhof TC, Takahashi M, Rosenmund C, Brose N. Cell. 2002;108:121–133. doi: 10.1016/s0092-8674(01)00635-3. [DOI] [PubMed] [Google Scholar]
  • 16.Rosenmund C, Sigler A, Augustin I, Reim K, Brose N, Rhee JS. Neuron. 2002;33:411–424. doi: 10.1016/s0896-6273(02)00568-8. [DOI] [PubMed] [Google Scholar]
  • 17.Wu MN, Littleton JT, Bhat MA, Prokop A, Bellen HJ. EMBO J. 1998;17:127–139. doi: 10.1093/emboj/17.1.127. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Miyamoto MD. J Physiol (London) 1975;250:121–142. doi: 10.1113/jphysiol.1975.sp011045. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Del Castillo J, Katz B. J Physiol (London) 1954;124:560–573. doi: 10.1113/jphysiol.1954.sp005129. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Ashery U, Betz A, Xu T, Brose N, Rettig J. Eur J Cell Biol. 1999;78:525–532. doi: 10.1016/s0171-9335(99)80017-x. [DOI] [PubMed] [Google Scholar]
  • 21.Naldini L, Blomer U, Gallay P, Ory D, Mulligan R, Gage FH, Verma IM, Trono D. Science. 1996;272:263–267. doi: 10.1126/science.272.5259.263. [DOI] [PubMed] [Google Scholar]
  • 22.Meyer AC, Neher E, Schneggenburger R. J Neurosci. 2001;21:7889–7900. doi: 10.1523/JNEUROSCI.21-20-07889.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Pyle JL, Kavalali ET, Piedras-Renteria ES, Tsien RW. Neuron. 2000;28:221–231. doi: 10.1016/s0896-6273(00)00098-2. [DOI] [PubMed] [Google Scholar]
  • 24.Mozhayeva MG, Sara Y, Liu X, Kavalali ET. J Neurosci. 2002;22:654–665. doi: 10.1523/JNEUROSCI.22-03-00654.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Schikorski T, Stevens CF. Nat Neurosci. 2001;4:391–395. doi: 10.1038/86042. [DOI] [PubMed] [Google Scholar]
  • 26.Stevens CF, Sullivan JM. Neuron. 1998;21:885–893. doi: 10.1016/s0896-6273(00)80603-0. [DOI] [PubMed] [Google Scholar]
  • 27.Virmani T, Ertunc M, Sara Y, Mozhayeva M, Kavalali ET. J Neurosci. 2005;25:10922–10929. doi: 10.1523/JNEUROSCI.3766-05.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Kushner SA, Elgersma Y, Murphy GG, Jaarsma D, van Woerden GM, Hojjati MR, Cui Y, LeBoutillier JC, Marrone DF, Choi ES, et al. J Neurosci. 2005;25:9721–9734. doi: 10.1523/JNEUROSCI.2836-05.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Augustin I, Rosenmund C, Südhof TC, Brose N. Nature. 1999;400:457–461. doi: 10.1038/22768. [DOI] [PubMed] [Google Scholar]
  • 30.Geppert M, Goda Y, Stevens CF, Südhof TC. Nature. 1997;387:810–814. doi: 10.1038/42954. [DOI] [PubMed] [Google Scholar]
  • 31.Reim K, Mansour M, Varoqueaux F, McMahon HT, Südhof TC, Brose N, Rosenmund C. Cell. 2001;104:71–81. doi: 10.1016/s0092-8674(01)00192-1. [DOI] [PubMed] [Google Scholar]
  • 32.Tyler WJ, Pozzo-Miller LD. J Neurosci. 2001;21:4249–4258. doi: 10.1523/JNEUROSCI.21-12-04249.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Voets T, Toonen RF, Brian EC, de Wit H, Moser T, Rettig J, Südhof TC, Neher E, Verhage M. Neuron. 2001;31:581–591. doi: 10.1016/s0896-6273(01)00391-9. [DOI] [PubMed] [Google Scholar]
  • 34.Bouwman J, Maia AS, Camoletto PG, Posthuma G, Roubos EW, Oorschot VM, Klumperman J, Verhage M. Neuroscience. 2004;126:115–126. doi: 10.1016/j.neuroscience.2004.03.027. [DOI] [PubMed] [Google Scholar]
  • 35.Calakos N, Schoch S, Südhof TC, Malenka RC. Neuron. 2004;42:889–896. doi: 10.1016/j.neuron.2004.05.014. [DOI] [PubMed] [Google Scholar]
  • 36.Gitler D, Takagishi Y, Feng J, Ren Y, Rodriguiz RM, Wetsel WC, Greengard P, Augustine GJ. J Neurosci. 2004;24:11368–11380. doi: 10.1523/JNEUROSCI.3795-04.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Toonen RF, Kochubey O, de Wit H, Gulyas-Kovacs A, Konijnenburg B, Sorensen JB, Klingauf J, Verhage M. EMBO J. 2006;25:3725–3737. doi: 10.1038/sj.emboj.7601256. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Chen YA, Scales SJ, Patel SM, Doung YC, Scheller RH. Cell. 1999;97:165–174. doi: 10.1016/s0092-8674(00)80727-8. [DOI] [PubMed] [Google Scholar]
  • 39.Schoch S, Deak F, Konigstorfer A, Mozhayeva M, Sara Y, Südhof TC, Kavalali ET. Science. 2001;294:1117–1122. doi: 10.1126/science.1064335. [DOI] [PubMed] [Google Scholar]
  • 40.Washbourne P, Thompson PM, Carta M, Costa ET, Mathews JR, Lopez-Bendito G, Molnar Z, Becher MW, Valenzuela CF, Partridge LD, Wilson MC. Nat Neurosci. 2002;5:19–26. doi: 10.1038/nn783. [DOI] [PubMed] [Google Scholar]
  • 41.Schoch S, Deak F, Konigstorfer A, Mozhayeva M, Sara Y, Südhof TC, Kavalali ET. Science. 2001;294:1117–1122. doi: 10.1126/science.1064335. [DOI] [PubMed] [Google Scholar]
  • 42.Hata Y, Slaughter CA, Südhof TC. Nature. 1993;366:347–351. doi: 10.1038/366347a0. [DOI] [PubMed] [Google Scholar]
  • 43.Pevsner J, Hsu SC, Scheller RH. Proc Natl Acad Sci USA. 1994;91:1445–1449. doi: 10.1073/pnas.91.4.1445. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Burrone J, Murthy VN. Curr Opin Neurobiol. 2003;13:560–567. doi: 10.1016/j.conb.2003.09.007. [DOI] [PubMed] [Google Scholar]
  • 45.Sons MS, Plomp JJ. Brain Res. 2006;1089:126–134. doi: 10.1016/j.brainres.2006.03.055. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting Information
pnas_0608507103_index.html (36.5KB, html)
pnas_0608507103_1.pdf (1MB, pdf)
pnas_0608507103_2.pdf (248KB, pdf)
pnas_0608507103_3.pdf (508.1KB, pdf)
pnas_0608507103_4.pdf (1.2MB, pdf)
pnas_0608507103_5.pdf (348.4KB, pdf)
pnas_0608507103_6.pdf (15.2MB, pdf)
pnas_0608507103_08507Fig5.jpg (986.1KB, jpg)

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences

RESOURCES