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Reexamination of N-terminal domains of syntaxin-1 in vesicle fusion from central murine synapses

  1. Gülçin Vardar  Is a corresponding author
  2. Andrea Salazar-Lázaro
  3. Marisa Brockmann
  4. Marion Weber-Boyvat
  5. Sina Zobel
  6. Victor Wumbor-Apin Kumbol
  7. Thorsten Trimbuch
  8. Christian Rosenmund  Is a corresponding author
  1. Universität Berlin, Humboldt-Universität zu Berlin, Germany
  2. Berlin Institute of Health, Germany
  3. Einstein Center for Neurosciences Berlin, Germany
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Cite this article as: eLife 2021;10:e69498 doi: 10.7554/eLife.69498

Abstract

Syntaxin-1 (STX1) and Munc18-1 are two requisite components of synaptic vesicular release machinery, so much so synaptic transmission cannot proceed in their absence. They form a tight complex through two major binding modes: through STX1’s N-peptide and through STX1’s closed conformation driven by its Habc- domain. However, physiological roles of these two reportedly different binding modes in synapses are still controversial. Here we characterized the roles of STX1’s N-peptide, Habc-domain, and open conformation with and without N-peptide deletion using our STX1-null mouse model system and exogenous reintroduction of STX1A mutants. We show, on the contrary to the general view, that the Habc-domain is absolutely required and N-peptide is dispensable for synaptic transmission. However, STX1A’s N-peptide plays a regulatory role, particularly in the Ca2+-sensitivity and the short-term plasticity of vesicular release, whereas STX1’s open conformation governs the vesicle fusogenicity. Strikingly, we also show neurotransmitter release still proceeds when the two interaction modes between STX1A and Munc18-1 are presumably intervened, necessitating a refinement of the conceptualization of STX1A–Munc18-1 interaction.

Introduction

The synaptic vesicle (SV) fusion is the fundamental process in synaptic transmission, and it is catalyzed by the merger of plasma and vesicular membranes by the neuronal SNAREs syntaxin-1 (STX1 collectively refers to STX1A and STX1B throughout this study), synaptobrevin-2 (Syb-2), and SNAP25 (Rizo and Sudhof, 2012; Rizo and Xu, 2015; Baker and Hughson, 2016). STX1 is the most important neuronal SNARE because not only synaptic transmission grinds to a halt in its absence, but also neurons cannot survive (Vardar et al., 2016). Compared to the other SNAREs, it also has a unique structure with its regulatory region composed of a bulky three helical Habc-domain and a short N-peptide preceding its SNARE motif (Figure 1A; Fernandez et al., 1998).

Figure 1 with 1 supplement see all
STX1A’s Habc-domain is essential and N-peptide is dispensable for neurotransmitter release.

(A) Domain structure of STX1A. The protein consists of a short N-peptide (aa 1–9 or 1–28), Habc domain (aa 29–144) formed by three helices, Ha, Hb, and Hc, followed by the H3 helix (aa 189–259; SNARE domain) and a transmembrane region (aa 266–288; TMR). (B) Example images of immunofluorescence labeling for Bassoon, STX1A, and Munc18-1 shown as red, green, and blue, respectively, in the corresponding composite pseudocolored images obtained from high-density cultures of STX1-null hippocampal neurons either not rescued or rescued with STX1AWT, or STX1A∆2-9; STX1ALEOpen; or STX1A∆Habc. Scale bar: 10 µm (C, D) Quantification of the immunofluorescence intensity of STX1A and Munc18-1 as normalized to the immunofluorescence intensity of Bassoon in the same ROIs as shown in (B). The values were then normalized to the values obtained from STX1AWT neurons. (E) Example traces (left) and quantification of the amplitude (right) of EPSCs obtained from hippocampal autaptic STX1-null neurons either not rescued or rescued with STX1AWT, STX1B∆2-9, STX1ALEOpen, or STX1A∆Habc. (F) Example traces (left) and quantification of the charge transfer (right) of 500 mM sucrose-elicited readily releasable pools (RRPs) obtained from the same neurons as in (E). (G) Quantification of probability of vesicular release (Pvr) determined as the percentage of the RRP released upon one AP. (H) Example traces (left) and quantification of the frequency (right) of mEPSCs recorded at –70 mV. (I) Example traces (left) and quantification (right) of short-term plasticity (STP) determined by high-frequency stimulation at 10 Hz and normalized to the EPSC1 from the same neuron. Data information: the artifacts are blanked in example traces in (D) and (H). The example traces in (G) were filtered at 1 kHz. In (C–H), data points represent single observations, the bars represent the mean ± SEM. In (I), data points represent mean ± SEM. Red and black annotations (stars and n.s.) on the graphs show the significance comparisons to STX1-null and to STX1AWT rescue, respectively (nonparametric Kruskal–Wallis test followed by Dunn’s post hoc test, *p≤0.05, ***p≤0.001, ****p≤0.0001). Two-way ANOVA was applied for data in (I). The numerical values are summarized in Figure 1—source data 1.

Figure 1—source data 1

Quantification of the STX1AWT and mutant STX1A expression induced by lentiviral transduction of STX1-null neurons and the consequent neurotransmitter release properties.

https://cdn.elifesciences.org/articles/69498/elife-69498-fig1-data1-v4.xlsx

Besides its interaction with the other SNAREs, STX1 also binds to its cognate SM protein Munc18-1 forming a tight binary complex with an affinity in the nanomolar range (Pevsner et al., 1994; Burkhardt et al., 2008). Munc18-1, which is an assistor of SNARE-mediated vesicular release, is an equally important protein as its absence also leads to inhibition of synaptic transmission (Verhage et al., 2000). Two major modes for STX1 binding to Munc18-1 have been defined: one through its N-peptide, the other through its closed conformation driven by the intramolecular interaction between its Habc- and SNARE domains (Dulubova et al., 1999; Misura et al., 2000). However, several issues regarding these reportedly different binding modes of STX1 to Munc18-1 are still subjects of dispute.

It is evident that STX1’s Habc-domain is required for proper folding of STX1 and for proper co-recruitment of STX1–Munc18-1 complex to the active zone (AZ) (Han et al., 2009; Meijer et al., 2012; Vardar et al., 2020; Zhou et al., 2013), yet it has been deemed to play a secondary role in synaptic transmission, to the point that it is dispensable for vesicle fusion per se (Rathore et al., 2010; Shen et al., 2010; Meijer et al., 2012; Zhou et al., 2013). However, an increasing number of mutations discovered in the Habc-domain of STX1B in patients with epilepsy (Schubert et al., 2014; Wolking et al., 2019; Vardar et al., 2020) points to greater importance for this region in neurotransmitter release.

The physiological significance of Munc18-1 binding to STX1’s N-peptide is less clear, even though the general view leans towards its indispensability for synaptic transmission. Firstly, the STX1 N-peptide does not majorly contribute to its overall affinity for Munc18-1 (Burkhardt et al., 2008; Christie et al., 2012; Colbert et al., 2013), yet liposome fusion cannot proceed without the N-peptide in reconstitution experiments (Shen et al., 2007; Rathore et al., 2010; Shen et al., 2010). On the other hand, interfering with STX1-N-peptide–Munc18-1 interaction by mutations either on STX1 (Zhou et al., 2013; Park et al., 2016) or on Munc18-1 (Khvotchev et al., 2007; Shen et al., 2007; Han et al., 2009; Meijer et al., 2012) in synapses in diverse model systems disclosed either its essentiality or its dispensability. Thus, a collective consensus as to what function the binding of STX1’s highly conserved N-peptide to Munc18-1 plays in synaptic transmission has not been reached.

So far, the physiological roles of STX1’s N-peptide, Habc-domain, and open-closed conformation were not assessed in central synapses completely devoid of STX1. Rather, studies have been conducted either in synapses with normal STX1 expression but mutant Munc18-1 (Khvotchev et al., 2007; Meijer et al., 2012; Shen et al., 2018) or in synapses with only severely reduced expression of STX1 (Zhou et al., 2013). Furthermore, in vitro studies do not contain the full panel of native synaptic proteins and mostly do not use full-length STX1 (Shen et al., 2007; Rathore et al., 2010; Shen et al., 2010). Therefore, we addressed the contribution of different domains of STX1 to neurotransmission using our STX1-null mouse model system and exogenous reintroduction of STX1A mutants either lacking N-peptide or the Habc-domain, or STX1 mutants forced into the open conformation (LEOpen mutation) with or without an N-peptide deletion. We show that the Habc-domain is absolutely required for STX1’s stability and/or expression and thus neurotransmitter release. Furthermore, in contrast to the general view, we find that N-peptide is not indispensable for synaptic transmission; however, we propose that STX1’s N-peptide plays a regulatory role, particularly in the Ca2+-sensitivity of vesicular release and generally in vesicle fusion, which is only unmasked by STX1’s open conformation.

Results

STX1’s Habc-domain is essential and N-peptide is dispensable for neurotransmitter release

Vesicle fusion does not occur in the absence of STX1 (Vardar et al., 2016) providing a null background in terms of neurotransmitter release. Thus, we used STX1A constitutive, STX1B conditional knockout (STX1-null) mouse neurons and lentiviral expression of different STX1 mutants in conjunction with Cre recombinase (Vardar et al., 2016; Vardar et al., 2020) to study the structure–function relationship of STX1 domains. With the focus on the effects of different Munc18-1 binding modes, we expressed STX1A mutants either with the deletion of the N-peptide (STX1A∆N2-9) or the Habc-domain (∆29–144; STX1A∆Habc) or with the introduction of well-described LEOpen (L165A, E166A; STX1ALEOpen) mutation (Figure 1A).

Firstly, we utilized immunocytochemistry in high-density hippocampal neuronal culture to quantify the exogenous expression of STX1A∆N2-9, STX1A∆Habc, and STX1ALEOpen at presynaptic compartments as defined by Bassoon-positive puncta and normalized fluorescence signals to the signals caused by expression of STX1AWT, all in STX1-null neurons. As expected from previous studies (Meijer et al., 2012; Zhou et al., 2013), deletion of the N-peptide had no significant effect on STX1A expression compared to STX1AWT, whereas STX1A∆Habc did not produce a measurable signal (Figure 1B and C). Loss of STX1 leads to a severe reduction in Munc18-1 expression, which can be rescued by the expression of either STX1A or STX1B (Zhou et al., 2013; Vardar et al., 2016; Vardar et al., 2020). Consistent with the expected relative binding states of STX1A∆N2-9 and STX1A∆Habc to Munc18-1 (Burkhardt et al., 2008), N-peptide deletion did not cause a significant change in Munc18-1 expression at Bassoon positive puncta, whereas the Habc-domain deletion was unable to rescue Munc18-1 levels back to WT-like levels (Figure 1B and D). Rendering STX1B constitutively open by LEOpen mutation is also known to decrease STX1B as well as Munc18-1 levels (Gerber et al., 2008) and the expression of STX1ALEOpen was severely low and inefficient to rescue Munc18-1 levels (Figure 1B–D).

To assess how the manipulation of the different Munc18-1 binding domains of STX1A affect the release of presynaptic vesicles, we measured Ca2+-triggered and spontaneous vesicle fusion, vesicle priming, and short-term plasticity (STP) in autaptic hippocampal neurons using electrophysiology as described previously (Vardar et al., 2016; Vardar et al., 2020). Compared to STX1AWT neurons, STX1ALEOpen neurons exhibited a trend towards a 40% increase in EPSC (Figure 1E) and towards a 30% decrease in hypertonic-sucrose measured readily releasable pool (RRP) (Figure 1F), trending towards an approximately threefold increase in probability of vesicular release (Pvr) (Figure 1G). The increase in Pvr, though not significant, was also evident in the observed enhancement of short-term depression (Figure 1I) as well as in the trend towards increased mEPSC frequency (Figure 1H). These findings are consistent with the previous studies on the LEOpen mutation on STX1A or STX1B (Gerber et al., 2008; Zhou et al., 2013).

Surprisingly, loss of N-peptide of STX1A showed only a trend towards 30% decrease in Ca2+-evoked vesicular release (Figure 1E), but not its full arrest. Similarly, RRP and spontaneous neurotransmission, which is assessed by the frequency of single-vesicle release events, were not completely inhibited by N-peptide deletion, but only trended towards a decrease by 30 and 50%, respectively, (Figure 1F and H). Proportionally similar trends in the reduction of both EPSC and RRP resulted in comparable Pvr between STX1A∆N2-9 and STX1AWT neurons (Figure 1G). Despite the lack of a net difference in the Pvr, however, STX1A∆N2-9 neurons exhibited an altered STP in response to the 10 Hz stimulation, with no depression to latter stimuli (Figure 1I).

Previous studies have suggested that the Habc-domain of STX1A and particularly its interaction with Munc18-1 is dispensable for vesicle fusion both in vitro and in vivo (Rathore et al., 2010; Shen et al., 2010; Meijer et al., 2012; Zhou et al., 2013). However, our analysis of the neurotransmission properties of the STX1A∆Habc neurons in comparison to the STX1AWT neurons showed that STX1A∆Habc was incapable of rescuing neurotransmitter release as it produced no detectable EPSC, RRP, or mEPSC; a phenotype similar to the STX1-null neurons (Figure 1E–G).

STX1 Habc-domain is indispensable for neuronal viability and the organization of synaptic ultrastructure

STX1 has also an obligatory function in neuronal maintenance and complete loss of both STX1A and STX1B leads to neuronal death (Vardar et al., 2016). To address the overall functionality of STX1A, we assessed the survivability of the high-density cultured STX1-null neurons expressing STX1A∆Habc and determined the cell number at different time intervals starting at DIV 8 (Figure 2A–C), at which time point all the groups had an average of ~40 neurons per mm2 (Figure 2B). Then we calculated the ratio of the cell number at DIV 15, 22, and 29 to the cell number at DIV 8 as a read-out for neuronal viability. STX1-null neurons showed a dramatic loss between DIV 8 and DIV 15 (Figure 2C) as reported before (Vardar et al., 2016). Even though at DIV 15 the number of surviving STX1A∆Habc neurons was slightly but significantly higher compared to that in STX1-null group, eventually STX1A∆Habc failed to rescue neuronal survival as by DIV 22 almost all STX1A∆Habc neurons were dead (Figure 2C).

STX1’s Habc-domain is essential for the overall function of STX1A.

(A) Example images of high-density cultures of STX1-null, STX1AWT, and STX1A∆Habc hippocampal neurons at DIV 8, 15, 22, and 29 represented with immunofluorescent labeling of microtubule associated protein 2 (MAP2) . Red and green nuclei serve as a marker for NLS-RFP-P2A-Cre recombinase expression and for NLS-GFP-P2A-STX1A (either WT or mutants), respectively. Scale bar: 50 µm. (B) Quantification of neuronal density at DIV 8. (C) Quantification of the percentage of the surviving neurons at DIV 8, 15, 22, and 29 as normalized to the neuronal density at DIV 8 in the same well. (D) Example high-pressure freezing fixation combined with electron microscopy (HPF-EM) images of nerve terminals from high-density cultures of STX1-null hippocampal neurons either not rescued or rescued with STX1AWT or STX1A∆Habc. (E–G) Quantification of active zone (AZ) length, number of synaptic vesicles (SVs) within 200 nm distance from AZ, and number of docked SVs. (H, I) SV distribution of STX1-null and STX1A∆Habc neurons compared to that of STX1AWT neurons. Data information: in (B, E–G), data points represent single observations, the bars represent the mean ± SEM. In (C, H, I), data points represent the mean ± SEM. Red and black annotations (stars and n.s.) on the graphs show the significance comparisons to STX1-null and to STX1AWT neurons, respectively (nonparametric Kruskal–Wallis test followed by Dunn’s post hoc test, *p≤0.05, **p≤0.01, ***p≤0.001, ****p≤0.0001). The numerical values are summarized in Figure 2—source data 1.

Figure 2—source data 1

Quantification of the neuronal density at different time intervals and quantification of ultrastructural synaptic properties in high density cultures of STX1-null, STX1AWT, and STX1AΔΗabc neurons.

https://cdn.elifesciences.org/articles/69498/elife-69498-fig2-data1-v4.xlsx

Furthermore, we analyzed vesicle docking by morphological assessment of synaptic ultrastructure to determine whether STX1A∆Habc expression could reverse the impairment in the vesicle docking observed in STX1-null neurons (Vardar et al., 2016). To circumvent the reduction in cell number and the synapse number thereof, we transduced the neurons at DIV 2–3 to postpone the cell death as previously shown (Vardar et al., 2016) and analyzed the synapses using high-pressure freezing fixation (DIV 14–16) combined with electron microscopy (HPF-EM; Figure 2D–I). Firstly, we observed no difference in the postsynaptic density (PSD) length, which is an indirect measurement of the opposing AZ length, among STX1-null, STX1AWT, and STX1A∆Habc neurons (Figure 2E). On the other hand, the total SV number within 200 nm from the AZ was significantly reduced in STX1A∆Habc neurons compared to that in STX1AWT neurons (Figure 2F). STX1A∆Habc also did not restore vesicle docking, which remained at ~50% of the STX1AWT neurons (Figure 2G). Similarly, the SV distribution within 100 nm of the AZ were comparable between STX1-null and STX1A∆Habc neurons, with both significantly altered number of SVs compared to the STX1WT neurons, especially in the 15, 40, and 100 nm range from AZ (Figure 2H and I). This suggests a general alteration of the synaptic organization even though the length of AZs was unaltered.

Based on the lack of immunofluorescent signal (Figure 1C) together with the lack of any rescue activity in any release parameters (Figures 1E–I2G and I) and neuronal survivability for STX1A∆Habc (Figure 2C), we again examined the expression level of STX1A∆Habc in comparison with STX1AWT, this time using constructs with a C-terminal FLAG tag (Figure 1—figure supplement 1). C-terminal FLAG tag did not reveal significant changes in the expression of STX1AWT (Figure 1—figure supplement 1). We then measured the immunofluorescent signal using a FLAG antibody in the neurons expressing FLAG-tagged STX1AWT, STX1A∆N2-9, STX1ALEOpen, or STX1A∆Habc, all of which showed similar levels of reduction in the expression as compared to the non-tagged constructs (Figure 1—figure supplement 1), suggesting that the lack of immunofluorescent signal in STX1A∆Habc (Figure 1C) is not due to a loss of antibody binding epitope, but rather due to the low level of protein.

Deletion of the entire N-terminal stretch does not impair neurotransmitter release

It is striking that deletion of the 2–9 amino acids (aa), namely the N-peptide, of STX1A revealed no significant phenotype in synaptic transmission from central synapses (Figure 1), even though this domain has been designated as a crucial factor for neurotransmitter release. Though the aa 2–9 has been defined as the residues binding to the outer surface of Munc18-1 (Hu et al., 2007; Burkhardt et al., 2008), the whole 2–28 aa stretch manifests an unstructured nature in NMR studies (Misura et al., 2000), suggesting a potential involvement in protein–protein interactions. Thus, we extended our analysis of the function of N-peptide by constructing STX1A with longer deletions in the N-terminus (STX1A∆N2-19 and STX1A∆N2-28) and probed the effects of these mutants on synaptic transmission.

Compared to the exogenous expression of STX1AWT, deletion of 19 or 28 aa from the N-terminus reduced the expression of STX1A to ~60% (Figure 3A–B), suggesting a modulatory effect of the unstructured N-terminal domain on STX1’s expression or stability. However, neither the reduction in STX1A expression nor loss of the putative Munc18-1 binding domain influenced the Munc18-1 levels, which was effectively rescued back to WT-like levels (Figure 3A and C).

Figure 3 with 2 supplements see all
Deletion of the entire N-terminal stretch does not impair neurotransmitter release.

(A) Example images of immunofluorescence labeling for Bassoon, STX1A, and Munc18-1 shown as red, green, and blue, respectively, in the corresponding composite pseudocolored images obtained from high-density cultures of STX1-null hippocampal neurons either not rescued or rescued with STX1AWT, STX1A∆2-9, STX1A∆2-19, or STX1A∆2-28. Scale bar: 10 µm. (B, C) Quantification of the immunofluorescence intensity of STX1A and Munc18-1 as normalized to the immunofluorescence intensity of Bassoon in the same ROIs as shown in (A). The values were then normalized to the values obtained from STX1AWT neurons. (D) Example traces (left) and quantification of the amplitude (right) of EPSCs obtained from hippocampal autaptic STX1-null neurons either not rescued or rescued with STX1AWT, STX1A∆2-9, STX1A∆2-19, or STX1A∆2-28. (E) Example traces (left) and quantification of the charge transfer (right) of sucrose-elicited readily releasable pools (RRPs) obtained from the same neurons as in (D). (F) Quantification of probability of vesicular release (Pvr) determined as the percentage of the RRP released upon one action potential (AP). (G) Example traces (left) and quantification (right) of paired-pulse ratio (PPR) measured at 40 Hz. The artifacts are blanked in the example traces. (H) Example traces (left) and quantification of the frequency (right) of mEPSCs. The example traces were filtered at 1 kHz. (I) Quantification of mEPSC rate as spontaneous release of one unit of RRP. Data information: the artifacts are blanked in example traces in (D) and (G). The example traces in (H) were filtered at 1 kHz. In (B–I), data points represent single observations, the bars represent the mean ± SEM. Red and black annotations (stars and n.s.) on the graphs show the significance comparisons to STX1-null and to STX1AWT neurons, respectively (nonparametric Kruskal–Wallis test followed by Dunn’s post hoc test, ****p≤0.0001). The numerical values are summarized in Figure 3—source data 1.

Figure 3—source data 1

Quantification of the lentiviral expression of STX1AWT and STX1AΔΝ mutants in STX1-null neurons and the consequent neurotransmitter release properties.

https://cdn.elifesciences.org/articles/69498/elife-69498-fig3-data1-v4.xlsx

Strikingly, similar to the deletion of the N-peptide, neither deletion of 2–19 aa nor 2–28 aa led to full inhibition of vesicle fusion nor of vesicle priming, but only a graded trend towards a decrease by 20–30% (Figure 3D and E). STX1AWT and STX1A∆N2-9 neurons had an average EPSC of ~6 nA and an average RRP of ~0.5 nC, while STX1A∆N2-19 and STX1A∆N2-28 had an average EPSC of ~4 nA and an average RRP of ~0.4 nC (Figure 3D and E). A trend towards a reduction in release was also expressed in Pvr, such that STX1A∆N2-19 and STX1A∆N2-28 neurons manifested Pvr of ~6%, whereas STX1AWT and STX1A∆N2-9 neurons released with a Pvr of ~8% and ~7%, respectively (Figure 3F). As another measure of Pvr, we induced paired action potentials (APs) at 40 Hz and observed no difference in paired-pulse ratio (PPR) of EPSCs between STX1AWT and STX1A∆N neurons (Figure 3G). Similarly, spontaneous release inclined to be impaired by 30–45% but not significantly, remaining at around 3–4 Hz compared to ~6 Hz of STX1AWT (Figure 3H). A similar level of reduction both in mEPSC frequency and RRP size recorded from STX1A∆N2-19 and STX1A∆N2-28 neurons led to no difference in spontaneous vesicle fusion rate compared to that recorded from STX1AWT neurons (Figure 3I).

Apart from STX1A’s first nine aa, the STX1-N-peptide–Munc18-1 interaction is also proposed to be regulated by the phosphorylation of STX1’s S14 residue by CKII (Rickman and Duncan, 2010). To test whether the phosphorylation of S14 affects Munc18-1 trafficking and neurotransmitter release, we generated phosphonull (S14A) and phosphomimetic (S14E) STX1A mutants. We again measured the STX1A and Munc18-1 levels at synapses, which revealed no impact of the phosphorylation status of S14 on either STX1A or Munc18-1 levels (Figure 3—figure supplement 1), consistent with the finding that S14A causes only a minor decrease in the affinity of STX1A to Munc18-1 (Burkhardt et al., 2008). As a direct function of STX1A S14 phosphorylation on vesicular release from neurons or neuroendocrine cells has been also proposed (Rickman and Duncan, 2010; Shi et al., 2020), we tested whether it would also influence the fusion of presynaptic vesicles. Both STX1AS14A and STX1AS14E efficiently restored all the release parameters to WT-like levels in STX1-null neurons (Figure 3—figure supplement 1), which suggests that the modulation of the STX1A N-peptide–Munc18-1 interaction by S14 phosphorylation does not alter its function in neurotransmitter release from central synapses. Neither N-peptide deletion nor phosphorylation modulation mutants compromised the neuronal survival (Figure 3—figure supplement 2).

‘Opening’ of STX1A in combination with the deletion of its entire N-terminal stretch does not impair neurotransmitter release

Munc18-1 binding to the N-peptide or to the closed conformation of STX1 constitutes the two well-defined interaction modes between these proteins, yet neither mutation causes a major deficit in synaptic release (Figures 1 and 3). However, Munc18-1 interacts with STX1AWT through multiple interaction points including the SNARE motif of STX1A (Misura et al., 2000; Burkhardt et al., 2008; Liang et al., 2013). To test whether or not the modulation of both ‘closed’ and ‘N-peptide’ binding modes would result in a drastic loss of the STX1A–Munc18-1 binary complex (Rickman et al., 2007) and thereby a loss of neurotransmitter release, we constructed STX1A mutants in which the N-peptide is deleted at differing lengths in conjunction with the LEOpen mutation. Firstly, we observed that N-peptide deletion in addition to the LEopen mutation decreased the STX1A and Munc18-1 levels further than that already caused by LEOpen mutation alone (Figure 4A–C).

Figure 4 with 3 supplements see all
‘Opening’ of STX1A in combination with the deletion of its entire N-terminal stretch does not impair neurotransmitter release.

(A) Example images of immunofluorescence labeling for Bassoon, STX1A, and Munc18-1 shown as red, green, and blue, respectively, in the corresponding composite pseudocolored images obtained from high-density cultures of STX1-null hippocampal neurons either not rescued or rescued with STX1AWT, STX1ALEOpen, STX1ALEOpen + ∆N2-9, STX1ALEOpen + ∆N2-19, or STX1ALEOpen + ∆N2-28. Scale bar: 10 µm (B, C) Quantification of the immunofluorescence intensity of STX1A and Munc18-1 as normalized to the immunofluorescence intensity of Bassoon in the same ROIs as shown in (A). The values were then normalized to the values obtained from STX1AWT neurons. (D) Example traces (left) and quantification of the amplitude (right) of EPSCs obtained from hippocampal autaptic STX1AWT, STX1ALEOpen, STX1ALEOpen + ∆N2-9, STX1ALEOpen + ∆N2-19, or STX1ALEOpen + ∆N2-28 neurons. (E) Example traces (left) and quantification of the charge transfer (right) of sucrose-elicited readily releasable pools (RRPs) obtained from the same neurons as in (D). (F) Quantification of probability of vesicular release (Pvr) determined as the percentage of the RRP released upon one action potential (AP). (G) Example traces (left) and quantification (right) of paired-pulse ratio (PPR) measured at 40 Hz. (H) Example traces (left) and quantification of the frequency (right) of mEPSCs. (I) Quantification of mEPSC rate as spontaneous release of one unit of RRP. (I) Quantification of mEPSC rate as spontaneous release of one unit of RRP.

Figure 4—source data 1

Quantification of lentiviral expression of STX1ALEOpen and STX1ALEOpen + ΔΝ mutants in STX1-null neurons and the consequent neurotransmitter release properties.

https://cdn.elifesciences.org/articles/69498/elife-69498-fig4-data1-v4.xlsx

Despite the presumed loss of the two STX1A–Munc18-1 interaction modes, it is remarkable that all LEOpen-∆N combination mutants rescued Ca2+-evoked neurotransmitter release to almost STX1AWT levels with only a trend towards a reduction by 25–35% (Figure 4D). Because STX1ALEOpen neurons showed increased EPSCs – albeit not significant – with an average of ~7 nA compared to ~4 nA of STX1AWT, EPSCs recorded from STX1ALEOpen+∆N neurons were significantly smaller than that of STX1ALEOpen neurons and remained at ~3 nA (Figure 4D). This suggests that the small enhancement of Ca2+-evoked release by the presumed open conformation by LEOpen mutation was reversed by additional N-peptide deletions (Figure 4D). On the other hand, the reduction in RRP observed in neurons that express LEOpen mutation was not reverted back to WT-like levels by the addition of N-peptide deletions, but instead was further exaggerated as the RRP size significantly decreased in STX1ALEOpen+∆N neurons compared to that in STX1AWT neurons (Figure 4E). As a result, increased Pvr, which is the hallmark phenotype of the LEOpen mutation (Gerber et al., 2008), was reversed back to WT-like levels with only a trend toward a small increase (Figure 4F). Increased Pvr in STX1ALEOpen-expressing neurons led to decreased PPR when measured at 40 Hz, and N-peptide deletions in STX1ALEOpen reverted PPR back to levels comparable to neurons expressing STX1AWT (Figure 4G). Similarly, mEPSC frequency and mEPSC release rate obtained from the STX1ALEOpen+∆N mutants were significantly smaller than that of STX1ALEOpen mutant (Figure 4H and I).

Surprisingly, putative disruption of the two supposedly main interaction points between STX1A and Munc18-1 – by deleting N-peptide in its entirety in LEOpen STX1A – ultimately led to neuronal death (Figure 4—figure supplement 1) indicative of independence of STX1’s functions in neurotransmitter release and neuronal maintenance of one another. However, the onset of cell death was postponed by the expression of STX1ALEOpen+∆N mutants compared to that observed in STX1-null neurons as at DIV15 almost all neurons expressing the STX1ALEOpen+∆N mutants were still alive (Figure 4—figure supplement 1). Because the electrophysiological recordings are mostly conducted at DIV 13–20, the compromised cell viability is unlikely to account for the reduction in neurotransmission in STX1ALEOpen+∆N mutants compared to that of in STX1ALEOpen mutant.

A severe reduction in STX1 expression induced by in vitro knock-down (Arancillo et al., 2013; Zhou et al., 2013) or transgenic knock-in (Arancillo et al., 2013) strategies results in a strong impairment in neurotransmitter release. Based upon that, we argued that the reduction of release parameters (Figure 4D–I) of STX1ALEOpen by additional N-peptide deletions may be due to decreased expression of STX1A (Figure 4B). To test this hypothesis, we down-titrated the viral load from 1× (~400 × 103 viral particles per 35 mm well) to 1/12× for STX1AWT and to 1/3× and 1/6× for STX1ALEOpen to reach expression of STX1A at a level comparable to that in STX1ALEOpen+∆N2-28 neurons (Figure 4—figure supplement 2). As our viral constructs include NLS-GFP before the P2A sequence followed by STX1A, nuclear GFP showed a decrease when the amount of virus was reduced (Figure 4—figure supplement 2). Immunofluorescent labeling in autaptic neurons revealed that reducing the viral amount was effective in reducing expression levels of either STX1AWT or STX1ALEOpen to the levels comparable to that of STX1ALEOpen+∆N2-28. However, reducing the exogenous expression level of STX1AWT or STX1ALEOpen down to the level of STX1ALEOpen+∆N2-28 did not cause a difference in their neurotransmitter release properties (Figure 4—figure supplement 2).

We have previously reported that STX1 level becomes a rate-liming factor in neurotransmission when endogenous STX1B expression is knocked-down by shRNA to a level below 20% on an STX1A-null background (Arancillo et al., 2013). Because reducing the lentiviral exogenous expression of STX1A down to ~20% of the initial experimental conditions did not show any alterations in synaptic release properties (Figure 4—figure supplement 2) and thus not reconcile with our previous hypothesis (Arancillo et al., 2013), we compared the endogenous STX1A expression in STX1A+/+; STX1B+/+ neurons to the exogenous expression level in STX1-null neurons transduced with 1× STX1A (Figure 4—figure supplement 3). We found that transduction of STX1-null neurons with STX1A using 1× viral volume leads to approximately threefold higher STX1A exogenous expression compared to that of WT neurons. Thus, using 1/12th of the initial viral volume for STX1A transduction leads to an expression level of ~60% compared to the endogenous level, suggesting that indeed one copy of either STX1A or STX1B is enough to drive normal synaptic transmission while being insufficient to rescue Munc18-1 levels back to the WT-like levels (Figure 4—figure supplement 3). However, please note that our model system does not include STX1B expression and therefore exogenous expression of STX1A in STX1-null neurons by 1× viral volume does not fall within overexpression studies. In summary, our STX1A down-titration experiments show that the reduction in neurotransmitter release properties observed in STX1ALEOpen+∆N2-28 neurons compared to that of STX1ALEOpen neurons does not stem from lower copy number of STX1A but rather from a functional deficit (Figure 4—figure supplement 2).

It is known that Munc18-1 also functions upstream of the vesicle docking step (Toonen et al., 2006; Gulyas-Kovacs et al., 2007). Therefore, we analyzed the state of docked vesicles in neurons that express either STX1A∆N2-28, STX1ALEOpen, or STX1ALEOpen+∆N2-28 using HPF-EM (Figure 5A). PSD length, and thus AZ length, was again comparable between all the mutants and STX1AWT (Figure 5B), whereas STX1ALEOpen+∆N2-28 neurons showed a small but significant reduction in total SV number within 200 nm of AZ (Figure 5C). Strikingly, the neurons in which two Munc18-1 binding modes were modulated by the STX1ALEOpen+∆N2-28 mutation showed docked vesicles were reduced to ~50% of those in STX1AWT synapses (Figure 5D). On the other hand, neither LEOpen mutation nor N-peptide deletion alone did not influence vesicle docking (Figure 5D). Furthermore, vesicle distribution analysis revealed an accumulation of vesicles at 5 nm distance from AZ in STX1A∆N2-28 neurons but a reduction in STX1ALEOpen+∆N2-28 neurons (Figure 5F and G), whereas STX1ALEOpen neurons did not show a major alteration in their vesicle distribution within 100 nm from AZ (Figure 5H).

Figure 5 with 1 supplement see all
‘Opening’ of STX1A in combination with the deletion of its entire N-terminal stretch reduces the number of docked synaptic vesicles (Svs).

(A) Example high-pressure freezing fixation combined with electron microscopy (HPF-EM) images of nerve terminals from high-density cultures of STX1AWT, STX1A∆N2-28, STX1ALEOpen, and STX1ALEOpen + ∆N2-28 Neurons. (B–D) Quantification of active zone (AZ) length, number of SVs within 200 nm distance from AZ, and number of docked SVs. (E) Correlation of the number of docked SVs obtained by HPF-EM to the size of readily releasable pool (RRP) obtained by electrophysiological recordings. (F–H) SV distribution of STX1A∆N2-28, STX1ALEOpen, and STX1ALEOpen + ∆N2-28 neurons compared to that of STX1AWT neurons. Data information: in (B–D), data points represent single observations, the bars represent the mean ± SEM. In (E–H), data points represent mean ± SEM. Black annotations on the graphs show the significance comparisons to STX1AWT rescue (nonparametric Kruskal–Wallis test followed by Dunn’s post hoc test in B–D, multiple t-tests in F–H *p≤0.05, **p≤0.01, ***p≤0.001). The numerical values are summarized in Figure 5—source data 1.

Figure 5—source data 1

Quantification of the ultrastructural synaptic properties of STX1AWT, STX1AΔΝ2-28, STX1ALEOpen, and STX1ALEOpen + ΔΝ2-28 neurons.

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It is possible that a reduction of RRP might reflect a reduction in synapse number if synapse loss precedes neuronal loss in the case of STX1ALEOpen and STX1ALEOpen+∆N2-28 neurons. In that scenario, number of docked vesicles, which is morphologically assessed by evaluating the existing synapses in mass culture, would not be affected and thus lead to differential outcomes for vesicle docking and vesicle fusion in neurons destined to death. To test this, we analyzed the synapse number in autaptic neurons for which we used the images shown in Figure 4—figure supplement 2 and found no difference neither in the synapse number nor in the synapse area among STX1AWT, STX1ALEOpen, and STX1ALEOpen+∆N2-28 neurons as determined by VGlut1-positive puncta (Figure 5—figure supplement 1). Previously, we have shown that vesicle priming can be completely abolished by a STX1A mutant (A240V, V244A) with the vesicle docking remaining intact (Vardar et al., 2016). We also have reported that the vesicle priming is more prone to impairments by mutations in the vesicle release machinery than is vesicle docking, which suggests a separation or a different cooperativity between these events (Zarebidaki et al., 2020). In this light, we plotted the number of docked SVs versus the RRP size and observed that the RRP is also more susceptible to a reduction than is vesicle docking for the STX1A–Munc18-1 binding mutants (Figure 5E).

STX1’s N-peptide has a modulatory function in short-term plasticity and Ca2+-sensitivity of synaptic transmission

So far, our analysis has shown that STX1’s N-peptide is not indispensable for neurotransmitter release (Figures 1 and 3), but plays a modulatory role in protein expression (Figure 3) and, when STX1’s open conformation is facilitated by LEOpen mutation, in vesicle fusion and Pvr (Figure 4). To elucidate the modulation of neurotransmitter release by STX1’s N-peptide, we took a closer look at Pvr and its effect on STP (Figure 6A). Even though the neurons expressing any STX1A∆N mutants showed only a trend towards decreased Pvr compared to that of STX1AWT neurons (Figure 3), their STP behavior in response to 50 stimuli at 10 Hz differed greatly (Figure 6A). Both STX1A∆N2-19 and STX1A∆N2-28 showed first zero then only ~10% depression following the first stimulus and STX1A∆N2-9 exhibited less depression than STX1AWT after the first 10 stimuli as analyzed by normalizing the EPSC responses to the first response (Figure 6A). Because STX1A∆N2-19 and STX1A∆N2-28 neurons have a reduced initial EPSC compared to that of STX1AWT neurons, we also plotted the absolute values of EPSCs elicited at 10 Hz (Figure 6—figure supplement 1). STX1A∆N2-28 tended to remain to elicit smaller EPSCs throughout the high-frequency stimuli (HFS) compared to those of STX1AWT (Figure 6—figure supplement 1). Zoomed-in example traces for the representation of the first and last five stimuli can be found in Figure 6—figure supplement 2.

Figure 6 with 2 supplements see all
STX1A’s N-peptide has a modulatory function in short-term plasticity and Ca2+-sensitivity of synaptic transmission.

(A) Example traces (left) and quantification (right) of STP measured by 50 stimulations at 10 Hz from STX1AWT, STX1A∆N2-9, STX1A∆N2-19, or STX1A∆N2-28 neurons. The traces show the absolute values, whereas the quantification shows normalized EPSC to EPSC1. (B) Example traces (left) and quantification (right) of the recovery of readily releasable pool (RRP) determined as the fraction of RRP measured at a second pulse of 500 mM sucrose solution after 2 s of initial depletion from STX1AWT, STX1A∆N2-9, STX1A∆N2-19, or STX1A∆N2-28 neurons. (C) Example traces (left) and quantification (right) of the ratio of the charge transfer triggered by 250 mM sucrose over that of 500 mM sucrose as a read-out of fusogenicity of the synaptic vesicles (SVs). (D) Example traces (left) and quantification (right) of Ca2+-sensitivity as measured by the ratio of EPSC amplitudes at [Ca2+]e of 0.5, 1, 2, 4, and 10 mM recorded from STX1AWT, STX1A∆N2-9, or STX1A∆N2-28 neurons. The responses were normalized to the response at [Ca2+]e of 10 mM. (E) Paired-pulse ratio (PPR) of EPSC amplitudes at [Ca2+]e of 0.5, 1, 2, 4, and 10 mM recorded at 40 Hz. (F) Example traces (left) and quantification (right) of STP measured by 50 stimulations at 10 Hz from STX1AWT, STX1ALEOpen, STX1ALEOpen + ∆N2-9, STX1ALEOpen + ∆N2-19, or STX1ALEOpen + ∆N2-28 neurons. The traces show the absolute values, whereas the quantification shows normalized EPSC to EPSC1. (G) Example traces (left) and quantification (right) of the recovery of RRP determined as the fraction of RRP measured at a second pulse of 500 mM sucrose solution after 2 s of initial depletion from STX1AWT, STX1ALEOpen, STX1ALEOpen + ∆N2-9, STX1ALEOpen + ∆N2-19, or STX1ALEOpen + ∆N2-28 neurons. (H) Example traces (left) and quantification (right) of the ratio of the charge transfer triggered by 250 mM sucrose over that of 500 mM sucrose as a read-out of fusogenicity of the SVs. (I) Example traces (left) and quantification (right) of Ca2+-sensitivity recorded from STX1AWT, STX1ALEOpen, STX1ALEOpen + ∆N2-9, or STX1ALEOpen + ∆N2-28 neurons. The responses were normalized to the response at [Ca2+]e of 10 mM. (J) PPR of EPSC amplitudes at [Ca2+]e of 0.5, 1, 2, 4, and 10 mM recorded at 40 Hz from STX1AWT, STX1ALEOpen, or STX1ALEOpen + ∆N2-28 neurons. Data information: the artifacts are blanked in example traces in (A, D, F, I). In (A, D, E, F, I, J), data points represent the mean ± SEM. In (B, C, G, H), data points represent single observations, the bars represent the mean ± SEM. Black and red annotations on the graphs show the significance comparisons to STX1AWT or STX1ALEOpen, respectively. (either nonparametric Kruskal–Wallis followed by Dunn’s post hoc test or one-way ANOVA followed by Holm–Sidak’s post hoc test was applied based on the normality of the data, *p≤0.05, **p≤0.01, ***p≤0.001, ****p≤0.0001). The numerical values are summarized in Figure 6—source data 1.

Figure 6—source data 1

Quantification of the STP, recovery of RRP, RRP fraction released by 250 mM sucrose solution application and Ca2+-sensitivity of the vesicles in neurons expressing STX1AWT, STX1AΔΝ- or STX1ALEOpen mutants.

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Whereas Pvr shapes the STP curve starting from the initial phase, the late phase of STP is affected not only by Pvr, but also by the rate of SVs newly arriving at the AZ. Because all the STX1A∆N neurons showed an altered behavior in the late phase of STP, we hypothesized that these neurons might keep up better with the high-frequency stimulus than STX1AWT does because of an increase in newly arrived SVs or replenishment of the RRP. To study the efficacy of replenishment of the primed vesicles, we stimulated the neurons with a double pulse of 500 mM sucrose solution with a time interval of 2 s (Figure 6B) as the replenishment of the whole pool of the primed vesicles after sucrose depletion takes at least 10 s (Stevens and Tsujimoto, 1995). STX1A∆N2-9 showed no effect on the fraction of the RRP recovered after full depletion when compared to that of STX1AWT (Figure 6B). On the other hand, STX1A∆N2-19 and STX1A∆N2-19 slightly decreased the vesicle replenishment rate by ~10%, which is contrary to our initial expectation (Figure 6B). This suggests that an increased replenishment rate does not account for the decreased depression in the STP curves of STX1A∆N mutants.

Pvr and the degree of STP depend on both vesicle fusogenicity and Ca2+-sensitivity of the vesicular release, thus we scrutinized the effects of N-peptide deletions on these variables. Calculating the RRP fraction released in response to a sub-saturating, 250 mM sucrose solution application revealed no difference between STX1AWT and STX1A∆N neurons, suggesting no decrease in SV fusogenicity with these mutants (Figure 6C). Interestingly, when we generated Ca2+-dose–response curves by evoking AP driven EPSCs in the presence of 0.5, 1, 2, 4, or 10 mM Ca2+-containing extracellular solutions, we observed a slightly lowered apparent Ca2+-sensitivity in the STX1A∆N2-19 (Figure 6—figure supplement 1) and STX1A∆N2-28 neurons (Figure 6D) when compared to STX1AWT neurons. On the other hand, STX1A∆N2-9 neurons showed a normal pattern of increase in EPSCs in relation to increasing extracellular Ca2+-concentration (Figure 6D). We also measured the PPR, which is inversely related to Pvr, at different extracellular Ca2+-concentrations and determined that STX1A∆N2-28 had a significantly higher PPR at 0.5 mM [Ca2+]e compared to that of STX1AWT (Figure 6E). This was also evident in STP behavior elicited by 5 AP stimulation at 40 Hz as STX1A∆N2-28 neurons showed a greater facilitation at 0.5 mM [Ca2+]e compared to that of STX1AWT (Figure 6—figure supplement 1). Whereas increasing [Ca2+]e to 2 mM was not sufficient to drive the STP behavior of STX1A∆N2-28 neurons towards STX1AWT-like pattern, at the highest [Ca2+]e tested all the groups STX1AWT, STX1A∆N2-9, and STX1A∆N2-28 showed a similar level of depression upon 40 Hz stimulation (Figure 6—figure supplement 1).

It is well documented that the presumed facilitation of opening of STX1 and thus the increase in Pvr by LEOpen mutation enhances short-term depression (Acuna et al., 2014; Gerber et al., 2008). Deletion of N-peptide at any length in STX1ALEOpen did not change the degree of the depression in the late phase of the high-frequency stimulus; however, all the deletions decreased the slope of the depression in the initial phase compared to that of STX1ALEOpen alone (Figure 6F). Whereas the EPSCs recorded from STX1ALEOpen neurons by 10 Hz stimulation tended to be initially larger, they declined further compared to that of STX1AWT neurons (Figure 6—figure supplements 1 and 2). On the other hand, STX1ALEOpen+∆N2-28 mutants remained to produce smaller EPSCs throughout the HFS compared to those of both STX1AWT and STX1ALEOpen (Figure 6—figure supplements 1 and 2). The recovery of the RRP after sucrose depletion was enhanced by the open conformation of STX1A but reverted back to the WT-like levels by the expression of STX1ALEOpen+∆N mutants (Figure 6G). Strikingly, the increase in fusogenicity was not influenced by the N-peptide deletions (Figure 6H), which is consistent with the observation that those mutants also did not change LEOpen mutation-driven enhancement of short-term depression at the late phase of the HFS (Figure 6F). On the other hand, N-peptide deletions imposed a right-shift in the Ca2+-dose–response curves on the STX1ALEOpen, which markedly increased the Ca2+-sensitivity, making them approach again WT-like levels (Figure 6I, Figure 6—figure supplement 1). Consistent with increased fusogenicity and Ca2+-sensitivity, STX1ALEOpen neurons always showed a greatly reduced PPR when compared to STX1AWT neurons in all extracellular Ca2+-concentrations tested (Figure 6J). However, at low extracellular Ca2+-concentrations STX1ALEOpen+∆N2-28 neurons exhibited a PPR comparable to that of STX1AWT neurons, but at high Ca2+-concentrations it was comparable to that of STX1ALEOpen neurons (Figure 6J). Contrary to STX1AWT neurons, STX1ALEOpen neurons showed no facilitation at 0.5 mM [Ca2+]e and a greater depression at 2 mM [Ca2+]e as well as at 10 mM [Ca2+]e. Whereas STX1ALEOpen+∆N2-28 showed a similar pattern of facilitation at 0.5 mM [Ca2+]e to STX1AWT, at higher [Ca2+]e their short-term depression approached the level of STX1ALEOpen (Figure 6—figure supplement 1). This and the observation that N-peptide deletion leads to an altered behavior only in the initial phase of the 10 Hz stimuli – when STX1A’s open conformation is facilitated – is consistent with the reduced Ca2+-sensitivity (Figure 6I) but unaltered fusogenicity (Figure 6F) of the vesicles.

Decreased Ca2+-sensitivity can arise from either reduced Ca2+-influx as a result of alterations in Ca2+-channel localization or gating, or from a disturbance in Ca2+-secretion coupling. To address this issue, we expressed the Ca2+-reporter GCamp6f coupled to Synaptophysin (SynGCamp6f) in STX1-null neurons with or without STX1A rescue constructs and measured the immunofluorescence at the synapses at baseline or upon 1, 2, 5, 10, or 20 AP stimulation at 10 Hz (Figure 7A and B). Surprisingly, STX1-null neurons showed a decreased global Ca2+-influx compared to neurons rescued with STX1AWT (Figure 7C). However, STX1A∆N2-9 or STX1A∆N2-28 did not influence the SynGCamp6f signal at any AP number elicited (Figure 7D), whereas global Ca2+-influx was reduced in synapses in STX1ALEOpen and STX1ALEOPen+∆N2-28 neurons (Figure 7E).

Figure 7 with 1 supplement see all
Ca2+-influx is reduced in STX1-null and in STX1LEOpen neurons.

(A, B) Example images and average of SynGCaMP6f fluorescence as (ΔF/F0) in STX1-null neurons either not rescued or rescued with STX1AWT, STX1A∆N2-9, STX1A∆N2-19, STX1ALEOpen, or STX1ALEOpen + ∆N2-28. The images were recorded at baseline, and at 1, 2, 5, 10, and 20 action potentials (APs). Scale bar: 10 µm (C–E) Maximum fluorescence changes (ΔF/F0) in STX1-null, STX1A∆N2-9, STX1A∆N2-28, STX1ALEOpen, or STX1ALEOpen + ∆N2-28 in comparison to that in STX1AWT neurons recorded at 1, 2, 5, 10, and 20 APs. (F–I-J) Summary plots of STX1A expression level, neuronal viability, readily releasable pool (RRP) charge, number of docked synaptic vesicles (SVs), and maximum SynGCaMP6f ΔF/F0 at 20 AP from STX1-null, STX1AWT, TX1A∆N2-28, STX1ALEOpen, and STX1ALEOpen + ∆N2-28. All the values were normalized to the one obtained from STX1AWT neurons in each individual culture. Data information: data points in all graphs represent the mean ± SEM. Black annotations on the graphs show the significance comparisons to STX1AWT (either unpaired t-test or Mann–Whitney test was applied in C based on the normality of the data; in D and E, nonparametric Kruskal–Wallis test followed by Dunn’s post hoc test was applied, *p≤0.05, ****p≤0.0001). The numerical values are summarized in Figure 7—source data 1.

Figure 7—source data 1

Quantification of the increase in SynGCaMP6f signal recorded at baseline or different numbers of APs in neurons expressing STX1AWT, STX1AΔΝ- or STX1ALEOpen mutants.

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The reduction in Ca2+-influx at the presynaptic terminals in STX1-null, STX1ALEOpen and STX1ALEOPen+∆N2-28 neurons compared to that of STX1AWT neurons is indicative of involvement of STX1 in the vesicular release processes upstream of vesicle docking (Figure 7A–E). As these STX1A mutants also showed severely decreased expression levels (Figure 4), we hypothesized that the synaptic structural properties might be affected by the expression level of STX1A. To test this, we measured the global Ca2+-influx in neurons expressing low level of STX1AWT by using again 1/12th of the initial viral load and observed no effect of reduced expression level of STX1A on SynGCamp6f signal (Figure 7—figure supplement 1), which indicates a functional account of open conformation of STX1A for global Ca2+-influx reduction rather than an expressional account (Figure 7E). As a summary, we plotted expression level of STX1A, neuronal viability, size of the RRP, number of docked SV, or the level of Ca2+-influx at 20 AP in relation to N-peptide deletion and/or open conformation of STX1A (Figure 7F–J). Almost all the parameters showed a decreased degree of rescue by the expression of STX1A with combined mutation of LEOpen and N-peptide deletion (Figure 7F–J), suggesting a cooperative function of STX1A’s closed conformation and N-peptide.

Discussion

The tight interaction between STX1 and Munc18-1 is not dictated through a single contact point but rather spans a large area both on STX1 and Munc18-1 (Misura et al., 2000), to which STX1’s N-peptide and closed conformation largely contribute. Using our STX1-null mouse model system, we can draw several conclusions from mutant STX1 rescue experiments: (1) STX1’s Habc-domain is essential for the stability of STX1 and Munc18-1, and thus for neurotransmitter release and overall STX1 function; (2) STX1’s N-peptide is dispensable for neurotransmitter release, but has a modulatory function for STX1’s stability, for Ca2+-sensitivity of vesicular release, and importantly for STP; and (3) neurotransmitter release can proceed even when the two interaction modes are presumably intervened by N-peptide deletions in conjunction with LEOpen mutation in STX1A (Figure 8).

Speculative model of effects of N-peptide deletions and LEOpen mutation on vesicular release.

(A) Native state of STX1A. (B) N-peptide deletion of STX1A leads to a decrease in Ca2+-sensitivity of vesicular release and short-term depression (STD) upon 10 Hz stimulation potentially through increased distance of Ca2+-channel synaptic vesicle (SV) coupling. (C) LEOpen mutation on STX1A increases fusogenicity and Ca2+-sensitivity of SVs and thus leads to a high degree of STD. It also leads to reduced global Ca2+-influx. (D) SV fusion proceeds normal when LEOpen mutation is combined with N-peptide deletion. LEOpen mutation dictates SV fusogenicity and Ca2+-influx by increasing the former and decreasing the latter.

STX1’s Habc-domain is essential for the overall function of STX1

The three helical Habc-domain constitutes a major portion of STX1. As the main driving force for vesicle fusion is the zippering of the SNARE domains of STX1, SNAP25, and Syb2 (Rizo and Sudhof, 2012; Rizo and Xu, 2015; Baker and Hughson, 2016), STX1 that lacks only the Habc-domain sufficiently mediates liposome fusion in reconstitution experiments (Rathore et al., 2010; Shen et al., 2010). Even in the synaptic environment, a crucial function of the Habc-domain has been suggested only for spontaneous neurotransmitter release (Zhou et al., 2013; Meijer et al., 2012). However, the picture in intact synapses is more complex because neurotransmitter release proceeds as a result of multiple steps dependent on protein folding and trafficking, inter- and intramolecular interactions, relative conformations, and the proper localization of multiple synaptic proteins. Due to the lack of significant expression of STX1A∆Habc (Figure 1), we cannot draw a certain conclusion on whether or not Habc-domain of STX1 is directly involved in neurotransmitter release. However, it has to be noted that even though Munc18-1 with a E259K point mutation, which interferes with its interaction with STX1’s Habc-domain as well as with the SNARE complex, could mediate Ca2+-triggered neurotransmitter release in Munc18-1 knock-out neurons to some extent, more than half of the neurons were reported as synaptically silent (Meijer et al., 2012). Additionally, while liposome fusion can proceed without STX1’s Habc-domain (Rathore et al., 2010), the function of Munc18-1 as a template for SNARE complex formation (Ma et al., 2013; Ma et al., 2015) is greatly perturbed by the absence of STX1A’s Habc-domain (Jiao et al., 2018).

Overall, what is clear from our study and the previous studies is the importance of Habc-domain in proper folding of STX1 and its co-recruitment to the AZ with Munc18-1. Severely decreased expression levels of STX1, Munc18-1, or both occur when the interaction between the Habc-domain of STX1 and Munc18-1 is interrupted (Gulyas-Kovacs et al., 2007; Meijer et al., 2012; Vardar et al., 2020; Zhou et al., 2013). Even improper folding of the Habc-domain by an insertion/deletion (InDel) mutation, identified in relation to epilepsy, leads to a high degree of STX1 instability (Vardar et al., 2020). Consistently, both the STX1B InDel mutant (Vardar et al., 2020) and STX1A∆Habc mutant (Figure 2) were incapable of sustaining neuronal viability. Thus, we argue that the major role of Habc-domain of STX1 is to drive it into its correct folding and to recruit it together with Munc18-1 to the AZ.

STX1’s N-peptide role in neurotransmitter release is only detectable in STX1’s LEOpen configuration

On the contrary to the general view, we show here that the STX1’s N-peptide is not indispensable for neurotransmitter release, but rather only modulates STX1’s expression and the Ca2+-sensitivity of SVs. Above all, the dispensability of STX1’s N-peptide in vesicle fusion and particularly for proper recruitment of Munc18-1 to the AZ is consistent with the estimated contribution of N-peptide to the overall affinity of STX1 to Munc18-1, which is only minor (Burkhardt et al., 2008; Christie et al., 2012; Colbert et al., 2013).

Remarkably, the putative loss of two canonical interaction modes between STX1–Munc18-1 has little or no effect on synaptic transmission in general. This is not unprecedented as STX1A∆N and STX1ALEOpen mutants exhibit a largely unaltered binding affinity to Munc18-1 (Burkhardt et al., 2008). Additionally, these mutants have also been proposed to maintain the closed conformation when bound to Munc18-1 (Colbert et al., 2013; Dawidowski and Cafiso, 2013; Lai et al., 2017; Wang et al., 2017) potentially through additional contact points on STX1A including its SNARE motif (Misura et al., 2000; Burkhardt et al., 2008; Liang et al., 2013). Given that and the flexibility of STX1–Munc18-1 interaction, which induces large conformational changes on these proteins not only when STX1 is isolated but also when it enters the SNARE complex (Jakhanwal et al., 2017), it is conceivable that even for the STX1ALEOpen+∆N mutants a level of interaction between STX1–Munc18-1 must be retained.

Nevertheless, our analysis shows that even though both N-peptide deletion and LEOpen mutation produce the same degree of reduction in the binding affinity of STX1 to Munc18-1 (Burkhardt et al., 2008; Christie et al., 2012; Colbert et al., 2013), it is the native conformation that commands the Munc18-1 recruitment and/or stability at the synapse (Figures 1, 3 and 4). This led us to interpret Munc18-1’s binding to STX1 and its ultimate effect on SNARE complex formation as a two-step process, which is a convolution of the affinity and the efficacy of this interaction. Consistently, it has been thought that LEOpen mutation exposes the SNARE domain of STX1 (Dulubova et al., 1999), whereas absence of N-peptide tightens its Munc18-1-driven closed conformation (Khvotchev et al., 2007; Christie et al., 2012; Colbert et al., 2013) potentially resulting in opposing effects in SNARE complex formation. Indeed, our observation that N-peptide deletion reverses the STX1ALEOpen-dependent facilitation of neurotransmitter release parameters, which is generally attributed to its promotion of SNARE complex formation (Dulubova et al., 1999; Gerber et al., 2008, Acuna et al., 2014), hints at a reduction in the number of SNARE complexes formed by STX1ALEOpen. However, N-peptide likely plays only a minor role in determining the equilibrium of open-closed conformations of STX1WT in a membranous environment when STX1’s TMR is present (Dawidowski and Cafiso, 2013), and thus the modulation of neurotransmitter release by the N-peptide cannot be observed in STX1WT but can be only unmasked by the introduction of LEOpen mutation.

Importantly, Munc18-1 does not bind only to STX1, but it is also thought to bind to the SNARE complex formed by STX1, SNAP-25, and Syb-2 (Zilly et al., 2006; Dulubova et al., 2007; Shen et al., 2007; Burkhardt et al., 2008) to provide a template as a scaffold together with Munc13 (Ma et al., 2013; Ma et al., 2015; Lai et al., 2017; Wang et al., 2017). Given that the SNARE complex formation is a dynamic process, which involves not only the assistance but also the protection by Munc18-1 and Munc13 against NSF-dependent dissociation (Ma et al., 2013; He et al., 2017), such a two-step process is also applicable for the back- and forward shift in the number of SNARE complexes. Furthermore, one of Munc13’s primary functions is priming the vesicles (Varoqueaux et al., 2002) in addition to templating proper SNARE complex formation for which it assists opening of Munc18-1-bound STX1 (Ma et al., 2011; Ma et al., 2013; Lai et al., 2017; Wang et al., 2017). In this context, it has been shown that STX1ALEOpen recovers neurotransmitter release in Munc13-1/2-deficient mouse or worm neurons albeit only minimally (Lai et al., 2017; Tien et al., 2020), while it impairs Munc13’s proper assistance of parallel SNARE complex formation (Wang et al., 2017) and further reduces the locomotor activity and neurotransmitter release in Munc18-1-deficient worms (Tien et al., 2020). Thus, it is plausible that the stability of the SNARE complex is ensured by Munc18-1’s and Munc13’s efficient binding to STX1, which may account for the additive effects of N-peptide deletion and LEOpen conformation on the size of RRP (Figures 4 and 7). However, this regulation process must be upstream of the vesicle priming as the fusogenicity of the primed vesicles was predominantly dictated by the LEOpen mutation.

STX1A’s N-peptide regulates Ca2+-sensitivity of SVs and short-term plasticity, whereas LEOpen mutation dictates the SV fusogenicity

How the SVs become more fusogenic in the presence of STX1ALEOpen is not known, though one simple explanation is its propensity to produce reactive SNARE complexes with a higher number and efficacy (Dulubova et al., 1999; Acuna et al., 2014; Gerber et al., 2008). This hypothesis is appealing as it can also account for the faster recovery of the SVs to the RRP by LEOpen mutation (Figure 6). Surprisingly, however, addition of N-peptide deletions not only in STX1ALEOpen but also in STX1AWT exclusively slowed down the RRP replenishment without an effect on the SV fusogenicity (Figure 6), uncoupling the regulation of these two processes. We cannot explain this phenomenon based on our data but would like to draw attention to that there are still unsolved questions regarding the regulation of SV fusogenicity. In fact, it is thought that at the state of primed and even docked vesicles the SNAREs are zippered up to hydrophobic layer +4 (Sorensen et al., 2006; Walter et al., 2010; Vardar et al., 2016) and thus include already ‘opened’ STX1. Therefore, the increase in SV fusogenicity by STX1ALEOpen- and STX1ALEOpen+∆N-mutants might involve yet an unknown mechanism, which does not employ STX1’s N-peptide.

It is remarkable that the deletion of the N-peptide of 19 or 28 aa reduced the Ca2+-sensitivity of the vesicular release both in default and LEOpen STX1A (Figure 6). Ca2+-sensitivity of the vesicular release is estimated by the assessment of Ca2+-dose–response, which is a convoluted measurement of SV fusogenicity and Ca2+-channel–SV distance coupling. Accordingly, the increased Ca2+-sensitivity of vesicular release by LEOpen mutation stems partly from the increased fusogenicity of SVs (Figure 6). However, the rightward shift in Ca2+-dose–response curve caused by N-peptide deletions was not accompanied by an altered fusogenicity of SVs neither on default nor on LEOpen background of STX1A. Thus, it is conceivable that the N-peptide deletions might have led to disrupted Ca2+-channel–SV coupling, without effecting the fusogenicity of the vesicles. Consistently, N-peptide deletions in the LEOpen conformation, which do not alter LEOpen-dependent increase in vesicle fusogenicity, lead to an increase in PPR compared to that of LEOpen mutation alone only at low external Ca2+-concentrations. This suggests that when the vesicles are positioned at a greater distance to Ca2+-channels in the absence of N-peptide, the enhancement of fusogenicity governed by the opening of STX1A by LEOpen mutation remains insufficient to increase the Pvr at low Ca2+-concentrations. At higher Ca2+-concentration, on the other hand, the wider distance of the vesicles to Ca2+-channels becomes negligible and the LEOpen mutation-dependent enhancement of vesicle fusogenicity dominates the Pvr and thus reduces PPR in the case of STX1ALEOpen+ΔΝ mutants (Figure 6—figure supplement 1) as speculatively illustrated in Figure 8. This can also explain why the N-peptide deletions on STX1ALEOpen background slow down depression during the initial phase of the 10 Hz stimuli, while they affect the STP only at the late phase on STX1AWT background (Figure 6).

Interestingly, our data reveal a new function of STX1 in synaptic transmission in that it controls global Ca2+-entry into the presynapse as STX1 deficiency and LEOpen mutation led to a decreased Ca2+-influx (Figure 7). This can be explained either by alterations in Ca2+-channel gating and/or abundance. In fact, a direct interaction between STX1 and Ca2+-channels (Bachnoff et al., 2013; Cohen et al., 2007; Sheng et al., 1994; Wiser et al., 1996; Sajman et al., 2017) has been proposed to contribute to the overall Ca2+-sensitivity of the vesicular release machinery, where STX1 deemed an inhibitory role in baseline activity of Ca2+-channels (Trus et al., 2001). However, we observe a general decrease in Ca2+-entry also for higher numbers of AP elicited, not only after single AP (Figure 7). This is evocative of the phenotype of loss of RIMs, which are tethering factors of Ca2+-channels to the AZ (Kaeser et al., 2011; Brockmann et al., 2020). Thus, it is likely that decreased Ca2+-influx into the presynapse might be due to reduced number of Ca2+-channels at AZ in STX1-deficient neurons rather than due to an altered Ca2+-channel gating.

In fact, it is known that STX1 clusters together with Munc18-1 and SNAP25 also outside of AZ (Pertsinidis et al., 2013) and that it interacts with endoplasmic reticulum (ER) SNARE Sec22 at the ER-plasma membrane contact sites (Petkovic et al., 2014), both having potential functions in constitutive intracellular trafficking and regulation of the membrane lipid composition. In this regard, an impairment in general intracellular trafficking and/or membrane lipid composition as a result of loss of STX1, its presumed conformational change, and/or its deficient Munc18-1 binding imposed by LEOpen mutation might potentially lead to a decreased number of Ca2+-channels. However, just as there is not always a direct correlation between the Ca2+-channel abundance and the level of Ca2+-entry, there is not always a relationship between the Ca2+-entry and the level of EPSC and Pvr. An example for that is the overexpression of Ca2+-channel subunit α2δ also leading to overexpression of Ca2+-channels in the synapse (Hoppa et al., 2012). In these synapses, surprisingly however, a reduction in Ca2+-influx has been observed together with an increase both in EPSC and Pvr – similar to the phenotype of STX1ALEOpen neurons (Hoppa et al., 2012). Therefore, it is plausible that the increased SV fusogenicity might overcome the effect of low Ca2+-entry into the synapse and still lead to an increased EPSC and Pvr when the SVs are localized at a proper distance to the Ca2+-channels in STX1ALEOpen neurons (Figure 8). Plausibly, on the other hand, if increased SV fusogenicity is accompanied by a greater SV-Ca2+-channel distance as thought for the case of STX1ALEOpen + ΔΝ neurons, the Ca2+-sensitivity and the amplitude of EPSCs might approach back to the WT-like levels (Figure 8).

Together, our data suggest that even though deletion of N-peptide potentially reduces the number of reactive SNARE complexes (Burkhardt et al., 2008), which could explain the slower rate of the recovery of the RRP in neurons that express STX1A∆N2-19 or STX1A∆N2-28 (Figure 6), the level of this reduction appears to be not enough to decrease the baseline neurotransmitter release in STX1AWT (Figure 3) but only in STX1ALEOpen (Figure 4). The increase in the apparent Ca2+-sensitivity of the vesicular release in STX1ALEOpen neurons has also been attributed to the increased number of SNARE complexes (Acuna et al., 2014), yet the increased fusogenicity of the SVs in those neurons beclouds this hypothesis. Whether or not reduced number of SNARE complexes can lead to the robust effect of N-peptide deletions on STP is not clear, but likely, as the longer N-peptide deletions showed a trend towards smaller absolute values of EPSCs throughout the high-frequency stimulus (Figure 6—figure supplement 1). Since N-peptide mutations do not significantly change the initial PPR, the effect during 10 Hz trains can also be explained by an impaired Ca2+-channel–vesicle distance coupling and an accumulation of global Ca2+ during the train. Therefore, whether the facilitation–hindrance of SNARE complex formation leads to changes in STP and/or Ca2+-sensitivity of vesicular release should be investigated in depth. However, STX1AΔΝ2-9 neurons did not show any difference in the initial EPSCs but only in the latter phase of STP by up to ~30% larger EPSCs compared to that of STX1AWT neurons (Figure 6—figure supplement 1) and also no difference in the Ca2+-sensitivity (Figure 6) of the vesicular release. This suggests that the regulation of STP might be an important function of STX1A’s N-peptide aa 2–9 independent of Munc18-1 because Munc18-1 mutants that cannot bind to the STX1’s N-peptide do not manifest any regulatory effect on STP (Meijer et al., 2012).

Materials and methods

Animal maintenance and generation of mouse lines

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All procedures for animal maintenance and experiments were in accordance with the regulations of and approved by the animal welfare committee of Charité-Universitätsmedizin and the Berlin state government Agency for Health and Social Services under license number T0220/09. The generation of STX1-null mouse line was described previously (Arancillo et al., 2013; Vardar et al., 2016).

Neuronal cultures and lentiviral constructs

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Hippocampal neurons were obtained from mice of either sex at postnatal day (P) 0–2 and seeded on the already prepared continental or micro-island astrocyte cultures as described previously (Xue et al., 2007; Vardar et al., 2016). The neuronal cultures were then incubated for 13–20 DIV in NeurobasalA supplemented with B-27 (Invitrogen), 50 IU/ml penicillin and 50 µg/ml streptomycin at 37°C before experimental procedures. Neuronal cultures for EM and Ca2+-influx and those for neuronal viability, immunofluorescence labeling, and electrophysiology experiments were transduced with lentiviral particles at DIV 2–3 and DIV 1, respectively. Lentiviral particles were provided by the Viral Core Facility (VCF) of the Charité-Universitätsmedizin, Berlin, and were prepared as previously described (Vardar et al., 2016). The cDNA of mouse STX1A (NM_016801) was cloned in frame after an NLS-GFP-P2A sequence within the FUGW shuttle vector (Lois et al., 2002) in which the ubiquitin promoter was replaced by the human synapsin 1 promoter (f(syn)w). The improved Cre recombinase (iCre) cDNA was c-terminally fused to NLS-RFP-P2A. SynGCamp6f was generated analogous to synGCamp2 (Herman et al., 2014), by fusing GCamp6f (Chen et al., 2013) to the C terminus of synaptophysin and within the f(syn)w shuttle vector (Grauel et al., 2016).

Neuronal viability

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The in vitro viability of the neurons was defined as the percentage of the number of neurons alive at DIV15, 22, 29, 36, and 43 compared to the number of neurons at DIV 8. Phase-contrast bright-field images and fluorescent images with excitation wavelengths of 488 and 555 nm were acquired with a DMI 400 Leica microscope, DFC 345 FX camera, HCX PL FLUOTAR 10 objectives, and LASAF software (all from Leica). Fifteen randomly selected fields of 1.23 mm2 per well and two wells per group in each culture were imaged at different time points and the neurons were counted offline with the 3D Objects Counter function in Fiji software as described previously (Vardar et al., 2016). Sample size estimation was done as previously published (Vardar et al., 2016). MAP2 immunofluorescence labeling as shown in the figures is used only for representative purposes.

Immunocytochemistry

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The high-density cultured STX1A-/-; STX1Bflox/flox hippocampal neurons were co-transduced with Cre recombinase and with either STX1AWT or mutants at DIV 1–2. All the cultures were fixed with 4% paraformaldehyde (PFA) in 0.1 M phosphate-buffered saline, PH 7.4, for 10 min at DIV14-16. The neurons were then permeabilized with 0.1 % Tween–20 in PBS (PBST) for 45 min at room temperature (RT) and then blocked with 5% normal goat serum (NGS) in PBST. Primary antibodies were applied overnight at 4°C and subsequently secondary antibodies were applied for 1 hr at RT in the dark. High-density hippocampal cultures of 50 × 103 seeded neurons for neuronal viability analysis were treated with chicken polyclonal anti-MAP2 (1:2000; M2694; Merck Millipore) and then with Alexa Fluor (A) 647 donkey anti-chicken IgG (Jackson ImmunoResearch). High-density hippocampal cultures of 25 × 103 seeded neurons for protein expression analysis were treated with guinea pig polyclonal anti-Bassoon (1:1000; Synaptic Systems), mouse monoclonal anti-STX1A (1:1000; Synaptic Systems), and rabbit polyclonal Munc18-1 (1:1000; Sigma-Aldrich) and then with rhodamine red donkey anti-guinea pig IgG, A488 donkey anti-mouse IgG, and A647 donkey anti-rabbit IgG (all from Jackson ImmunoResearch). Autaptic neurons were treated with guinea pig polyclonal anti-VGlut1 (1:4000; Synaptic Systems), mouse monoclonal anti-STX1A (1:1000; Synaptic Systems), and rabbit polyclonal Munc18-1 (1:1000; Sigma-Aldrich) and subsequently with rhodamine red donkey anti-guinea pig IgG, A647 donkey anti-mouse IgG (both from Jackson ImmunoResearch), and Pacific-Blue goat anti-rabbit IgG (ThermoFisher). All secondary antibodies were diluted in 1:500 in PBST. The coverslips were mounted on glass slides with Mowiol mounting agent (Sigma-Aldrich). The images were acquired with an Olympus IX81 epifluorescence-microscope with MicroMax 1300YHS camera using MetaMorph software (Molecular Devices). Exposure times of excitations were kept constant for each wavelength throughout the images obtained from individual cultures. Data were analyzed offline with ImageJ as previously described (Vardar et al., 2016). Sample size estimation was done as previously published (Vardar et al., 2016). The number of synapses was analyzed by using Object Analyzer macro plug-in in ImageJ.

Electrophysiology

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The hippocampal autaptic neurons were co-transduced with Cre recombinase and with either STX1AWT or mutants at DIV 1–3. Sample size estimation was done as previously published (Rosenmund and Stevens, 1996). Whole-cell patch-clamp recordings were performed on glutamatergic autaptic hippocampal neurons at DIV 14–20 at RT with a Multiclamp 700B amplifier and an Axon Digidata 1550B digitizer controlled by Clampex 10.0 software (both from Molecular Devices). The recordings were analyzed offline using Axograph X Version 1.7.5 (Axograph Scientific).

Prior to recordings, the transduction of the neurons was verified by RFP and GFP fluorescence. Membrane capacitance and series resistance were compensated by 70%, and only the recordings with a series resistance smaller than 10 MΩ were used for further recordings. Data were sampled at 10 kHz and filtered by low-pass Bessel filter at 3 kHz. The standard extracellular solution was applied with a fast perfusion system (1–2 ml/min) and contained the following: 140 mM NaCl, 2.4 mM KCl, 10 mM HEPES, 10 mM glucose, 2 mM CaCl2, and 4 mM MgCl2 (300 mOsm; pH 7.4). Borosilicate glass patch pipettes were pulled with a multistep puller, yielding a final tip resistance of 2–5 MΩ when filled with KCl-based intracellular solution containing the following: 136 mM KCl, 17.8 mM HEPES, 1 mM EGTA, 4.6 mM MgCl2, 4 mM ATP-Na2, 0.3 mM GTP-Na2, 12 mM creatine phosphate, and 50 U/ml phosphocreatine kinase (300 mOsm; pH 7.4).

The neurons were clamped at –70 mV in steady state. To evoke EPSCs, the neurons were depolarized to 0 mV for 2 ms. The size of the RRP was determined by a 5 s application of 500 mM sucrose in standard external solution (Rosenmund and Stevens, 1996) and the total charge transfer was calculated as the integral of the transient current. Fusogenicity measurement was conducted by application of 250 mM sucrose solution for 10 s and calculation of the ratio of the charge transfer of the transient current over RRP. For the analysis of the RRP recovery, a paired pulse of 5 s long 500 mM sucrose was applied with a time interval of 2 s. Spontaneous release was determined by monitoring mEPSCs for 30–60 s at –70 mV. To correct false-positive events, mEPSCs were recorded in the presence of 3 µM AMPA receptor antagonist NBQX (Tocris Bioscience) in standard external solution. The spontaneous release rate was assessed by the division of the mEPSC frequency over the number of primed vesicles to determine the fraction of the RRP released per second by spontaneous release.

For Ca2+-sensitivity assays, EPSCs were evoked in extracellular solution containing 1 mM MgCl2 and either 0.5, 1, 2, 4, or 10 mM CaCl2. In between the test extracellular solution applications, standard extracellular solution was applied. For each concentration, six APs were elicited at 0.2 Hz. To control for rundown and cell-to-cell variability, the test solutions were applied either in increasing or decreasing concentration order for equal number of neurons and test responses were normalized to average EPSCs priorly recorded in standard external solution. Normalized responses were then normalized to the response in 10 mM CaCl2. The normalized values were fitted into a standard Hill equation.

SynGcamp6f-imaging

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Imaging experiments were performed at DIV 13–16 on autapses in response to a single stimulus and stimuli trains of 10 Hz as described previously for SynGcamp2-imaging (Herman et al., 2014). Images were acquired using a 490 nm LED system (pE2; CoolLED) at a 5 Hz sampling rate with 25 ms of exposure time. The acquired images were analyzed offline using ImageJ (National Institute of Health), Axograph X (Axograph), and Prism 8 (GraphPad, San Diego, CA). Sample size estimation was done as previously published (Herman et al., 2014).

High-pressure freezing and electron microscopy

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The high-density cultured hippocampal neurons were co-transduced with Cre recombinase and with either STX1 WT or mutants at DIV 2–3 and high-pressure fixed under a pressure of 2100 bar in standard extracellular recording solution using an HPM 100 Leica or ICE Leica high-pressure freezer at DIV 14–16. Samples were then transferred into cryovials containing 1% glutaraldehyde, 1% osmium tetroxide, 1% ddH20 (Millipore) in anhydrous acetone and processed in an AFS2 automated freeze-substitution device (Leica) followed by a temperature ramp from −90°C for 5 hr, from −90°C to −50°C (8°C/hr), from −50°C to −20°C (6°C/hr), for 12 hr at −20°C, from −20 to +20°C (10°C/hr). After the freeze-substitution step, samples were embedded in epoxy epon 812 (EMS). Finally, samples were placed into capsules filled with pure epoxy epon 812 and further polymerized for 48 hr at 60°C. Randomly selected areas of ~250 μm2 containing neurons were ultracut into 40 nm thick slices using an Ultracut UCT ultramicrotome (Leica) and collected on 0.5% formvar-coated 200-mesh copper grids (EMS). Those sections were contrasted with 0.1% uranyl acetate for 1 hr and lead citrate (0.15 lead citrate, 0.12 m sodium citrate in ddH2O). Images were collected blindly in an FEI Tecnai G20 electron microscope operating at 200 keV and digital images taken with a Veleta 2k × 2k CCD camera (Olympus). Synapses were analyzed blindly using an analysis program developed for ImageJ and MATLAB (Watanabe et al., 2013). AZs were defined as the membrane stretch directly opposite to the postsynaptic density, and docked vesicles were defined as those in direct contact with the plasma membrane. SV distribution was analyzed by calculating the shortest distance of each vesicle to the AZ membrane and binned with 5 nm. Sample size estimation was done as published previously (Watanabe et al., 2013).

Statistical analysis

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Data in bar graphs present single observations (points) and means ± standard error of the mean (SEM; bars). Data in x–y plots present means ± SEM. All data were tested for normality with Kolmogorov–Smirnov test. Data from two groups with normal or nonparametric distribution were subjected to Student’s two-tailed t-test or Mann–Whitney nonparametric test, respectively. Data from more than two groups were subjected to Kruskal–Wallis followed by Dunn’s post hoc test when at least one group showed a nonparametric distribution. For data in which all the groups showed a parametric distribution, one-way ANOVA test followed by Tukey’s post hoc test was applied. For STP measurements, two-way ANOVA test was used. All the tests were run with GraphPad Prism 8.3, and all the statistical data are summarized in the corresponding source data tables.

Data availability

All data generated or analysed during this study are included in the manuscript and supporting files. We uploaded source data files which show summary tables of mean, SEM, median, number of independent cultures, number of independent measurements, real p value for each test performed, and statistical test used for each separate figure.

References

Decision letter

  1. Axel T Brunger
    Reviewing Editor; Stanford University, United States
  2. Gary L Westbrook
    Senior Editor; Oregon Health and Science University, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

In this study Vardar et al., use patch-clamp electrophysiology in autaptic neurons to provide a systematic analysis of the different roles of the N-terminal domains and different Munc18 binding modes of Syntaxin-1. The complexity of the interaction between Munc18 and syntaxin and their isoforms in other species has led to a vexing complexity involving different interactions that appear to range from essential to dispensable in different experiments and contexts. This study compares many aspects of these interactions in the same neuronal system. The paper confirms previous observations, but also arrives at new conclusions. The authors show that the Habc-domain is essential for syntaxin's role in neurotransmitter release, while the N-peptide has a modulatory role. Disrupting both binding modes of Syntaxin-1 with M18 leads to strongly reduced levels of both proteins while neurotransmitter release can still occur.

Decision letter after peer review:

Thank you for submitting your article "Reexamination of N-terminal domains of Syntaxin-1 in vesicle fusion from central murine synapses" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Gary Westbrook as the Senior Editor. The reviewers have opted to remain anonymous. The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential Revisions (for the authors):

1. The authors attempt to address the reduced protein expression by titrating down WT levels by using only 1/12 of the viral load. Strikingly, reducing WT STX1A levels by almost 6-fold had no effect on synaptic transmission. This raises important questions. At which point does STX1A become rate-limiting? And how do these protein levels compare to endogenous STXA levels? The authors should show the expression levels of their mutants relative to endogenous STX levels. Furthermore, a previous paper from the same group concluded that titrating Synatxin1 did impact synaptic transmission (Arancillo et al., J Neurosci 2013). The authors should discuss how their expression levels and findings compare to the findings in their previous paper.

2. While the effect of the N-peptide on most release parameters is absent/small, a robust effect was observed on short-term plasticity: up to70% larger EPSC's at the end of a 10Hz train in N-peptide mutants. This substantial effect is undervalued by the authors. Regulation of STP could be an important function of this domain. The authors mention this effect only briefly in the abstract, but should discuss this biological relevance in the Discussion. Second, typical traces of trains in Figure 6 should be of higher resolution and zoomed in more to critically evaluate. For instance, the standing (asynchronous) current that usually develops during 10Hz trains in autapses seems to be absent. Why is this the case? Since N-peptide mutations do not change PPR, the most logical explanation of the effect during 10Hz trains is a difference in the accumulation of global Ca during the train. This is plausible given the (disputed) interaction of syntaxin with calcium channels. Ideally, the experiments should be repeated +/- EGTA-AM. In this same experiment, the author can examine the potential effect on Ca-dependent replenishment or increase in fusogenicity (measuring RRP size and fusogenicity before and after 10Hz trains). At the minimum, the observed effect should be more pointed out more clearly in the text.

3. The main hypothesis put forward for reduced Ca-sensitivity and impaired RRP recovery in N-peptide mutants is an impairment in SNARE complex formation. How does this reconcile with up to 70% larger EPSC's during 10 Hz trains? Please comment and discuss.

4. The data in figure 6 on the interaction between the two M18 binding modes of Stx1 are intriguing. However, different effects of N-peptide mutations in the WT and LE_Open background are difficult to reconcile, and not further addressed in the discussion. It is therefore difficult to interpret these data in the light of the N-peptide function. Please comment and discuss.

5. In Figure 6F the N-peptide mutations in Stx1 LE_Open slow down depression during the first few stimulations, but have no effect on STP at the end of the train. This is opposite from the effect found at the WT background, where these mutations affect STP at the end of the train (Figure 6A). How these seemingly different results can be reconciled should be addressed in the discussion.

6. Given the twofold higher EPSC amplitude and Pvr for Stx1 LE_Open, this mutant will most likely end up with similar EPSC amplitudes at the end of the train as for the WT. However, the LE_Open Δ_N mutants will have lower EPSC amplitudes at the end of the train. This could lead to different conclusions about the role of the N-peptide, especially when combined with similar experiments with these mutants as suggested in point 2.

7. It is interesting to see in Figure 6J that, for low external Ca, N-peptide mutations on a LE_open background rescue the paired-pulse ratio to WT levels, while for increasing external Ca levels this gradually reverts to the LE_open phenotype. This should be addressed in the discussion in the light of the different Ca sensitivities and fusogenicities for the different experimental groups.

8. If reduced Ca-influx explains reduced Ca affinity in LE_open mutant (Figure 6I), how does this reconcile with the LE_open-deltaN mutant, which has significant less Ca influx compared to WT (Figure 7E), but no significant difference in Ca sensitivity? Some explanation should be added to the Discussion.

9. The authors' interpretation that the effect of calcium influx relates to expression levels and survivability is somewhat challenged by the fact that the LE mutant has the strongest effect on calcium influx, while it is by far not the mutant with the lowest expression levels, nor is it experiencing a decrease in survivability. Please comment and discuss.

10. Throughout the manuscript, the authors quantify cell numbers as a measure of survival. Some mutations, for example syntaxin knockout and the DeltaHabc mutant rescue, strongly affect cell survival. For the interpretation of the synaptic physiology, a more meaningful parameter would be synapse numbers, especially for evaluating the relationship between docking (measured per synapse profile) and sucrose RRP (measured per cell). It appears possible, if not likely, that synapses are lost before cells die, and in this case correlating RRP and docking may be strongly confounded. The authors may have the data in mass cultures already, and it should be possible to quantify synapse numbers (for example via Bassoon puncta densities) in these existing data. This should be accompanied by a discussion of the systems used (mass cultures for morphology and autaptic cultures for electrophysiology). If synapse numbers are changed, the RRP charge and # of docked vesicles should not be directly correlated (as done in Figure 5E), because RRP charge strongly depends on synapse numbers, while the number of docked vesicles per synapse does not. The discussion should be adjusted accordingly.

11. The authors present the double mutation of opening syntaxin (LE mutant) and N-peptide deletion as "interruption of both Munc18-1 binding modes" (caption line 185, figure captions Figures 4 and 5). While we appreciate where the terminology for considering this a "null for Munc18 binding" comes from, it would be much better to stay closer to the experiments in the data presentation and use wording like "opening of syntaxin combined with N-peptide deletion" instead. As the authors discuss, Munc18-syntaxin interactions are complex, and most data suggest that Munc18 also binds to SNARE complexes (and open syntaxin) while they are being assembled. Opening syntaxin and deleting the N-peptide does unlikely fully abolish Munc18 binding, but biases binding of syntaxin to Munc18 alone towards binding of Munc18 to syntaxin as it is being incorporated into SNARE complexes. The authors should do justice to this point in figure captions, and should discuss it as such.

12. The effect of the LE mutant of syntaxin has been characterized by several biophysical methods that show that it shifts the conformational equilibrium towards the open state, but it still adopts the closed state upon Munc18 binding as observed by SAXS (Colbert et al. 2013) and single molecule FRET (Wang et al., EMBO J. 36, 816-, 2017). Moreover, the LE mutant only partially compensates for deletion of Munc13, i.e., the rescue is rather limited as observed in autaptic neuronal cultures and by single molecule FRET experiments (Lai et al., Neuron 95, 591-, 2017). Moreover, the results in Wang et al., 2017 suggest that the LE mutant interferes with proper Munc13 function (via its interactions with syntaxin), which may be related to the observed phenotypes of this mutant. Please discuss the experiments presented here in the context of these biophysical findings.Reviewer #1 (Recommendations for the authors):

In this study Vardar et al., use patch-clamp electrophysiology in autaptic neurons to provide a systematic analysis of the different roles of the N-terminal domains and different Munc18 binding modes of Syntaxin-1. The complexity of the interaction between Munc18 and Syntaxin has led to a long-standing controversy about the importance of certain interactions, ranging from absolutely essential to dispensable. This study is the first to compare all aspects in a side-by-side comparison in the same, native model system. The paper confirms previous observations but also arrives at new conclusions. The authors show that the Habc-domain is essential for Stx1's role in neurotransmitter release, while the N-peptide has a minor modulatory role. Disrupting both binding modes of Stx1 with M18 leads to strongly reduced levels of both proteins while neurotransmitter release can still occur.

This manuscript provides the most comprehensive study to date on the effects of mutant Syntaxin variants to assess their effects on synaptic transmission, -efficacy, and a lot of other synaptic determinants. The manuscript contains an extensive set of experiments of high-quality, which are clearly described and presented in the text and figures. All conclusions are justified and built on solid evidence. The main limitations of the current study are the strongly reduced expression levels of some mutants, an issue the authors also raise, which limit some of the interpretations, and the effects of N-peptide mutants which are underdeveloped. However, taken together, this study is of great interest to the synapse field and has all the ingredients to provide the final verdict on some controversial issues.

Major points:

1. The authors attempt to address the reduced protein expression by titrating down WT levels by using only 1/12 of the viral load. Strikingly, reducing WT STX1A levels by almost 6-fold had no effect on synaptic transmission. This raises important questions. At which point does STX1A become rate-limiting? And how do these protein levels compare to endogenous STXA levels? The authors should show the expression levels of their mutants relative to endogenous STX levels. Furthermore, a previous paper from the same group concluded that titrating Synatxin1 did impact synaptic transmission (Arancillo et al., J Neurosci 2013). The authors should discuss how their expression levels and findings compare to the findings in their previous paper.

2. While the effect of the N-peptide on most release parameters is absent/small, a robust effect was observed on short-term plasticity: up to70% larger EPSC's at the end of a 10Hz train in N-peptide mutants. This substantial effect is undervalued by the authors. Regulation of STP could be an important function of this domain. The authors mention this effect only briefly in the abstract, but should discuss this biological relevance in the Discussion. Second, typical traces of trains in Figure 6 should be of higher resolution and zoomed in more to critically evaluate. For instance, the standing (asynchronous) current that usually develops during 10Hz trains in autapses seems to be absent. Why is this the case? Since N-peptide mutations do not change PPR, the most logical explanation of the effect during 10Hz trains is a difference in the accumulation of global Ca during the train. This is plausible given the (disputed) interaction of syntaxin with calcium channels. To test this the experiments should be repeated +/- EGTA-AM. In this same experiment, the author can examine the potential effect on Ca-dependent replenishment or increase in fusogenicity (measuring RRP size and fusogenicity before and after 10Hz trains).Reviewer #2 (Recommendations for the authors):

This manuscript by Vardar, Rosenmund et al. provides a systematic analysis of syntaxin-1 function in exocytosis. It probes the major molecular roles of syntaxin through mutational analyses of three of its key features: the N-terminal peptide interaction with Munc18 (using short deletions), the role of the Habc domain (using a larger deletion), and the role of a confirmational switch (using the established LE open mutation). It probes the synaptic functions of these features either on their own or in combination with one another in rescue experiments in cultured neurons of syntaxin-1A and 1B null mutants, assessing synaptic transmission and its constituents, synaptic ultrastructure and neuronal survival.

The main finding is that the N-terminal peptide interaction with Munc18 is dispensable, and its role is only revealed when combined with the LE open mutation. Another finding is that syntaxin function is quite resilient, as even combining mutations (for example LE open and N-peptide deletion) does not fully disrupt synaptic transmission. In general, the finding that the N-peptide deletion is dispensable is important as it differs from previous studies that have used different manipulations. A key strength is the systematic comparison of the various mutants alone and in combination upon syntaxin-1 ablation and the testing of a series of different N-terminal deletions. The authors provide a balanced discussion of their and previous results, how they relate to one another, and how discrepancies could be reconciled.

Weaknesses in the current methodology include that the manuscript does not distinguish well between effects on cell health, effects on synapse numbers, and effects on synaptic properties. Furthermore, some of the data presentation relies on assumptions in the field rather than directly relating to the experimental approach. For example, the data describing the LE mutant and the N-peptide deletion assume that these mutations together essentially reveal a "complete loss-of-Munc18-binding" phenotype, but this may not necessarily be that case.Reviewer #3 (Recommendations for the authors):

In this manuscript, Vardar et al., investigated the role of different syntaxin I (STX1) domains in neurotransmitter release using a STX1-null mouse model system and exogenous reintroduction of STX1A mutants, the latter lacking either the N-peptide or the Habc domain of STX I. In addition, the STX1 mutant, which is present in the ‚open' conformation (LE open mutant) was examined with or without deletion of the N-peptide. The results show that the Habc domain is absolutely necessary for the stability or expression of STX1 and thus for neurotransmitter release. Moreover, it became clear, contrary to earlier work, that the N-peptide is dispensable for synaptic transmission, but assumes a regulatory role in controlling ca2+ sensitivity of vesicular release and generally in vesicle fusion when syntaxin is in the 'open' conformation. The manuscript provides a very comprehensive structure-function analysis using a wide variety of high-resolution techniques (e.g. HPF-EM, synaptic ca2+ imaging). Remarkably, perturbation of the STXI-Munc18 interaction by interfering with the canonical interaction modes has surprisingly little effect on synaptic transmission. Several results in this present manuscript correct observations of previous studies that used less quantitative and systematic approaches in the attempt to unravel the molecular mechanisms of syntaxin-Munc18 interaction in ca2+-driven exocytosis.

In general, the combined set of data is particularly valuable and allows new insights into the complexity of neuronal transmission and its underlying components.

https://doi.org/10.7554/eLife.69498.sa1

Author response

Essential Revisions (for the authors):

1. The authors attempt to address the reduced protein expression by titrating down WT levels by using only 1/12 of the viral load. Strikingly, reducing WT STX1A levels by almost 6-fold had no effect on synaptic transmission. This raises important questions. At which point does STX1A become rate-limiting? And how do these protein levels compare to endogenous STXA levels? The authors should show the expression levels of their mutants relative to endogenous STX levels. Furthermore, a previous paper from the same group concluded that titrating Synatxin1 did impact synaptic transmission (Arancillo et al., J Neurosci 2013). The authors should discuss how their expression levels and findings compare to the findings in their previous paper.

The aim of our down-titration experiments was not to determine which level of exogenous STX1A expression limits the neurotransmitter release, but we rather aimed to test whether the relative reduction in EPSC and in other release parameters in STX1ALEOpen + ΔΝ neurons compared to that of STX1ALEOpen neurons was due to the relative reduction in their STX1A levels. Therefore, we have reduced the number of viral particles used for 35 mm well from ~400.000 (1X) to ~35.000 for STX1AWT and to ~ 65.000 for STX1ALEOpen and successfully reached the levels of STX1ALEOpen + ΔΝ2-28. At this point, we did not observe any influence of expression levels on the release parameters neither by using STX1AWT nor STX1ALEOpen. Therefore, we suggested that the reduction in the release parameters due to the N-peptide deletions on STX1ALEOpen background is not due to its relatively lower expression levels but rather due to a functional deficit.

However, we thank to the reviewers for their suggestion to compare the endogenous STX1A levels to the exogenous STX1A levels used in this study to solve a possible discrepancy between this study and our previous study (Arancillo et al., 2013). It is important to note that our mouse line has the genotype of STX1BFL/FL; STX1A-/-. We do not currently hold a mouse line with the genotype of STX1BFL/FL; STX1A+/- for breeding. Therefore, we cannot compare the endogenous and exogenous expression levels of STX1A using littermates. Because of that, we analyzed the endogenous STX1A levels in WT neurons in comparison to the lentiviral expression obtained with the help of 1X volume of lentiviral particles encoding STX1A in STX1-null neurons. However, we cultured both WT and the transgenic neurons on the same batch of astrocyte cultures and fixed them with PFA at the same DIV. We have observed that 1X volume of viral particles led to ~3 fold higher expression of STX1A compared to the endogenous levels (Figure 4 – Supplement 3). This shows that even the lowest expressing construct STX1ALEOpen+ΔN2-28 reaches an expression level of ~60% of endogenous STX1AWT. This also suggests that with our down-titration experiment we have reached again 60% of endogenous levels. We have explained and discussed our new data in relation to our previous study (Arancillo et al., 2013) in lines 236-252.

It should be noted that the WT neurons also express STX1B. It is fair to interpret our data as such that our experiments do not fall within over expression studies, because the neurons used in this study do not contain STX1B. Therefore, a ~3 fold higher expression of STX1A in the absence of STX1B constitutes a total level of STX1 comparable to that in WT neurons. However, because the STX1ALEOpen+ΔN2-28 do show a 60% expression of STX1A, we have removed our statement that ‘neurotransmitter release precedes normal even when both STX1A and Munc18-1 show a severely low expression’. Now, we have included our new interpretation that our down-titration experiments are insufficient to reach the rate-limiting level as determined by our previous study and that 60% of STX1A in the absence of STX1B is inadequate to rescue Munc18-1 levels (lines 242-249).

2. While the effect of the N-peptide on most release parameters is absent/small, a robust effect was observed on short-term plasticity: up to70% larger EPSC's at the end of a 10Hz train in N-peptide mutants. This substantial effect is undervalued by the authors. Regulation of STP could be an important function of this domain. The authors mention this effect only briefly in the abstract, but should discuss this biological relevance in the Discussion. Second, typical traces of trains in Figure 6 should be of higher resolution and zoomed in more to critically evaluate. For instance, the standing (asynchronous) current that usually develops during 10Hz trains in autapses seems to be absent. Why is this the case? Since N-peptide mutations do not change PPR, the most logical explanation of the effect during 10Hz trains is a difference in the accumulation of global Ca during the train. This is plausible given the (disputed) interaction of syntaxin with calcium channels. Ideally, the experiments should be repeated +/- EGTA-AM. In this same experiment, the author can examine the potential effect on Ca-dependent replenishment or increase in fusogenicity (measuring RRP size and fusogenicity before and after 10Hz trains). At the minimum, the observed effect should be more pointed out more clearly in the text.

Firstly, we thank to the reviewers for pointing out the importance of our finding of STX1A’s N-peptide’s regulation of STP. Now, we have discussed this issue also in our discussion with a greater emphasis (lines 520-538).

Secondly, we have now included zoomed-in example traces of absolute EPSCs for the first and last 5 stimuli for all the groups (Figure 6 – Supplement 2) for a better inspection, as suggested by the reviewers. It can also be observed in those zoomed-in traces that 10 Hz stimulation of autaptic neurons is not adequate for production of a measurable standing current. Usually, the currents elicited by 1 AP reach back the baseline in less than 100 ms, which is the time-interval for 10 Hz stimulation. Additionally, the decay-time of the EPSC affects the time required for the current to be back at baseline level and in autaptic neurons the bigger EPSCs can lead to longer decay-times and thus can contribute to the standing current (in-house information based on our previous experiments using low concentration of AMPA antagonists to produce smaller EPSCs). A reason for the effect of EPSC size on the decay-time and thus the standing current is the spill-over of glutamate because the autaptic neurons are not a fully-isolated system on the contrary of slice cultures. We understand that the reviewers would like to know the effect of N-peptide deletions and the LEOpen mutation on asynchronous release. However, more elaborate experiments such as the usage of Strontium instead of Calcium in the extracellular solution (Friedmann and Regehr, 1999, Biophys J) would be required.

Finally, we would like to raise some of our concerns regarding the usage of EGTA-AM for high-frequency stimulation. First of all, EGTA-AM experiments require pre-incubation of the neurons in EGTA-AM before electrophysiological recordings and thus leads to poorly controlled concentration of this ca2+-buffer in the neurons which ultimately may contaminate the results of high-frequency stimulation. Whereas small differences in the intracellular EGTA-AM concentration among individual neurons of one group might be negligible when the differences in the ca2+-sensitivity of the vesicular release and the RRP replenishment between two groups are high, they might greatly influence the interpretation of the data when those differences between two groups are small as in our study. Furthermore, increasing ca2+-buffering leads to complex changes in ca2+-transients during the trains of APs. During the train, ca2+-buffer saturates and removal of ca2+ through plasmalemmal pumps gets impaired leading to elevated intracellular ca2+-levels due to the extended time of the removal of ca2+ through the pumps at the end of the train. This further complicates the interpretation of the 10 Hz data using EGTA-AM particularly when the purpose of the study is the dissection of the mechanisms leading to small changes in STP and RRP replenishment. Thus, we prefer not to utilize EGTA-AM to examine the potential effect on ca2+-dependent replenishment or increase in fusogenicity. We would like to keep the focus of our study in that N-peptide is not indispensable for neurotransmitter release but rather regulates the STP and ca2+-sensitivity of the vesicular release.

However, we have discussed the N-peptides’s role in STP more with with a greater emphasis (lines 520-538) as mentioned before.

3. The main hypothesis put forward for reduced Ca-sensitivity and impaired RRP recovery in N-peptide mutants is an impairment in SNARE complex formation. How does this reconcile with up to 70% larger EPSC's during 10 Hz trains? Please comment and discuss.

We thank to the reviewers for raising this issue. We have calculated the net depression of EPSCs during 10 Hz trains for N-peptide deletions and saw that the responses recorded from STX1AWT neurons depress down to 60% of the initial EPSC size and EPSCs recorded from N-peptide mutants depress down to 80-90%. We also have plotted the absolute EPSCs and observed that the EPSCs recorded from STX1AΔN2-19 and STΧ1AΔN2-28 did not become larger but inclined to remain smaller compared to those recorded from STX1AWT neurons (Figure 6 – Supplement 1). However, because STX1AΔN2-9 had initial EPSC comparable to that of STX1AWT and because those neurons showed short-term depression of only 20%, the absolute EPSCs became ~30 % larger compared to that of STX1AWT. We have now discussed this issue in the light of their capability of SNARE complex formation and tuned down our interpretation (lines 520-529).

4. The data in figure 6 on the interaction between the two M18 binding modes of Stx1 are intriguing. However, different effects of N-peptide mutations in the WT and LE_Open background are difficult to reconcile, and not further addressed in the discussion. It is therefore difficult to interpret these data in the light of the N-peptide function. Please comment and discuss.

We have already discussed this issue in our discussion (lines 426-441, especially lines 437-441). In brief, the facilitation of the SNARE complex formation by N-peptide and the tightening of the Munc18-1 bound closed conformation by the absence of N-peptide are only minor (Burkhardt, 2008; Colbert, 2013). Furthermore, N-peptide’s effect might become negligible in a membranous environment when STX1’s TMR is present (Dawidowski and Cafiso, 2013).

We also propose that N-peptide regulates ca2+-sensitivity of the vesicular release, which is a common phenotype of N-peptide deletions on the WT and LEOpen background. However, we rather remain conservative for that issue and only speculate about this ‘newly discovered’ function of N-peptide.

5. In Figure 6F the N-peptide mutations in Stx1 LE_Open slow down depression during the first few stimulations, but have no effect on STP at the end of the train. This is opposite from the effect found at the WT background, where these mutations affect STP at the end of the train (Figure 6A). How these seemingly different results can be reconciled should be addressed in the discussion.

STP behavior upon a high-frequency stimulus depends on the initial Pvr, fusogenicity, and the ca2+-sensitivity of the SVs. We show in our study that deletion of N-peptide does not influence the fusogenicity of the vesicles neither on STX1AWT nor on STX1ALEOpen background (Figure 6). However, LEOpen mutation increases the degree of the SV fusogenicity, which remains high also when N-peptide is deleted (Figure 6). Therefore, the STX1ALEOpen+ΔΝ mutants show a high degree of depression upon 10 Hz stimulation. On the other hand, we also show that N-peptide deletion decreases the ca2+-sensitivity of the vesicular release both on STX1AWT and on STX1ALEOpen background (Figure 6). However, the decrease of the ca2+-sensitivity on STX1AWT background is relatively minor compared to that on STX1ALEOpen background. This is also evident in that the decrease of the ca2+-sensitivity on STX1AWT is not enough to significantly reduce the initial Pvr and EPSC. Therefore, it is plausible that the effect of global ca2+-accumulation in the presynapse can be observed only in the late phase of the stimulus, when N-peptide is deleted on STX1AWT background. However, STX1ALEOpen+ΔΝ mutants also show a high degree of fusogenicity and that might nullify the effect of reduced ca2+-sensitivity at the end of the train of APs. Our interpretations regarding this issue can be found at lines 335-338, 348-350, and 473-491.

6. Given the twofold higher EPSC amplitude and Pvr for Stx1 LE_Open, this mutant will most likely end up with similar EPSC amplitudes at the end of the train as for the WT. However, the LE_Open Δ_N mutants will have lower EPSC amplitudes at the end of the train. This could lead to different conclusions about the role of the N-peptide, especially when combined with similar experiments with these mutants as suggested in point 2.

We already have shown the absolute values of EPSCs during a 10 Hz stimulus for STX1AWT and for all the mutants (Figure 6 – Supplement 1). Whereas STX1A∆N2-19 and STX1A∆N2-28 tended to remain to elicit smaller EPSCs throughout the high-frequency stimuli compared to those of STX1AWT (Figure 6- supplement 1), EPSCs recorded from STX1A∆N2-9 neurons became ~30 % larger at the end of the stimulus compared to that of STX1AWT (lines 290-294). On the other hand, the EPSCs recorded from STX1ALEOpen neurons by 10 Hz stimulation tended to be initially larger, but they declined further compared to that of STX1AWT neurons (Figure 6 —figure supplement 1). STX1ALEOpen+∆N2-28 mutant remained to produce smaller EPSCs throughout the high-frequency stimuli compared to those of both STX1AWT and STX1ALEOpen (Figure 6 —figure supplement 1) (lines 328-333).

These findings render the hypothesis likely that N-peptide deletions both on STX1AWT and STX1ALEOpen background reduce the efficacy of SNARE complex formation and therefore leads to smaller EPSCs throughout the train. The exception for that observation is that STX1AΔΝ2-9 neurons show EPSCs initially comparable to that of STX1AWT, which does not reconcile with the hypothesis that the ca2+-sensitivity of SVs and STP is affected by N-peptide deletions due to an impairment in SNARE complex formation. We have now discussed this issue more in detail in lines (520-535).

7. It is interesting to see in Figure 6J that, for low external Ca, N-peptide mutations on a LE_open background rescue the paired-pulse ratio to WT levels, while for increasing external Ca levels this gradually reverts to the LE_open phenotype. This should be addressed in the discussion in the light of the different Ca sensitivities and fusogenicities for the different experimental groups.

We already have discussed this issue in lines 348-350 and suggested that the observation that N-peptide deletion leads to an altered behavior only in the initial phase of the 10 Hz stimuli – when STX1A’s open conformation is facilitated – is consistent with the reduced ca2+-sensitivity (Figure 6I) but unaltered fusogenicity (Figure 6F) of the vesicles. A cartoon illustrating a speculative model for the effects of fusogenicity, ca2+-channel-SV coupling, and ca2+-sensitivity of vesicular release can be found in Figure 8.

8. If reduced Ca-influx explains reduced Ca affinity in LE_open mutant (Figure 6I), how does this reconcile with the LE_open-deltaN mutant, which has significant less Ca influx compared to WT (Figure 7E), but no significant difference in Ca sensitivity? Some explanation should be added to the Discussion.

We would like to emphasize that our findings show that LEOpen mutant does not reduce but enhances the ca2+-affinity of the vesicular release, which is in line with previous reports (Gerber, et al., 2008; Acuna et al., 2014). In Figure 6I, the ca2+-dose response curve of STX1ALEOpen deviates from that of STX1AWT with a leftward shift showing that STX1ALEOpen neurons release a bigger fraction of their release capacity at lower ca2+-concentration, thus they have a higher ca2+-sensitivity.

ca2+-sensitivity of the vesicular release is a convoluted function of SV fusogenicity and ca2+-channel–SV distance coupling. Our sub-saturating sucrose application shows that STX1ALEOpen leads to a higher fusogenicity, which contributes to the higher apparent ca2+-sensitivity. However, we cannot conclude from our experiments that the vesicles are docked and primed at a closer distance to ca2+-channels, which should be further investigated. Therefore, we argue that even though ca2+-channel concentration might be reduced in STX1ALEOpen neurons, which should be further corroborated with further experiments, ca2+-sensitivity might be increased with adequate ca2+-channel–SV distance and increased SV fusogenicity.

On the other hand, STX1ALEOpen+∆N mutants might lead to an increase in the distance between docked/primed vesicles and the ca2+-channels, which is likely due to the loss of N-Peptide. Because those neurons retain high SV fusogenicity due to the ‘opening’ of STX1A, ca2+-sensitivity of the SV fusion as a convoluted function of SV fusogenicity and ca2+-channel–SV distance coupling would approach back towards the WT-like level.

We have already had discussed this issue in our previous version of the manuscript (previous version 468-483). However, we incorporated a more elaborate discussion regarding the relationship between ca2+-influx, fusogenicity, and ca2+-sensitivity (lines 509-519) and we hope we could address your concern. Furthermore, a cartoon illustrating a speculative model for the effects of fusogenicity, ca2+-channel-SV coupling, ca2+-influx, and ca2+-sensitivity of vesicular release can be found in Figure 8.

9. The authors' interpretation that the effect of calcium influx relates to expression levels and survivability is somewhat challenged by the fact that the LE mutant has the strongest effect on calcium influx, while it is by far not the mutant with the lowest expression levels, nor is it experiencing a decrease in survivability. Please comment and discuss.

We haven’t directly related the expression levels of STX1A to the level of global ca2+-influx, but rather proposed it as a possible hypothesis (lines 359-363). Therefore, we measured ca2+-influx from neurons expressing STX1AWT but at a level as reduced as STX1ALEOpen+∆N2-28 and found no difference. In this respect, we have interpreted our data that STX1ALEOpen reduces ca2+-influx due to a functional impairment rather than due to low expression level (lines 363-372). We are sorry if there was a confusion.

10. Throughout the manuscript, the authors quantify cell numbers as a measure of survival. Some mutations, for example syntaxin knockout and the DeltaHabc mutant rescue, strongly affect cell survival. For the interpretation of the synaptic physiology, a more meaningful parameter would be synapse numbers, especially for evaluating the relationship between docking (measured per synapse profile) and sucrose RRP (measured per cell). It appears possible, if not likely, that synapses are lost before cells die, and in this case correlating RRP and docking may be strongly confounded. The authors may have the data in mass cultures already, and it should be possible to quantify synapse numbers (for example via Bassoon puncta densities) in these existing data. This should be accompanied by a discussion of the systems used (mass cultures for morphology and autaptic cultures for electrophysiology). If synapse numbers are changed, the RRP charge and # of docked vesicles should not be directly correlated (as done in Figure 5E), because RRP charge strongly depends on synapse numbers, while the number of docked vesicles per synapse does not. The discussion should be adjusted accordingly.

We thank to the reviewers for raising the point of a possible alteration of synapse number in STX1ALEOpen and STX1ALEOpen+∆N2-28 neurons as a possible reason for the reduction in their RRP. To test this hypothesis, we analyzed our VGlut1 immunocytochemical images of autaptic neurons using the images which were shown before in the Figure 4 – Supplement 2. We have found no difference in the synapse number nor in the synapse area among STX1AWT, STX1ALEOpen, and STX1ALEOpen+∆N2-28 neurons. We have included our finding as Figure 5—figure supplement 1 and the numerical values can be found in the respective source data. Because we found no reduction in synapse number but in RRP, we have kept our initial comment for the correlation between the number of docked vesicles and the RRP charge. On the other hand, we noted that the vesicle docking is assessed in mass cultures whereas RRP charge is assessed in autaptic cultures. Our explanation of our finding of the synapse number can be found in lines 265-272.

11. The authors present the double mutation of opening syntaxin (LE mutant) and N-peptide deletion as "interruption of both Munc18-1 binding modes" (caption line 185, figure captions Figures 4 and 5). While we appreciate where the terminology for considering this a "null for Munc18 binding" comes from, it would be much better to stay closer to the experiments in the data presentation and use wording like "opening of syntaxin combined with N-peptide deletion" instead. As the authors discuss, Munc18-syntaxin interactions are complex, and most data suggest that Munc18 also binds to SNARE complexes (and open syntaxin) while they are being assembled. Opening syntaxin and deleting the N-peptide does unlikely fully abolish Munc18 binding, but biases binding of syntaxin to Munc18 alone towards binding of Munc18 to syntaxin as it is being incorporated into SNARE complexes. The authors should do justice to this point in figure captions, and should discuss it as such.

We are sorry for using a terminology which might be misleading for the readers. We already have commented in our discussion that Munc18-1 is probably still bound to STX1A and/or SNARE complexes even when N-Peptide is deleted in STX1’s open conformation (lines 416-425). We have corrected the caption line 186 and the figure captions of Figures 4 and 5 as the reviewers suggested.

12. The effect of the LE mutant of syntaxin has been characterized by several biophysical methods that show that it shifts the conformational equilibrium towards the open state, but it still adopts the closed state upon Munc18 binding as observed by SAXS (Colbert et al. 2013) and single molecule FRET (Wang et al., EMBO J. 36, 816-, 2017). Moreover, the LE mutant only partially compensates for deletion of Munc13, i.e., the rescue is rather limited as observed in autaptic neuronal cultures and by single molecule FRET experiments (Lai et al., Neuron 95, 591-, 2017). Moreover, the results in Wang et al., 2017 suggest that the LE mutant interferes with proper Munc13 function (via its interactions with syntaxin), which may be related to the observed phenotypes of this mutant. Please discuss the experiments presented here in the context of these biophysical findings.

We have already discussed that STX1ALEOpen binds to Munc18-1 in closed conformation (lines 418-422). We now included the references Wang, EMBO, 2017 and Lai, Neuron, 2017 into our discussion of this point (line 420). We also included these references into our discussion of Munc18-1 and Munc13’s roles as a template for SNARE complex formation (line 444-445). Additionally, we discussed the possible impairment of Munc13-STX1 interaction when STX1 includes LEOpen mutation (lines 448-454).

https://doi.org/10.7554/eLife.69498.sa2

Article and author information

Author details

  1. Gülçin Vardar

    1. Universität Berlin, Humboldt-Universität zu Berlin, Berlin, Germany
    2. Berlin Institute of Health, Berlin, Germany
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Validation, Visualization, Writing – original draft, Writing – review and editing
    For correspondence
    gulcinv@gmail.com
    Competing interests
    None
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5295-1591
  2. Andrea Salazar-Lázaro

    1. Universität Berlin, Humboldt-Universität zu Berlin, Berlin, Germany
    2. Berlin Institute of Health, Berlin, Germany
    Contribution
    Formal analysis, Investigation
    Competing interests
    None
  3. Marisa Brockmann

    1. Universität Berlin, Humboldt-Universität zu Berlin, Berlin, Germany
    2. Berlin Institute of Health, Berlin, Germany
    Contribution
    Investigation
    Competing interests
    None
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1386-5359
  4. Marion Weber-Boyvat

    1. Universität Berlin, Humboldt-Universität zu Berlin, Berlin, Germany
    2. Berlin Institute of Health, Berlin, Germany
    Contribution
    Formal analysis, Investigation
    Competing interests
    None
  5. Sina Zobel

    1. Universität Berlin, Humboldt-Universität zu Berlin, Berlin, Germany
    2. Berlin Institute of Health, Berlin, Germany
    Contribution
    Formal analysis, Investigation
    Competing interests
    None
  6. Victor Wumbor-Apin Kumbol

    Einstein Center for Neurosciences Berlin, Berlin, Germany
    Contribution
    Formal analysis, Investigation
    Competing interests
    none
  7. Thorsten Trimbuch

    1. Universität Berlin, Humboldt-Universität zu Berlin, Berlin, Germany
    2. Berlin Institute of Health, Berlin, Germany
    Contribution
    Resources
    Competing interests
    None
  8. Christian Rosenmund

    1. Universität Berlin, Humboldt-Universität zu Berlin, Berlin, Germany
    2. Berlin Institute of Health, Berlin, Germany
    Contribution
    Conceptualization, Funding acquisition, Methodology, Project administration, Supervision, Writing – review and editing
    For correspondence
    christian.rosenmund@charite.de
    Competing interests
    None
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3905-2444

Funding

Deutsche Forschungsgemeinschaft (SFB958 TRR186)

  • Christian Rosenmund

Deutsche Forschungsgemeinschaft (Reinhart Koselleck Projects)

  • Christian Rosenmund

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

Acknowledgements

We thank the Charité Viral Core facility, Katja Pötschke, and Bettina Brokowski for virus production, Berit Söhl-Kielczynski and Heike Lerch for technical assistance, Melissa Herman and Marcial Camacho for their contribution to the manuscript, and to all the Rosenmund Lab members for the discussions. This work was supported by the German Research Council (DFG) grants 388271549, 399894546, 436260754, and 278001972.

Ethics

All procedures for animal maintenance and experiments were in accordance with the regulations of and approved by the animal welfare committee of Charité-Universitätsmedizin and the Berlin state government Agency for Health and Social Services under license number T0220/09. The generation of STX1-null mouse line was described previously (Arancillo et al. 2013, Vardar et al. 2016).

Senior Editor

  1. Gary L Westbrook, Oregon Health and Science University, United States

Reviewing Editor

  1. Axel T Brunger, Stanford University, United States

Publication history

  1. Preprint posted: February 19, 2021 (view preprint)
  2. Received: April 16, 2021
  3. Accepted: August 23, 2021
  4. Accepted Manuscript published: August 24, 2021 (version 1)
  5. Accepted Manuscript updated: August 27, 2021 (version 2)
  6. Version of Record published: September 3, 2021 (version 3)
  7. Version of Record updated: September 14, 2021 (version 4)

Copyright

© 2021, Vardar et al.

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

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