Neurons in the brain communicate with one another by passing molecules called neurotransmitters across the synapse connecting them together. Mutations in the machinery that controls neurotransmitter release can lead to epilepsy or developmental delays in early childhood, but how exactly is poorly understood.

Neurotransmitter release is primarily controlled by three proteins that join together to form the SNARE complex, and another protein called synaptotagmin-1. This assembly of proteins primes vesicles containing neurotransmitter molecules to be released from the neuron. When calcium ions bind to synaptotagmin-1, this triggers vesicles in this readily releasable pool to then fuse with the cell membrane and secrete their contents into the small gap between the communicating neurons.

Mutations associated with epilepsy and developmental delays have been found in all components of this release machinery. Here, Kádková, Murach, Østergaard et al. set out to find how three of these mutations, which are found in a protein in the SNARE complex called SNAP25, lead to aberrant neurotransmitter release.

Two of these mutations are located in the interface between the SNARE complex and synaptotagmin-1, while the other is found within the bundle of proteins that make up the SNARE complex. In vitro and ex vivo experiments in mice revealed that the two interface mutations led to defects in vesicle priming, while at the same time bypassing the control by synaptotagmin-1, resulting in vesicles spontaneously fusing with the cell membrane in an unregulated manner. These mutations therefore combine loss-of-function and gain-of-function features.

In contrast, the bundle mutation did not impact the number of vesicles in the releasable pool but reduced spontaneous and calcium ion evoked vesicle fusion. This was due to the mutation destabilizing the SNARE complex, which reduced the amount of energy available for merging vesicles to the membrane.

These findings reveal how SNAP25 mutations can have different effects on synapse activity, and how these defects disrupt the release of neurotransmitters. This experimental framework could be used to study how other synaptic mutations lead to diseases such as epilepsy. Applying this approach to human neurons and live model organisms may lead to the discovery of new therapeutic targets for epilepsy and delayed development.

The fusion machinery responsible for chemical synaptic transmission is well known: it consists of the SNARE complex, a ternary complex formed by the proteins VAMP2, syntaxin-1, and SNAP25 (Sutton et al., 1998). This complex is under tight control by upstream partner protein Munc18-1, which acts as a template for SNARE complex formation, and Munc13s, which assist in the transitions required along the pathway of assembly (Rizo, 2022). SNARE complexes assemble in a zipper-like manner; partially assembled SNAREpins bind synaptotagmin-1 (Syt1) and complexin within two separate interaction sites: the primary interface formed between a synaptotagmin-1 molecule and syntaxin-1 and SNAP25 (Zhou et al., 2015), and the tripartite interface formed by syntaxin-1 and VAMP2 with another Syt1 molecule and complexin (Zhou et al., 2017). Upon arrival of an action potential, Ca²⁺ binds to the two C2 domains of Syt1, which results in rapid vesicle–plasma membrane fusion and release of neurotransmitter within a fraction of a millisecond (Südhof, 2013).

The strong functional integration and specialization of the neuronal SNARE for speed have rendered the release machinery exquisitely susceptible to insults. De novo mutations in SNAREs and associated proteins lead to complex neurological disease, characterized by drug-resistant epilepsy, intellectual disability, movement disorders, and often autism; a syndrome which has been denoted ‘SNAREopathy’ (Verhage and Sørensen, 2020). Although rare, these are devastating conditions for the patients and their families. Consequently, there is considerable interest in revealing the molecular/cellular mechanisms for these conditions, which is seen as key to the development of treatment. Disease-causing mutations in the SNARE-machinery fall into distinct categories according to the nature of the defect (Verhage and Sørensen, 2020). These include different forms of haploinsufficiency, where the mutated protein is either lost altogether or has lost its functionality, and dominant-negative or recessive variants, as well as variants with new or changed protein interactions, referred to as neomorphs.

Most SNAREopathy mutations have been described in STXBP1 (Abramov et al., 2021a; Verhage and Sørensen, 2020; Xian et al., 2022), the gene encoding Munc18-1. These mutations are generally found to cause Munc18-1 hypo-expression, probably due to protein instability (Guiberson et al., 2018; Kovacevic et al., 2018; Martin et al., 2014; Saitsu et al., 2008), and thus the mutations belong in the haploinsufficiency category. Synaptic phenotypes of Munc18-1 hypo-expression have been identified in the Drosophila neuromuscular junction (Wu et al., 1998), in cultured mouse neurons (Toonen et al., 2006) and in brain slices of STXBP1 heterozygous mice (Chen et al., 2020; dos Santos et al., 2023), as well as in human neurons (Patzke et al., 2015), whereas expression of human missense mutations in STXBP1 null mouse neurons led to synaptic phenotypes for some mutations, but not for others (Kovacevic et al., 2018). Attempts to rescue the disease phenotypes have focused on mechanisms for increasing expression levels, for instance chemical chaperones that might prevent Munc18-1 misfolding and degradation (Abramov et al., 2021b; Guiberson et al., 2018).

The situation is different for disease mutations in Syt1 and SNAP25. In these cases, mutation generally does not cause protein instability, but instead changes function in different ways. For Syt1, disease mutations were found to cluster in the C2B domain around the top loops that coordinate Ca²⁺ (Baker et al., 2018). Expression of these mutants in neurons demonstrated reduced evoked release, in some cases an increase in spontaneous release, and a dominant-negative phenotype when co-expressed with wildtype (WT) protein (Bradberry et al., 2020). The molecular mechanism was identified as a decrease in Ca²⁺-dependent lipid binding (Bradberry et al., 2020). In SNAP25, disease-causing mutations are found within the SNARE domains (Hamdan et al., 2017; Klöckner et al., 2021; Rohena et al., 2013; Shen et al., 2014). Alten et al., 2021 studied a selection of SNAP25 mutants, and found no changes in expression levels, but changes in both spontaneous and evoked release. Specifically, mutations in the primary Syt1:SNARE interface (Zhou et al., 2015) caused an increase in spontaneous release rates, and a decrease in evoked release amplitudes. Conversely, C-terminal mutations in the so-called ‘layer residues’, whose side chains point to the center of the SNARE complex and are involved in SNAREpin zippering (Sutton et al., 1998), led to a decrease in both evoked and spontaneous release (Alten et al., 2021). This is expected because the C-terminal end of the SNARE complex is required for both types of release (Weber et al., 2010).

Alten et al., 2021 described striking phenotypes for most SNAP25 mutations tested, but the molecular reason for these phenotypes remains incompletely understood, and a few findings were surprising. For instance, Alten et al. reported that the V48F and D166Y mutants supported an unchanged readily releasable pool (RRP) of vesicles. The RRP is the pool of vesicles available for immediate release upon invasion of the presynapse by the action potential, and it is often assessed by applying a hyperosmotic solution (often 0.5 M sucrose), which causes primed vesicles to fuse (Rosenmund and Stevens, 1996). Previously, we concluded that the primary Syt1:SNARE interface is involved in vesicle priming (Schupp et al., 2016), which agrees with the suggestion that Syt1 binds to the SNAREs before Ca²⁺ arrival (Zhou et al., 2015). Conversely, Alten et al. reported a smaller RRP for the I67N mutant, but the I67 residue is present in the internal of layer +4, which we expected to affect final SNARE complex zippering causing membrane fusion rather than priming (Sørensen et al., 2006; Weber et al., 2010). These discrepancies might be explained by the fact that Alten et al. used mixed cultures and mainly studied GABAergic transmission. Glutamatergic transmission is hard to investigate in mixed cultures due to reverberating activity, but it can be studied using autaptic neurons, which was our approach (Schupp et al., 2016; Weber et al., 2010). Thus, there is a need to further study these disease mutations in glutamatergic neurons. Another open question is whether the phenotype of the mutants in the primary interface (V48F and D166Y) is explainable solely by the loss of Syt1 binding, or whether other features of these mutations add to, or detract from, the phenotype.

Here, we reexamined three SNAP25 disease mutations using glutamatergic autaptic neurons: I67N, V48F, and D166Y. We confirmed the increase in spontaneous release rate and decrease in evoked release by V48F and D166Y reported previously (Alten et al., 2021). Through in vitro analysis we find that both mutations lower Syt1 association to SNARE-protein liposomes. Additional experiments both in vitro and in cells demonstrate that V48F and especially D166Y represent partial gain-of-function mutations that increase association to partner SNAREs and lower the energy barrier for fusion, bypassing Syt1-dependent control. Thus, these mutants do not phenocopy the loss of Syt1, but combine loss of Syt1 binding with a gain-of-function phenotype. At the same time, the mutants act as loss-of-function in upstream reactions, through effects on priming. The I67N is a classical dominant-negative mutation, which increases the energy barrier for fusion, but does not change the size of the RRP if probed by a sufficiently high concentration of sucrose, nor does the I67N change the electrostatics of triggering itself. Thus, for V48F and D166Y, loss-of-function and gain-of-function features combine to change the energy landscape for vesicle priming and fusion in a complex way. These findings have consequences for our understanding of the simultaneous role of the primary SNARE:Syt1 interface in vesicle priming and release clamping. It further demonstrates the challenge faced by finding mechanism-based treatments of these disorders in the presence of multiple effects caused by single-point mutations.

We first investigated two SNAP25 disease-causing mutations within the primary Syt1:SNARE interface (V48F and D166Y; Figure 1A–C). The mutations occurred de novo and were identified in single heterozygous subjects. The V48F mutant was identified in a 15-year-old female with encephalopathy, intellectual disability and generalized epilepsy with seizures started at 5 months of age; Magnetic Resonance Imaging (MRI) was normal except for delayed myelination (Rohena et al., 2013). The D166Y mutants were found in a 23-year-old male with global developmental delay, nocturnal tonic–clonic seizures, and moderate intellectual disability; the MRI showed mild diffuse cortical atrophy (Hamdan et al., 2017). We aimed to achieve a detailed understanding of the reason for synaptic dysfunction.

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The V48F and D166Y mutants disinhibit spontaneous release and desynchronize evoked release

We constructed lentiviral vectors, which expressed WT or mutant SNAP25b fused N-terminally to EGFP as an expression marker (Delgado-Martínez et al., 2007). In the absence of SNAP25, neuronal viability is compromised (Delgado-Martínez et al., 2007; Peng et al., 2013; Santos et al., 2017; Weber et al., 2010) with low-density autaptic glutamatergic neurons dying within a few days, whereas Snap25 KO neurons growing at higher densities, or in the intact brain, can survive longer, but also eventually degenerate (Bronk et al., 2007; Hoerder-Suabedissen et al., 2019). We therefore first examined the morphology and survival of autaptic hippocampal neurons from Snap25 KO after expressing mutated or WT EGFP-SNAP25b (henceforth denoted ‘WT’ in rescue experiments; not to be confused with ‘Syt1 WT’, which refers to wildtype littermates of Syt1 KO mice).

As expected, Snap25 KO autaptic neurons did not survive in the absence of expression of exogeneous SNAP25 (Figure 2G), whereas expression of WT EGFP-SNAP25b restored survival (Delgado-Martínez et al., 2007; Ruiter et al., 2019; Weber et al., 2010). Both mutations (V48F and D166Y) caused rescue of survival; however, the number of neurons per islet was lower than in WT expressing neurons prepared in parallel. This difference was statistically significant for D166Y, but not for V48F (Figure 2G). Since patients harbor one mutated and one WT allele, we co-expressed WT EGFP-SNAP25b with mutated EGFP-SNAP25b, by combining infection with separate viruses encoding WT and mutant protein in a 1:1 ratio, as done previously by others for Syt1 (Bradberry et al., 2020). As a prerequisite for this, we demonstrated by Western blot analysis that both mutant viruses expressed similar amounts of protein as WT viruses (Figure 2A–C). In co-expressing neurons, we added half the volume of WT and mutant virus as compared to the WT condition, thus keeping the total amount of virus constant. We preferred to combine two single-cistronic viruses, rather than to construct bicistronic viruses, since the larger insert would be expected to lower the titer of viruses, which might compromise survival rescue of Snap25 KO neurons. Using co-expression of WT and mutant SNAP25b, neuronal survival was mildly and significantly reduced in the V48F + WT condition from WT (Figure 2G). Overall, mutations seemed to mildly reduce survivability of neurons. Staining of neurons against VGlut1 (a marker for glutamatergic synapses) and MAP2 (a dendritic marker) revealed no significant difference in the dendritic length or the number of synapses in neurons expressing V48F, and D166Y alone (Figure 2D–F).

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Patch-clamp was performed on days 10–14 on autaptic neurons expressing either the WT EGFP-SNAP25b, the mutants, or both the mutant and WT in a 1:1 ratio. Spontaneous miniature events were recorded at a holding voltage of −70 mV. Both primary interface mutations (V48F and D166Y) strongly increased the frequency of mEPSCs (Figure 3A, B, D, E); the mEPSC amplitude was significantly increased in the V48F, and insignificantly (p = 0.14) increased for the D166Y (Figure 3C, F). The mEPSC frequency was at least as high as in Syt1 KO neurons measured in separate experiments (Figure 3G–I). V48F and D166Y co-expressed with WT resulted in mEPSC frequencies close to the arithmetic mean between frequencies in WT and the mutant alone (Figure 3B, E), indicating that the mutations are incompletely dominant, or co-dominant.

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Brief depolarization elicits an unclamped action potential in the axon, which makes it possible to study evoked release, which is essentially absent in Snap25 KO neurons (Bronk et al., 2007; Delgado-Martínez et al., 2007). The V48F and D166Y mutants both supported evoked release, but the evoked EPSC (eEPSC) amplitude was significantly reduced in the mutant condition (Figure 4A, B, E, F), whereas when co-expressed with the WT, only the D166Y mutant had a significantly reduced eEPSC (Figure 4B, F). Integration of individual eEPSCs allowed determination of the total charge and the assessment of synchronous and asynchronous release components (Figure 4—figure supplement 1). The eEPSC charge was significantly reduced in the D166Y, but not in the V48F (Figure 4C, G), whereas in both cases the kinetics was significantly shifted in the direction of more asynchronous release, and the fast time constant was prolonged (Figure 4D, H; Figure 4—figure supplement 2). We compared the data obtained from V48F and D166Y SNAP25 to Syt1 KO neurons recorded in separate experiments. Syt1 WT neurons in this set of experiments had larger eEPSC amplitudes, and a larger charge than WT-rescued Snap25 KO neurons (Figure 4I–K), which might be caused by differences between cell cultures, or animal lines (SNAP25 vs. Syt1 lines). Note that our experiments using WT, mutant, and WT + mutant SNAP25 were always carried out in neurons prepared and recorded in parallel from the same Snap25 KO embryos (Materials and methods). In the Syt1 KO, the kinetic change was similar to the V48F and D166Y (Figure 4L), but the total charge was also strongly reduced (Figure 4K), unlike the two mutations that displayed at most a mild reduction (Figure 4C, G).

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Overall, D166Y and V48F caused a strong disinhibition of spontaneous release and a desynchronization of evoked release, as demonstrated before (Alten et al., 2021), and consistent with previous mutational studies of the primary interface, both in SNAP25 (Schupp et al., 2016) and Syt1 (Zhou et al., 2015). Accordingly, these phenotypes are similar to Syt1 KO, but the effect of V48F and D166Y on total evoked charge was milder than in the Syt1 KO. The preserved overall charge of evoked release in V48F and mild reduction in D166Y might point to a compensatory gain-of-function aspect to these mutations, in addition to the impaired Syt1 interaction.

V48F and D166Y mutations lower the energy barrier for release and reduce the RRP size

The role of Syt1 and Syt1:SNARE interactions in vesicle priming has been controversially discussed. The RRP is often assessed by applying a pulse of hypertonic solution to the neurons, usually 0.5 M sucrose (Rosenmund and Stevens, 1996; Schotten et al., 2015). However, in some experiments this did not lead to a change in RRP size in the absence of Syt1 (Bacaj et al., 2015; Xu et al., 2009), whereas experiments both from our laboratories and others showed a decrease in RRP in the absence of Syt1 (Bouazza-Arostegui et al., 2022; Chang et al., 2018; Courtney et al., 2019; Huson et al., 2020; Liu et al., 2009; Ruiter et al., 2019). The reason for this discrepancy is likely partly technical and has to do with how fast sucrose can be applied to the dendritic tree (see also Discussion). Using neurons growing on small autaptic islands makes it possible to apply sucrose to the entire dendritic tree (which is confined to the 30–50 µm island) within tens of milliseconds, which is fast enough to dissect the RRP with a short (few s) sucrose application. We set out to understand whether the V48F and D166Y mutants changed the size of the RRP.

Application of 0.5 M sucrose to neurons with the V48F or D166Y mutation resulted in estimates of the RRP (denoted RRP_0.5) that were significantly reduced compared to the WT condition (Figure 5A, C, F, H). Also the co-expressed condition displayed significantly reduced RRP_0.5. Application of two different sucrose concentrations is used to probe the size of the energy barrier for release (Basu et al., 2007; Schotten et al., 2015), because the use of a lower sucrose concentration (typically 0.25 M) will only release a fraction of the RRP, and this fraction depends sensitively on the energy barrier for release; the size of the RRP at 0.5 M sucrose (RRP_0.5) is used for normalization. Application of 0.25 M sucrose to V48F and D166Y expressing neurons strikingly led to unchanged pool release (RRP_0.25) (Figure 5B, G). Consequently, the ratio of pools (RRP_0.25/RRP_0.5) was significantly increased for both the V48F and the D166Y mutations (Figure 5D, I), indicating that these two mutations decrease the apparent energy barrier for fusion (Schotten et al., 2015). The co-expressed conditions were in-between WT and mutant and did not reach statistical significance. Consistent with these results, both mutations on average increased the release probability, calculated as the ratio between the eEPSC charge and the RRP_0.5 charge measured in the same cell. The increase was statistically significant for the D166Y mutation (Figure 5J), but not for V48F (Figure 5E). This is different from the situation in the Syt1 KO, where the RRP_0.25/RRP_0.5 and the release probability were both significantly decreased (Figure 5K–O), although in this dataset the reduction in RRP_0.5 size by removing Syt1 did not reach statistical significance (p = 0.0548, unpaired t-test). Thus, the increased mEPSC release rate from the Syt1 KO does not correlate with a reduced energy barrier as assayed by sucrose, which was reported before (Huson et al., 2020), and the V48F and D166Y have specific gain-of-function features that lower the energy barrier for vesicle fusion.

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To investigate the reasons for the change in RRP size by D166Y and V48F, we considered a one-pool model for the RRP (Figure 6A), where the RRP is filled by priming (k₁) from an upstream pool and depleted by depriming (k₋₁) or spontaneous fusion (k_f). Dividing the miniature release rates (r_mini) with the size of the RRP yields the spontaneous fusion rate k_f. The current plateau during 0.5 M sucrose application (Figure 6B) essentially reports on the priming rate (k₁), providing that the fusion rate is sufficiently increased by sucrose (Materials and methods). However, high sucrose concentrations can change the baseline current level, which will cause an error in estimation of k₁. We therefore corrected the plateau level using a plot of the variance versus mean during the sucrose application (Figure 6C); this plot is linear for the type of noise generated by synaptic transmission (shot noise). Back-extrapolation of a regression line allows a determination of the baseline current level in the presence of sucrose (Figure 6B, C; see Materials and methods). Combining RRP size with the estimates of k₁ and k_f allows determining the depriming rate, k₋₁.

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These calculations showed that k_f, the spontaneous fusion rate was strongly increased in the V48F and D166Y, and this increase was much larger for the mutations than for the Syt1 KO (Figure 6D) fulfilling D166Y > V48F > Syt1 KO. Further analysis showed that in both mutations, the forward priming rate, k₁, and the depriming rate, k₋₁, were both decreased – the latter effect was only significant for D166Y (Table 1). Summing up the effects of changes in the three parameters on the RRP size for the V48F, D166Y, and Syt1 KO (Figure 6E), we can conclude that for the V48F and even more for the D166Y, spontaneous release contributes to the reduction in RRP size, whereas this effect is minimal in the Syt1 KO. However, the major reason for the smaller RRP size is a reduction in priming rate, k₁, which is partly counteracted by the decrease in k₋₁ (which would increase RRP size). Overall, changes in priming, depriming, and spontaneous fusion rates combine to change RRP size.

Table 1

Mean ± SEM	WT	V48F	WT	D166Y	Syt1 WT	Syt1 KO
k1 [vesicles/s]	385.6 ±52.2	79.87*** ±12.5	457.4 ±77.4	37.68**** ±7.33	1227 ±189	646* ±114
k−1 [1/s]	0.0903 ±0.0091	0.0605# ±0.011	0.1114 ±0.012	0.0294**** ±0.0122	0.140 ±0.016	0.114¤ ±0.013
kf [1/s]	0.000844 ±0.000178	0.0164**** ±0.00230	0.000398 ±0.000077	0.03522**** ±0.00352	0.000235 ±0.000043	0.00286*** ±0.00062

Repetitive stimulation to determine the RRPs often results in lower estimates than sucrose application, because action potentials draw on a sub-pool of the RRP (Moulder and Mennerick, 2005), whereas sucrose releases the entire RRP (Rosenmund and Stevens, 1996). To determine the RRP sub-pool that evoked release draws on (denoted RRP_ev) we applied repetitive stimulation (50 APs @ 40 Hz) and used back-extrapolation to determine the RRP (Neher, 2015). Performing this in our standard 2 mM Ca²⁺-containing extracellular solution resulted in overall smaller estimates for RRP_ev for V48F and D166Y compared to WT (Figure 7—figure supplement 1); however, the differences were not statistically significant. The back-extrapolation method works best with high release probabilities (Neher, 2015); therefore, we repeated these experiments in the presence of 4 mM extracellular Ca²⁺. Under these conditions, the RRP_ev was significantly reduced for both the V48F and D166Y mutation (Figure 7C, G; Figure 7—figure supplement 2). Strikingly, the release probabilities (calculated as the charge of the first eEPSC of the train divided by the RRP_ev) were decreased for both mutations (Figure 7D, H), which correlated with an increased paired-pulse ratio (Figure 7A, E, inserts) (Zucker and Regehr, 2002). Thus, although the release probability calculated by normalizing evoked charge to the sucrose-determined RRP (RRP_0.5) was increased (see above), the release probability when normalizing to RRP_ev was decreased. This points to a difference in the organization of the RRP_ev and RRP_0.5 (see Discussion). The forward priming rate is determined as the slope of the linear fit used for the back-extrapolation; this parameter was reduced in both mutations (Figure 7B, F; Figure 7—figure supplement 2). Thus, the difference in priming rate extends to the RRP_ev.

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Overall, these data demonstrate a rather complex phenotype of the D166Y and V48F mutations, which combine a lowering of the energy barrier – a gain-of-function feature – with a loss of vesicle priming – a loss-of-function feature. The D166Y and V48F phenotypes can be summarized as (1) desynchronized eEPSCs with at most mildly reduced total charge, (2) lowered energy barrier for fusion, (3) increased release probability when normalized to the sucrose pools, (4) decreased RRP size due to unclamped spontaneous release and lowered forward priming rates, and (5) short-term facilitation. These phenotypes are distinctive from the Syt1 KO, which does not have a preserved charge of the eEPSC, or lowered energy barrier when probed by sucrose, or an increased release probability. Thus, the D166Y and V48F cannot be understood solely in terms of a lack of Syt1 coupling; instead, gain-of-function features are present in the mutants which are absent upon deletion of Syt1.

V48F and D166Y mutants show increased partner SNARE interactions and cause unregulated fusion in vitro

We next tried to identify the biochemical properties of V48F and D166Y, which could support a gain-of-function phenotype during exocytosis. These data are displayed in Figures 8 and 9 – data on the I67N mutant were obtained in parallel and will be presented later. To test to which degree the mutants may change the stability of SNARE complexes, full-length t-SNAREs (syntaxin-1 and SNAP25) were incubated with the cytosolic domain of VAMP2 (VAMP2cd) overnight. Cis-SNARE complex stability was tested in the presence of SDS at the indicated temperatures (Figure 8A), and the release of syntaxin-1 as a single-protein band was used as a measure of the complex dissociation. The WT, V48F, and D166Y v-/t-SNARE complexes showed a similar stability with half-maximal dissociation occurring at approximately 71°C.

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Next, we asked whether the V48F and D166Y mutants would change interaction with Syt1. To this end, Atto647-labeled giant unilamellar vesicles (GUVs) filled with isosmotic sucrose and containing preassembled t-SNARE complexes were preincubated with Atto488/Atto550-labeled small unilamellar vesicles (SUVs) containing Syt1 as well as VAMP2 for 10 min on ice, followed by centrifugation to re-isolate GUVs with attached SUVs. Fusion was blocked by performing the assay on ice. Attachment of SUVs was determined by measuring the Atto550 fluorescence of SUVs. In the absence of PI(4,5)P₂, vesicle attachment occurs by Syt1:SNARE interactions (Kim et al., 2012; Parisotto et al., 2012), probably involving the primary interface (Zhou et al., 2015). In the cell, PI(4,5)P₂ binding by Syt1 appears to happen first (Honigmann et al., 2013), whereas subsequent Syt1:SNARE binding leads to a tightly docked state (Chen et al., 2021). Under our conditions, both SNAP25 mutants (V48F and D166Y) showed significantly impaired attachment of Syt1/VAMP2 SUVs to t-SNARE GUVs (Figure 8B). Both V48 and D166 directly interact with Syt1 (Figure 1B, C, Zhou et al., 2015), so that changing these two amino acids to more bulky or hydrophobic amino acids reduced the docking from 42.7 ± 2.6% WT docking efficiency to 29.6 ± 4.1% and 20.9 ± 3.7%, respectively. In the presence of 1% PI(4,5)P₂, the vesicle attachment increased from 42.7 ± 2.6% to 66.0 ± 0.5% for WT t-SNARE. Although the Syt1:PI(4,5)P₂ interaction predominated SUV–GUV docking, both mutants still showed significantly reduced vesicle attachment by approximately 6% compared to WT (Figure 8C) (V48F: 59.1 ± 1.4% and D166Y: 60.0 ± 2.0%).

To understand how the mutations affect fusion in a well-defined reconstituted membrane fusion system, we performed in vitro lipid mixing assays using GUVs containing both t-SNAREs (syntaxin-1 and SNAP25) with 1% PI(4,5)P₂, and 0.5% Atto488/0.5% Atto550-labeled SUVs containing Syt1 and VAMP2 in the presence or absence of complexin-II (6 µM) (Kedar et al., 2015; Malsam et al., 2012). Fusion was measured at 37°C by Atto488 fluorescence dequenching, which occurs upon lipid mixing with GUVs. Calcium (100 µM free Ca²⁺ final) was added after 2 min to the t-SNARE GUV assay. Measurements were continued for another 5 min. SUVs treated with botulinum toxin D, which cleaves VAMP2 and abolishes membrane fusion, served as negative control and the corresponding background fluorescence was subtracted. Measurements were normalized to total fluorescence after detergent lysis.

Complexin-II clamps spontaneous Ca²⁺-independent membrane fusion in the reconstituted assay (i.e. fusion before addition of Ca²⁺), via laterally binding the membrane-proximal C-terminal ends of SNAP25 and VAMP2 (Malsam et al., 2020). Remarkably, V48F and D166Y showed impaired clamping by complexin, as apparent by increased fusion before Ca²⁺ addition (Figure 8D). The decreased clamping is likely caused by the reduced interaction of V48F and D166Y with Syt1. As a note, the clamping function of Syt1 becomes only obvious in the presence of complexin, whereas the clamping function of complexin depends on the presence of Syt1 (Malsam et al., 2012); thus, the clamping function of the two proteins cannot be separated. After Ca²⁺ triggering, WT SNAP25 supported the largest amount of fusion, followed by V48F, whereas D166Y Ca²⁺-dependent fusion was clearly reduced (Figure 8D). Notably, this is the same sequence as found for evoked release in synapses, considering either the amplitude or the charge of the eEPSC (Figure 4B, C, F, G). Thus, the in vitro assay reproduces both the increased spontaneous release and the reduced Ca²⁺-dependent release found in neurons, indicating that these features are present within the minimal set of fusion proteins included in this assay (i.e. the SNAREs, Syt1 and complexin).

There is evidence that SNAP25 might enter the SNARE complex last, after syntaxin-1 and VAMP2 are joined by Munc18-1 (Baker et al., 2015; Jiao et al., 2018; Sitarska et al., 2017), although another view is that a syntaxin/SNAP25 dimer bound to Munc18-1 acts as an intermediary (Jakhanwal et al., 2017). In the former case, mutations changing association of SNAP25 to the SNAREs might change exocytosis efficiency. To test this, we used a syntaxin-1 GUV assay, where the incorporation of soluble SNAP25 into the SNARE complex becomes a rate-limiting step. In this assay, fusion kinetics are much slower when compared with fusion reactions containing GUVs with preassembled t-SNAREs. Accordingly, we allowed 30 min for pre-stimulation fusion to take place, and after addition of Ca²⁺ (100 µM free Ca²⁺), fusion was followed for another 30 min (Figure 9A). The assay was performed in the presence and absence of complexin-II. D166Y and to a lesser degree V48F revealed enhanced stimulation of fusion in comparison to WT before calcium was added, regardless of the presence or absence of complexin (Figure 9B). Comparing data with and without complexin established that complexin barely suppressed spontaneous fusion in reactions containing the V48F mutant. The presence of complexin partially reduced Ca²⁺-independent fusion, but D166Y still stimulated pre-Ca²⁺ fusion compared to WT. Overall, these data demonstrate that D166Y and V48F are gain-of-function mutants under conditions where SNAP25 association to the other SNAREs is rate limiting.

To test directly whether V48F and especially D166Y enhance SNARE interactions, co-flotation assays were performed. SUVs containing either syntaxin-1 (Stx-1), or VAMP2, or Syt1, or the Syt1/VAMP2 combination were incubated with SNAP25 and re-isolated by flotation using a Nycodenz density gradient. An additional reaction, reflecting the fusion assay, contained SNAP25 in combination with both Syt1/VAMP2 SUVs and Syntaxin-1 SUVs (Figure 9D). SNAP25 recruitment for each condition was determined by SDS–PAGE followed by Coomassie Blue and silver staining. Silver stained SNAP25 bands were quantified, and the mutants were plotted relatively to the WT SNAP25 (Figure 9D and Figure 9—figure supplement 1).

SNAP25 WT did not show any binding to Syt1 SUVs (Figure 9—figure supplement 1), although direct interactions with Syt1 would be predicted based on the primary interface, indicating that such interactions are not stable under the employed conditions, which is expected because SNAP25 is unstructured until it binds its SNARE partners (Fasshauer et al., 1997). D166Y showed profoundly increased interactions with SUVs containing either syntaxin-1, or VAMP2, or Syt1/VAMP2, and the combination used in the fusion assay (Figure 9D; Figure 9—figure supplement 1). V48F displayed mildly increased binding to syntaxin-1 and Syt1/VAMP1 SUVs, and a tendency to increased association to VAMP2 SUVs (p = 0.0629, one-sample t-test). These data show that loss of Syt1 interaction upon mutation in the primary interface can be accompanied by a gain-of-function phenotype stimulating interactions with the other SNARE partners. This association of SNAP25 to the other SNAREs might happen as one of the last steps toward fusion; consequently, when D166Y and V48F join the complex prematurely, it will bypass layers of control and result in uncontrolled fusion.

The I67N mutation supports an intact RRP, but an increased energy barrier for fusion

We next addressed the phenotype of the I67N disease mutation, which was found in an 11-year-old female, who suffered from myasthenia, cortical hyperexcitability, ataxia, and intellectual disability, but with normal brain MRI (Shen et al., 2014). The I67 is found within the interaction layer +4 (Fasshauer et al., 1998), which helps in assembly of the C-terminal of the SNARE complex (Gao et al., 2012), and it might therefore have a different synaptic phenotype than V48F and D166Y.

In in vitro experiments, SNARE complexes formed with the I67N mutant displayed a lower stability, with the melting temperature reduced from 71°C to approximately 56°C (Figure 8A), as expected for a mutation that destabilizes the SNARE complex. I67N also showed a strong decrease in SUV docking (Figure 8B, C), which is likely caused by the destabilization of the t-SNARE complex, which indirectly perturbs the Syt1-binding interface(s). In lipid mixing assays, I67N strongly reduced both Ca²⁺-independent and Ca²⁺-dependent fusion, whether the t-SNAREs were preassembled (Figure 8D, E) or not (Figure 9B, C). The co-flotation assay did not display any binding of I67N to t-SNAREs or Syt1 (Figure 9D). The binding to Syt1/VAMP2 SUVs was reduced compared to SNAP25 WT, but since binding of the WT protein is already very low, the biological significance of this result is unclear. Overall, these data indicate that I67N is inferior in membrane fusion.

Lentiviruses encoding the I67N mutant N-terminally fused to EGFP expressed similar amounts as WT EGFP-SNAP25b (Figure 10A–C). Expression in SNAP25 KO neurons resulted in reduced rescue of survival in neurons expressing I67N alone, whereas neurons co-expressing WT and I67N had intermediate survival, not significantly different from WT (Figure 10D). Staining against MAP2 (dendritic marker) and VGlut1 (synapse marker) showed that the number of synapses on average was reduced in the I67N (Figure 10E, F), and the dendritic length was on average reduced (Figure 10G), but the changes did not reach statistical significance. Patch-clamp measurements demonstrated strongly reduced spontaneous release frequencies (Figure 10H–J) and evoked release amplitude (Figure 10K, L) with the I67N mutation, see also Alten et al., 2021. mEPSCs were absent in most I67N expressing neurons, whereas WT and I67N co-expressing neurons had a very low mEPSC rate, much closer to the I67N than the WT phenotype; similar for eEPSC amplitudes and charges (Figure 10L-M). The I67N therefore is dominant negative for both types of release, in contrast to the incompletely dominant phenotypes of the V48F and D166Y mutants (see above). The fraction of synchronous release was unchanged in WT and I67N co-expressing neuron (Figure 10N); this number could not be estimated for I67N expressing neurons due to the low amount of release.

Figure 10

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Reduced spontaneous and evoked release could result from a decrease in priming, or fusion, or both. To distinguish between these possibilities, we turned to sucrose applications. Application of 0.25 M sucrose did not lead to any measurable release in I67N expressing cells, and only minimal release in cells co-expressing WT and I67N (Figure 11B). This indicates that the energy barrier is increased in amplitude, and therefore the RRP might be underestimated when probed by 0.5 M sucrose (Schotten et al., 2015). Indeed, 0.5 M sucrose displayed reduced release in the I67N and WT + I67N condition (Figure 11C), but this could be due to defects in priming or fusion. To investigate this, we applied a stronger stimulus, 0.75 M sucrose, to these cells (Schotten et al., 2015). Strikingly, the RRP as assessed by 0.75 M sucrose (RRP_0.75) was unchanged between WT, I67N, and WT + I67N co-expressed cells (Figure 11E, G). Application of 0.375 M sucrose led to small amounts of release in the I67N, but more in the WT + I67N co-expressed situation (Figure 11F). Forming the ratio RRP_0.375/RRP_0.75 revealed a statistically significant reduction in I67N compared to WT (Figure 11H). Therefore, the RRP per se appears intact in the I67N (when probed by sufficiently high concentrations of sucrose), but the vesicles face a higher apparent fusion barrier, which makes the RRP appear smaller if assessed by 0.5 M sucrose. The higher apparent fusion barrier explains the lower frequency of mEPSC in the I67N mutation, the lower degree of spontaneous fusion in in vitro assays, as well as the lower amount of Ca²⁺-dependent release in vitro and in the cell.

Figure 11

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The I67N mutation profoundly affected trains in 2 mM extracellular Ca²⁺ (Figure 11I) leading to strong facilitation, which is expected due to the strong phenotype of this mutation, which radically lowers release probability. Even when co-expressed with WT protein, the train facilitated over the first several stimulations, attesting to the strong dominant-negative feature of the I67N mutation (Figure 11I–J). Consequently, the paired-pulse ratio was increased in the I67N and intermediate in the WT + I67N condition (Figure 11K). Back-extrapolation of these trains was not reliable, because the low release probability in the I67N mutation made it impossible to achieve sufficient depletion of the RRP.

The energy barrier for fusion is exquisitely sensitive to the charges on the surface of the SNARE complex, with positive charges decreasing and negative charges increasing the fusion barrier amplitude (Ruiter et al., 2019). To investigate whether the same electrostatic mechanism applies to the I67N, we combined the I67N mutation with a mutation of four amino acids (‘4K’ = SNAP25 E183K/S187K/T190K/E194K) in the second SNARE domain of SNAP25, constructing the quintuple mutation (‘I67N/4K’ = SNAP25 I67N/E183K/S187K/T190K/E194K). The 4K mutation lowers the energy barrier for fusion by increasing the charge of the SNARE complex surface by +6 via charge introduction and charge reversal (Ruiter et al., 2019). The 4K mutation increased the mEPSC release rate compared to WT (Ruiter et al., 2019), whereas in the combined I67N/4K mutation the spontaneous release rate was indistinguishable from WT (Figure 12A–C), showing that increased positive charges rescued the defect of spontaneous release in the I67N mutant. Evoked release was also increased in the 4K mutation compared to WT (Ruiter et al., 2019), but in the I67N/4K mutation, evoked release was still strongly depressed compared to WT (Figure 12D, E). Nevertheless, evoked release was noticeable in the combined mutation, whereas it was almost absent in I67N expressing cells (comp. Figure 10K, L), indicating a positive effect of the 4K mutation, which amounted to an increase by a factor ~5 (eEPSC amplitude, I67N: 0.0475 ± 0.0087; I67N/4K: 0.2668 ± 0.12 nA; Mann–Whitney test, p = 0.035).

Figure 12

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Figure 12—source data 1

Excel file containing quantitative data.

https://cdn.elifesciences.org/articles/88619/elife-88619-fig12-data1-v2.zip

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We previously created a simple mathematical model that links the release rate to the number of charges added to the SNARE complex (Ruiter et al., 2019). This model includes both spontaneous and evoked release, which are separated by the addition of 35 positive charges in the latter case (Figure 12F, black points are WT spontaneous and evoked release rates; blue line is the model fitted to WT data). Placing the spontaneous release rates for I67N and I67N/4K on this curve (by dividing the spontaneous release rate with RRP size and finding a corresponding charge value using the model) resulted in two points (red) separated by 5.6 charges, which is close to the nominal 6 charges added by the 4K mutation (Figure 12F). Similarly, evoked release in the I67N and the I67N/4K were separated by 5.9 charges (Figure 12F). The fact that these numbers are close to the nominal 6 charges introduced shows that even for the I67N mutation, the same basal electrostatic model still applies, but the deleterious effect of the I67N on evoked release rates is larger than on spontaneous rates and therefore the rescue of evoked release rate by positive charges is insufficient to reach WT values. Moreover, because of the saturating form of the curve (i.e. the model) adding positive charges is an effective way of rescuing spontaneous, but not evoked release.

Overall, the I67N disease mutation increases the amplitude of the energy barrier for fusion, and it does so more for evoked than for spontaneous release, but the electrostatic mechanism, which we assume is part of release triggering (Ruiter et al., 2019), appears to be intact.

We have shown that two SNAP25 mutations (V48F and D166Y) that compromise interaction with Syt1 lead to complex phenotypes characterized by a combination of loss-of-function and gain-of-function features. Thereby, the mutations fall into the ‘neomorph’ category, where the mutated protein has novel or changed interactions or functions (Verhage and Sørensen, 2020). In contrast, the I67N substitution within the SNARE bundle is a dominant-negative mutation.

SNAP25 disease mutations change protein–protein interactions and the energy landscape of fusion

Both the V48F and the D166Y resulted in a decrease in the amplitude of the apparent energy barrier for fusion, whereas the I67N increased the amplitude of the apparent fusion barrier. A vesicle’s release willingness can only be assessed by fusing it; therefore, it is not possible to distinguish between effects on the fusion barrier per se, and effects on the fusion machinery. In recognition of this fact, we here refer to the ‘apparent energy barrier’. Using Arrhenius’ equation to convert fusion, priming and depriming rates to their respective energy barrier heights (see Materials and methods for the assumptions behind this procedure), we can derive the energy landscape for fusion of the three mutants (Figure 13A–D). This shows the multiple changes in the V48F and D166Y, which affect at least two different barriers (priming and fusion, Figure 13A, B), leading to a complex phenotype, whereas for I67N the fusion barrier is primarily (or solely) affected (Figure 13D).

Figure 13

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When combining the I67N with the 4K mutation, which introduces six extra positive fixed charges, we could place our data within the framework of our model for electrostatic triggering (Ruiter et al., 2019) and show that the effect of charge per se is approximately the same in the I67N mutant as in the WT. Note that there are endogenous positively charged amino acids toward the C-terminal end of SNAP25 that are important for release rates (Fang et al., 2015). Rescue of spontaneous release was completed by adding six positive charges, which is consistent with the idea that the assembly of the C-terminal end of the SNARE complex, which is compromised by I67N (Rebane et al., 2018), works against the electrostatic energy barrier, which is affected by the SNARE surface charge (Ruiter et al., 2019). In contrast, rescue of evoked release by charges were incomplete, due to the larger effect of I67N on evoked release, combined with the shallow effect of charges on evoked release (Ruiter et al., 2019). The larger susceptibility of evoked release to C-terminal mutation of the SNAREs might be partly due to the higher number of SNARE complexes involved in evoked than in sustained/spontaneous release (Mohrmann et al., 2010).

The effect of the I67N mutation in the energy domain at rest can be calculated from the spontaneous release rate, which was reduced by a factor 22.4 (from 1.31 ± 0.36 to 0.0583 ± 0.0274 Hz). This corresponds to an effect in the energy domain of 3.1 k_BT (where k_B is Boltzmann’s constant; assuming unchanged RRP size, pool normalization is not required). Work with single-molecular optical tweezers showed that the I67N mutation destabilizes the overall SNARE C-terminal and linker domain, which are supposed to deliver the power stroke for membrane fusion, by 14 k_BT (Rebane et al., 2018), which is substantially more. The reduction in spontaneous release rate is more comparable to the reduction in the transition rate of folding by the C-terminal and linker domain by a factor of ~10 (Rebane et al., 2018). Since at least three SNARE complexes, possibly more, contribute to vesicle fusion (Bao et al., 2018; Manca et al., 2019; Mohrmann et al., 2010; Shi et al., 2012), folding kinetics correlates better to spontaneous fusion rates than overall SNARE complex stability.

The D166Y and V48F mutations lead to increases in spontaneous release, and more asynchronous eEPSCs, consistent with their localization in the primary SNARE:Syt1 interface (Schupp et al., 2016; Zhou et al., 2015; Zhou et al., 2017), and the demonstrated impaired Syt1 binding (Figure 8B, C), see also Alten et al., 2021. These phenotypes are at first glance similar to Syt1 knockout/knockdown (Bouazza-Arostegui et al., 2022; Chang et al., 2018; Huson et al., 2020; Ruiter et al., 2019). However, when normalized to the RRP size, mEPSC frequencies were much higher in the V48F and the D166Y than in the Syt1 KO (Figure 6D), and dual sucrose applications indicated a decrease in the amplitude of the sucrose-probed apparent energy barrier and increased release probability, features not found in the Syt1 KO (Bouazza-Arostegui et al., 2022; Huson et al., 2020). This indicates a gain-of-function feature of these mutations, which fulfills D166Y > V48F; such a feature was identified as an increased interaction of V48F and D166Y with VAMP2- and syntaxin-1-containing SUVs (Figure 9D); the interaction was stronger for D166Y than for V48F. Molecular dynamics (MD) simulations of the SNAP25 mutants did not reveal major structural changes in the SNAP25 backbone compared to WT (Figure 9—figure supplement 2A–C). Nevertheless, the calculation of electrostatic (Coulomb) and van der Waals (Lennard–Jonson, LJ) interactions for residues 48–52 (Figure 9—figure supplement 2D) and residues 162–166 (Figure 9—figure supplement 2E) shows that for D166Y the interaction energy is substantially more negative. This suggests that the interaction between D166Y and nearby residue H162 is stronger than in WT. Overall, this may result in stabilization of a structure consistent with SNARE complex formation and explain why D166Y is a stronger gain-of-function mutation than V48F. However, note that the AlphaFold prediction (Jumper et al., 2021; Mirdita et al., 2022) used as a starting point for these simulations is identical to the structure of SNAP25 in the assembled SNARE complex (Zhou et al., 2015), whereas unassembled SNAP25 is likely less structured or unstructured (Fasshauer et al., 1997). Overall, these disease mutants do not only fail in their interaction with Syt1, they bypass fusion control, resulting in premature SNARE complex assembly. Although the increased spontaneous release rate might cause patient symptoms, Alten et al., 2021 suggested that the resulting postsynaptic depolarization might compensate for the smaller eEPSC amplitude to normalize the overall firing rate.

In an SUV:GUV fusion assay, where folding of SNAP25 onto syntaxin-1/VAMP2 is rate limiting, D166Y and V48F caused an increase in pre-stimulation fusion rates. This is consistent with recent data showing that folding of SNAP25 onto a template formed by VAMP2 and syntaxin-1 held in place by Munc18-1 might be a late, rate-limiting, step in exocytosis (Jiao et al., 2018). This process is regulated and sped up by Munc13-1 (Kalyana Sundaram et al., 2021; Shu et al., 2020; Wang et al., 2019). The fact that a similar effect on spontaneous fusion was seen in the assay with preformed t-SNAREs in the presence of complexin indicates that there is an assembly step, even in that assay, which can be sped up by the mutations. This aligns with the demonstration by single-molecule FRET of a further assembled (tighter) state of the trans-SNARE complex induced by Ca²⁺-unbound Syt1, which becomes committed for fusion once Ca²⁺ binds (Das et al., 2020).

All data generated and used during this study are included in the manuscript and supporting files. Source data are included for all figures. All structures generated using AlphaFold2, as well as the initial structures and topology files for atomistic molecular dynamics simulations, were deposited to ZENODO (https://doi.org/10.5281/zenodo.10051665).

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Author details

1. Anna Kádková

   Department of Neuroscience, University of Copenhagen, Copenhagen, Denmark

   Contribution

   Data curation, Formal analysis, Investigation, Writing – original draft, Writing – review and editing

   Contributed equally with

   Jacqueline Murach and Maiken Østergaard

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-6648-9679

2. Jacqueline Murach

   Heidelberg University Biochemistry Center, Heidelberg, Germany

   Contribution

   Data curation, Formal analysis, Investigation, Writing – review and editing

   Contributed equally with

   Anna Kádková and Maiken Østergaard

   Competing interests

   No competing interests declared

3. Maiken Østergaard

   Department of Neuroscience, University of Copenhagen, Copenhagen, Denmark

   Contribution

   Data curation, Formal analysis, Investigation, Writing – original draft, Writing – review and editing

   Contributed equally with

   Anna Kádková and Jacqueline Murach

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0009-0005-3869-9069

4. Andrea Malsam

   Heidelberg University Biochemistry Center, Heidelberg, Germany

   Contribution

   Formal analysis, Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

5. Jörg Malsam

   Heidelberg University Biochemistry Center, Heidelberg, Germany

   Contribution

   Formal analysis, Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

6. Fabio Lolicato

   1. Heidelberg University Biochemistry Center, Heidelberg, Germany
   2. Department of Physics, University of Helsinki, Helsinki, Finland

   Contribution

   Formal analysis, Investigation, Visualization, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-7537-0549

7. Walter Nickel

   Heidelberg University Biochemistry Center, Heidelberg, Germany

   Contribution

   Supervision, Funding acquisition, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-6496-8286

8. Thomas H Söllner

   Heidelberg University Biochemistry Center, Heidelberg, Germany

   Contribution

   Conceptualization, Supervision, Funding acquisition, Writing – review and editing

   Competing interests

   No competing interests declared

9. Jakob Balslev Sørensen

   Department of Neuroscience, University of Copenhagen, Copenhagen, Denmark

   Contribution

   Conceptualization, Supervision, Funding acquisition, Writing – original draft, Project administration, Writing – review and editing

   For correspondence

   jakobbs@sund.ku.dk

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5465-3769

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