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eLife. 2020; 9: e54506.
Published online 2020 May 13. doi: 10.7554/eLife.54506
PMCID: PMC7220375
PMID: 32401194

Synergistic roles of Synaptotagmin-1 and complexin in calcium-regulated neuronal exocytosis

Sathish Ramakrishnan,1 Manindra Bera,1 Jeff Coleman,1 James E Rothman,1 and Shyam S Krishnakumar1,2
Axel T Brunger, Reviewing Editor and Vivek Malhotra, Senior Editor
Axel T Brunger, Stanford University, United States;
Author information Article notes Copyright and License information PMC Disclaimer

Associated Data

Supplementary Materials
Data Availability Statement

Abstract

Calcium (Ca2+)-evoked release of neurotransmitters from synaptic vesicles requires mechanisms both to prevent un-initiated fusion of vesicles (clamping) and to trigger fusion following Ca2+-influx. The principal components involved in these processes are the vesicular fusion machinery (SNARE proteins) and the regulatory proteins, Synaptotagmin-1 and Complexin. Here, we use a reconstituted single-vesicle fusion assay under physiologically-relevant conditions to delineate a novel mechanism by which Synaptotagmin-1 and Complexin act synergistically to establish Ca2+-regulated fusion. We find that under each vesicle, Synaptotagmin-1 oligomers bind and clamp a limited number of ‘central’ SNARE complexes via the primary interface and introduce a kinetic delay in vesicle fusion mediated by the excess of free SNAREpins. This in turn enables Complexin to arrest the remaining free ‘peripheral’ SNAREpins to produce a stably clamped vesicle. Activation of the central SNAREpins associated with Synaptotagmin-1 by Ca2+ is sufficient to trigger rapid (<100 msec) and synchronous fusion of the docked vesicles.

Research organism: E. coli

Introduction

The controlled yet rapid (sub-millisecond) release of neurotransmitters stored in synaptic vesicles (SVs) is central to all information processing in the brain (Südhof, 2013; Kaeser and Regehr, 2014; Rizo, 2018). Synaptic release of neurotransmitters relies on the efficient coupling of SV fusion to the triggering signal – action potential (AP)-evoked Ca2+ influx into the pre-synaptic terminal (Kaeser and Regehr, 2014; Südhof, 2013). SV fusion is catalyzed by synaptic SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) proteins, VAMP2 on the vesicles (v-SNAREs) and Syntaxin/SNAP25 (t-SNAREs) on the pre-synaptic membrane (Söllner et al., 1993; Weber et al., 1998). On their own, SNARE proteins are constitutively active and intrinsically trigger fusion in the range of 0.5–1 s (Ramakrishnan et al., 2018; Ramakrishnan et al., 2019; Xu et al., 2016).

To achieve the requisite speed and precision of synaptic transmission, the nerve terminals maintain a pool of docked vesicles that can be readily released upon Ca2+ influx (Südhof, 2013; Kaeser and Regehr, 2014; Südhof and Rothman, 2009). The prevailing theory is that in a ‘release-ready’ vesicle, multiple SNARE complexes are firmly held (‘clamped’) in a partially assembled state. These ‘SNAREpins’ are then synchronously released by Ca2+ to drive fusion dramatically faster than any one SNARE alone (Rothman et al., 2017; Brunger et al., 2018; Volynski and Krishnakumar, 2018). Several lines of evidence suggest that two synaptic proteins, Synaptotagmin-1 (Syt1) and Complexin (Cpx) play a critical role in establishing Ca2+-regulated neurotransmitter release (Geppert et al., 1994; Xu et al., 2007; Bacaj et al., 2013; Yang et al., 2013; Huntwork and Littleton, 2007).

Synaptotagmin-1 is a SV-localized protein with a large cytoplasmic part containing tandem C2A and C2B domains that bind Ca2+ (Fuson et al., 2007; Sutton et al., 1995). It is well established that fast AP-evoked synchronous release is triggered by Ca2+ binding to Syt1 C2 domains (Brose et al., 1992; Geppert et al., 1994; Littleton et al., 1993). Genetic analysis shows that Syt1 also plays a crucial role in ‘clamping’ spontaneous release and Ca2+-evoked asynchronous release to ensure high fidelity of the Ca2+-coupled synchronous transmitter release (Xu et al., 2009; Bacaj et al., 2013; Littleton et al., 1993). Recently it has been demonstrated that C2B-driven self-oligomerization of Syt1 provides the structural basis for its clamping function (Wang et al., 2014; Bello et al., 2018; Tagliatti et al., 2019). Syt1 is also involved in initial stages of docking of SVs to the plasma membrane (PM), in part mediated by its interaction with the anionic lipids, phosphatidylserine (PS) and phosphatidylinositol 4, 5-bisphosphate (PIP2) on the PM (Honigmann et al., 2013; Parisotto et al., 2012; Park et al., 2012). Besides the membrane interaction, Syt1 also binds the neuronal t-SNAREs on the PM, both independently (primary binding site) and in conjunction with Complexin (tripartite binding site) (Zhou et al., 2015; Zhou et al., 2017; Grushin et al., 2019). The Syt1-SNARE interactions are important for Syt1 role in SV docking, clamping fusion and triggering Ca2+-dependent neurotransmitter release (Zhou et al., 2015; Zhou et al., 2017).

Complexin is a cytosolic α-helical protein that binds and regulates SNARE assembly (Chen et al., 2002; Kümmel et al., 2011; Li et al., 2011). Biochemical analyses reveal that Cpx catalyzes the initial stages of SNARE assembly, but then blocks complete assembly (Giraudo et al., 2009; Kümmel et al., 2011; Li et al., 2011). Physiological studies show that Cpx indeed clamps spontaneous fusion in invertebrate neurons (Huntwork and Littleton, 2007; Wragg et al., 2013; Cho et al., 2014), but its importance as a fusion clamp in mammalian neurons is still under debate (Yang et al., 2013; López-Murcia et al., 2019; Courtney et al., 2019). However, under all conditions, Cpx has been shown to facilitate vesicle priming and promote AP-evoked synchronous release (Yang et al., 2013; Yang et al., 2015; López-Murcia et al., 2019).

It is broadly accepted that Syt1 and Cpx are both involved in clamping spurious or delayed fusion events and synchronize neurotransmitter release to Ca2+-influx. However, there is presently no coherent understanding how these proteins assemble and operate together. To gain mechanistic insights into this process, a reductionist approach where the variables are limited, and components can be rigorously controlled is required. We recently described a biochemically-defined fusion system based on a pore-spanning lipid bilayer setup that is ideally suited for this purpose (Ramakrishnan et al., 2018). This reconstituted setup is capable of precisely tracking individual vesicle docking, clamping (delay from docking to spontaneous fusion) and Ca2+ triggered release at tens of milliseconds timescale (Coleman et al., 2018; Ramakrishnan et al., 2019). Critically, this setup allows us to examine these discrete sub-steps in the vesicular exocytosis process, independent of alterations in the preceding or following stages (Coleman et al., 2018; Ramakrishnan et al., 2019; Ramakrishnan et al., 2018).

Using the in vitro fusion system, we recently reported that under artificially low-VAMP2 conditions (~13 copies of VAMP2 and ~25 copies of Syt1 per vesicle), Syt1 oligomerization is both necessary and sufficient to establish a Ca2+-sensitive fusion clamp (Ramakrishnan et al., 2019). Here we extend this study to more physiologically-relevant conditions, using SV mimics reconstituted with ~25 copies of Syt1 and ~70 copies of VAMP2. We report that under these conditions, both Cpx and Syt1 oligomers are needed to stably clamp all SNARE complexes and the reversal of the Syt1 clamp is sufficient to achieve fast, Ca2+-triggered synchronized fusion.

Results

Synaptotagmin and complexin co-operate to clamp vesicle fusion

With the goal of approximating the physiological context, we chose a reconstitution condition for small unilamellar vesicles (SUV) resulting in an average of 74 copies and 25 copies of outward-facing VAMP2 and Syt1, respectively (Figure 1—figure supplement 1). We employed pre-formed t-SNAREs (1:1 complex of Syntaxin1 and SNAP-25) in the planar bilayers (containing 15% PS and 3% PIP2) to both simplify the experimental approach and to bypass the requirement of SNARE-assembling chaperones, Munc18 and Munc13 (Baker and Hughson, 2016; Rizo, 2018). In most of the experiments, we used fluorescently-labelled lipid (2% ATTO647-PE) included in the SUVs to track the docking, diffusion and fusion of individual SUVs (Figure 1A).

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Syt1 and Cpx co-operatively clamp SNARE-mediated vesicle fusion under physiologically-relevant conditions.

The effect of Syt1 and Cpx on SNARE-driven fusion was assessed using a single-vesicle docking and fusion analysis with a pore-spanning bilayer setup (Ramakrishnan et al., 2019; Ramakrishnan et al., 2018). (A) Representative fluorescence (ATTO647N-PE) traces showing the behavior of small unilamellar vesicles (SUV) containing VAMP2 (vSUV) or Syt1 and VAMP2 (Syt1-vSUV) on t-SNARE containing bilayer in the presence or the absence of Cpx. (B) The time between docking and fusion was measured for each docked vesicle and the results for the whole population are presented as a survival curve. vSUVs (black curve) are diffusively mobile upon docking (Figure 1—figure supplement 2) and fuse spontaneous with a half-time of ~1 s. Addition of soluble Cpx (2 µM) does not change this behavior (green curve). Inclusion of Syt1 in the v-SUV (red curve) does not block fusion but increases the time from docking-to-fusion (~5 s half-life), in effect delaying the kinetics of fusion. When included together Syt1 and Cpx (blue curves) fully arrest fusion to produce stably docked SUVs that attach and remain in place during the entire period of observation. (C) Syt1 and Cpx, both individually and collectively increase the number of docked vesicles. In all cases, a mutant form of VAMP2 (VAMP24X) which eliminated fusion was used to unambiguously estimate the number of docked vesicles after the 10 min interaction phase. The average values and standard deviations from three to four independent experiments are shown for each condition. In sum, 500–1000 vesicles were analyzed for each condition.

Figure 1—figure supplement 1.

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Coomaisse-stained SDS-PAGE analysis of the proteins used in this study.

(A) Gel image of the VAMP2 ±Syt1 vesicles. Based on the densitometry analysis, we estimated each vesicle contain 74 ± 4 and 26 ± 6 copies of outward-facing VAMP2 and Syt1 respectively under ‘normal’ physiologically-relevant condition and 13 ± 2 and 26 ± 6 copies of VAMP2 and Syt1 under low-copy conditions. (B) In all fusion experiments, co-purified t-SNAREs containing 1:1 complex of Syntaxin1a and SNAP25 was reconstituted into the free-standing bilayer and full-length Complexin 1 (2 µM final concentration) was added in solution.

Figure 1—figure supplement 2.

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Representative time-lapse fluorescence (ATTO647N-PE) images showing the behavior of VAMP2- containing SUVs in presence of Syt1 and/or Cpx.

Representative time-lapse fluorescence (ATTO67N-PE) images of a docked SUVs showing that all VAMP2-containg SUVs (vSUVs) are diffusively mobile upon docking and fuse spontaneous within 1 s. This behavior is un-altered upon addition of 2 µM Cpx (vSUV + Cpx). Inclusion of Syt1 in the v-SUVs (Syt1-vSUV) introduces a significant docking-to-fusion delay, but most vesicles proceed to fuse spontaneously. In contrast, Cpx, when added along with Syt1 (Syt1-vSUV + Cpx), produces a stable clamp and the vesicles are docked in-place and largely immobile and do not fuse during the observation period. Images corresponding to a single 5 µm suspended bilayer is shown.

Figure 1—figure supplement 3.

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Survival analysis showing that vSUVs in the presence of Syt1 and Cpx dock and remain stably clamped for up to 1 hr*.

The vesicle fluorescence (ATTO647N-PE) were continuously imaged for the first 60 s post-docking with 150 ms frame rate (A), beyond which the vesicles were imaged only every 10 min for up to 1 hr (B) to minimize photo-bleaching. Average and standard deviation from three independent experiments are shown. (C) Representative fluorescence (ATTO647N-PE) image (bottom row) showing that Syt1/Cpx clamped vesicles remain Ca2+-sensitive even 1 hr post-docking. At each time point, we also recorded the NBD-fluorescence (top row) to verify the stability of the bilayer. Images corresponding to a single 5 µm suspended bilayer is shown. *Note: 1 hr is the maximum time-point we have tested so far considering the stability of the suspended bilayer.

We initially focused on the kinetics of constitutive fusion to assess the ability of Syt1 and Cpx to ‘clamp’ SNARE-driven fusion in the absence of Ca2+. We monitored large ensembles of vesicles to determine the percent remaining unfused as a function of time elapsed after docking and quantified as ‘survival percentages’ (Figure 1B). Docked immobile vesicles that remained un-fused during the initial 10 min observation period were defined as ‘clamped’ (Ramakrishnan et al., 2019). Vesicles containing VAMP2 only (vSUV) that docked to the t-SNARE containing bilayer surface were mobile and a majority (>95%) spontaneously fused typically with a t1/2 ~ 1 sec post-docking (Figure 1B, Figure 1—figure supplement 2, Table 1, Table 2, Video 1).

Table 1.

Survival probabilities at specific points in time post-docking (Kaplan Meier estimators) for the VAMP2-containing vesicles (vSUV) in the presence of Cpx, Syt1 or both. The corresponding survival curves are shown in Figure 1B
Time (s)
(post-docking)		vSUVvSUV + CpxSyt1-vSUVSyt1-vSUV + Cpx
0.5880.79780.49450.99151.0000
1.0290.41340.22770.95190.9989
2.0580.09410.05440.80450.9963
3.0870.05830.03190.63610.9936
4.1160.04910.02670.51100.9917
5.1450.04390.02430.39840.9888
7.4970.04210.02150.24820.9821
9.9960.03880.02110.18530.9814
15.1410.03810.02090.15470.9794
20.1390.03790.02010.15050.9682

Table 2.

The survival curves in Figure 1B were compared pair-wise using the log-rank test to determine the statistical significance (p-values) of the observed effects of Syt1 and Cpx on v-SUV fusion
vSUVvSUV + cpxSyt1-vSUVSyt1-vSUV + cpx
vSUVn/ap=0.731p<0.001p<0.0001
vSUV + Cpxp=0.731n/ap<0.0001p<0.0001
Syt1-vSUVp<0.001p<0.001n/ap<0.001
Syt1-vSUV + Cpxp<0.0001p<0.0001p<0.001n/a

Video 1.

Download video file.(169K, mp4)
Representative video showing the docking, diffusion and spontaneous fusion of a vSUV to free-standing t-SNARE containing lipid bilayer containing t-SNARE complex.

The movie was acquired using a Leica confocal scanning microscope at a speed of 7 frames/sec and the ATTO647N-PE fluorescence was used to track the fate of the vesicle. For clarity, a single hole corresponding to 5 µm suspended bilayer marked by the white circle is shown.

Inclusion of wild-type Syt1 in the vesicles (Syt1-vSUVs) enhanced the vesicle docking rate, with ~8 fold increase in total number of docked vesicles (Figure 1C). The majority (~80%) of docked Syt1-vSUVs remained mobile on the bilayer surface and fused on an average ~5–6 s after docking (Figure 1B, Figure 1—figure supplement 2, Table 1, Table 2, Video 2). The remaining small fraction (~20%) were immobile and stably clamped. This is in stark contrast to our earlier finding under low-copy VAMP2 conditions wherein the bulk of the Syt1-vSUVs (>90%) were stably clamped (Ramakrishnan et al., 2019). The pronounced docking-to-fusion delay introduced by Syt1 (t1/2 ~5 sec for Syt1-vSUV compared to ~1 s for vSUV) suggests that under physiologically-relevant (‘normal’ VAMP copy number) conditions, Syt1 alone can meaningfully delay but not stably clamp fusion.

Video 2.

Download video file.(428K, mp4)
Representative video showing the docking, slow diffusion and delayed spontaneous fusion of a Syt1- containing vSUV to free-standing lipid bilayer containing t-SNAREs.

The movie was acquired using a Leica confocal scanning microscope at a speed of 7 frames/sec and the ATTO647N-PE fluorescence was used to track the vesicle. For clarity, a single hole corresponding to 5 µm suspended bilayer marked by the white circle is shown.

This unstable Syt1 clamp was stabilized by addition of Cpx (Figure 1B, Figure 1—figure supplement 2, Video 3). In the presence of 2 µM of soluble Cpx, all Syt1-vSUVs were immobile following docking (Figure 1B), and they rarely fused over the initial observation period (Figure 1B, Figure 1—figure supplement 2, Table 1, Table 2, Video 3). In fact, these vesicles remained stably docked up to 1 hr without fusing (Figure 1—figure supplement 3). Furthermore, Syt1 and Cpx together significantly increased (~18 fold) the total number of docked vesicles (Figure 1C). Thus, we find that Syt1 and Cpx act synergistically to increase the rate of vesicle docking and serve to effectively block fusion of the docked vesicles under resting conditions.

Video 3.

Download video file.(737K, mp4)
Representative video showing the immobile docking of a Syt1-vSUV to a free-standing t-SNARE containing lipid bilayer in the presence of 2 µM Cpx.

The ATTO647N-DOPE introduced in the SUV was used to track the vesicle and the movie was acquired using a Leica confocal scanning microscope at a speed of 7 frames/sec. For clarity, a single hole corresponding to 5 µm suspended bilayer marked by the white circle is shown.

Addition of soluble Cpx (2 µM) alone produced a ~10 fold increase in the number of docked vesicles (Figure 1C) but did not change the behavior of the docked vSUVs (Figure 1B, Figure 1—figure supplement 2, Table 1, Table 2, Video 4). In the presence of Cpx alone, virtually all docked vSUVs fused spontaneously typically within 1–2 s (Figure 1A,B). This meant that Syt1 somehow synergizes with Cpx to form the overall fusion clamp.

Video 4.

Download video file.(106K, mp4)
Representative video showing the docking, diffusion and spontaneous fusion of a vSUV to free-standing t-SNARE containing lipid bilayer in the presence of 2 µM Cpx.

The movie was acquiredusing a Leica confocal scanning microscope at a speed of 7 frames/sec and the ATTO647N-PE fluorescence was used to track the vesicle. For clarity, a single hole corresponding to 5 µm suspended bilayer marked by the white circle is shown.

Synaptotagmin and complexin establish fast, Ca2+-triggered vesicle fusion

We then investigated the effect of Ca2+ on the stably Syt1/Cpx-clamped vesicles (Figure 2). We estimated the time of arrival of Ca2+ at/near the docked vesicles using a lipid-conjugated Ca2+ indicator (Calcium green C24) attached to the planar bilayer (Figure 2—figure supplement 1). Influx of free Ca2+ (1 mM) triggered simultaneous fusion of >90% of the docked vesicles (Figure 2B, Video 5). These vesicles fused rapidly and synchronously, with a characteristic time-constant (τ) of ~110 msec following the arrival of Ca2+ locally (Figure 2C). This estimate is constrained by the temporal resolution limit (150 msec per frame) of our imaging experiment. Indeed, most of Ca2+-triggered fusion occurs within a single frame (Figure 2C). We thus suspect that the true Ca2+-driven fusion rate is <100 msec. Notably, the Syt1/Cpx clamped vesicles remained Ca2+-sensitive even 1 hr post-docking (Figure 1—figure supplement 3). Our data indicate that Syt1 and Cpx acting together synchronize vesicle fusion to Ca2+ influx and greatly accelerate the underlying SNARE-mediated fusion process (which typically occurs at a rate of ~1 s). We also tested and confirmed these findings with a content-release assay using sulforhodamine B-loaded SUVs (Figure 2—figure supplement 2) under similar experimental conditions. Overall, we find that Syt1 and Cpx act co-operatively to clamp the SNARE assembly process to generate and maintain a pool of docked vesicles that can be triggered to fuse rapidly and synchronously upon Ca2+ influx.

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Syt1/Cpx clamped vesicles fuse synchronously and rapidly following Ca2+-addition.

(A) Representative fluorescence images (top) and quantitation of change in fluorescence signal (bottom) before and after addition of 1 mM Ca2+ shows that vesicles clamped by Syt1/Cpx are sensitive to Ca2+. Fusion was attested by a burst and sudden decrease in fluorescence (ATTO647N-PE) intensity as the lipids diffuse away. To visualize this, fluorescence was simultaneously monitored in a circular region of interest (ROI) encompassing the docked vesicle (vesicle ROI, green and blue circles) and in a surrounding annular ROI (outer ROI, yellow circle). Corresponding to actual fusion events, we observed a sudden decrease of fluorescence intensity in the vesicle ROI with a concomitant increase of fluorescence in the annular outer ROI. Note that the two docked vesicles fuse synchronously in response to Ca2+-influx. (B) End-point analysis at 1 min post Ca2+-addition shows that >90% of all clamped vesicles fuse following Ca2+ addition. (C) Kinetic analysis shows that the Syt1/Cpx clamped vesicles fuse rapidly following Ca2+-addition with a characteristic time constant of 0.11 s. This represents the temporal resolution limit of our recordings and the true Ca2+-triggered fusion rate is likely well below 0.1 s. Thus, Syt1 and Cpx synchronize vesicle fusion to Ca2+-influx and accelerate the underlying fusion process. The average values and standard deviations from three independent experiments (with ~1000 vesicles in total) is shown.

Figure 2—figure supplement 1.

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Ca2+-sensor dye, Calcium Green, introduced in the suspended bilayer (via a lipophilic 24-carbon alkyl chain) was used to monitor the arrival of Ca2+ at/near the docked vesicles.

A representative fluorescence trace acquired at a speed of ~36 msec per frame using confocal microscopy equipped with resonant scanner focused at or near the bilayer membrane is shown. The rise in fluorescence intensity upon Ca2+ binding was observed ~3 frames (~100 ms) after the addition of Ca2+ at the top of chamber. This was used to benchmark and to accurately estimate the Ca2+-triggered fusion rate.

Figure 2—figure supplement 2.

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The effect of Syt1 and Cpx on SNARE-driven fusion assessed using a content-release assay with Sulforhodamine B loaded vesicles.

Sulforhodamine B is largely self-quenched when encapsulated inside a SUV. Fusion of vesicle results in dilution of the probe, which is accompanied by increasing fluorescence. Representative fluorescence trace showing that in the presence of 2 µM Cpx, the Syt1-vSUV (loaded with Sulphorhodamine B) dock and remain in place, until triggered to fuse by Ca2+ (1 mM) addition. This is in line with the lipid mixing data (Figure 2) showing that Syt1 and Cpx act together to establish Ca2+-regulated exocytosis.

Video 5.

Download video file.(2.1M, mp4)
Representative video showing the rapid and synchronous fusion of multiple Syt1/Cpx clamped vesicles to free-standing t-SNARE containing lipid bilayer triggered by Ca2+-influx (1 mM free).

The ATTO647N-DOPE introduced in the SUV was used to track the fate of the vesicle and the movie was acquired using a Leica confocal scanning microscope at a speed of 7 frames/sec. For clarity, a single hole corresponding to 5 µm suspended bilayer marked by the white circle is shown.

Synaptotagmin and complexin clamp different sets of SNARE complexes

We next examined if Syt1 and Cpx act on the same SNARE complexes sequentially or if they function separately to produce molecularly-distinct clamped SNAREpins under the same docked vesicles. To this end, we employed an accessibility-dependent competition assay (Figure 3). We washed Cpx out from stably clamped Syt1/Cpx vesicles (by dilution) in the absence or the presence of excess inhibitory soluble cytoplasmic domain of the t-SNARE complex (CDT). Cpx binds to half-zippered (clamped) SNAREpins with a Kd ~0.5 µM (Krishnakumar et al., 2011; Kümmel et al., 2011) and is therefore expected to freely dissociate when the bulk concentration of CPX is reduced well below that level. CDT will bind and sequester/inactivate any free VAMP2 on the vesicles and is also expected to effectively compete out bilayer-anchored t-SNAREs. This is because CDT can form fully-zippered SNARE complexes (stabilized by ~70 kBT) as compared to the half-zippered SNAREpins (~35 kBT) formed by bilayer-anchored t-SNAREs (Li et al., 2016). We reasoned that if Syt1 and Cpx act on the same SNARE complex, then CDT treatment (with Cpx wash-out) would irreversibly block all vesicle fusion. However, if Syt1 and Cpx clamp different SNARE complexes, then some SNAREpins might be sequestered/protected from CDT by Syt1, thereby keeping the vesicles clamped yet sensitive to Ca2+ influx.

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Syt1 and Cpx bind and clamp different pools of SNAREpins.

(A) In situ removal of Cpx from the Syt1/Cpx clamped state by extensive (40X) buffer wash triggers spontaneous fusion of the docked vesicles. This further confirms that both Syt1 and Cpx are required to produce a stable clamped state. (B) Inclusion of soluble cytoplasmic domain of t-SNAREs (CDT) blocked the spontaneous fusion events triggered by elimination of Cpx from the Syt1/Cpx clamped vesicles. The CDT-treated vesicles remain sensitive to Ca2+ influx and most of the vesicles fuse rapidly and synchronously following the addition of 1 mM Ca2+. This indicates that a sub-set of SNAREpins are protected against CDT even in the absence of Cpx, implying that Syt1 and Cpx likely engage and clamp different set of SNAREpins. It further shows that the Syt1-associated SNAREpins are sufficient to catalyze rapid Ca2+-triggered vesicle fusion. Data (average ± standard deviation) obtained from four to five independent experiments with at least 200 vesicles in total are shown. Note: Only single SNAREpins clamped by either Syt1 or Cpx is shown for illustrative purposes.

Figure 3—figure supplement 1.

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Representative fluorescence images showing that the extensive (40X) buffer wash results in complete washout of Cpx from clamped Syt1/Cpx vesicles.

This was assessed using fluorescently (Alexa488) labeled Cpx and fluorescence recorded prior to the addition of Cpx (left panel), upon addition of Cpx (middle) and after dilution by 40X buffer wash (right panel) are shown. Images corresponding to multiple 5 µm suspended bilayer is shown. Note: The vesicle and bilayer fluorescence are not shown for clarity.

Figure 3—figure supplement 2.

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Respresentative fluorescence images showing the effect of CDT wash and subsequent Ca2+ addition.

(A) Representative fluorescence (ATTO647N) image showing that clamped Syt1/Cpx vesicles (left panel) fuse spontaneously upon wash out of Cpx by dilution. Upon fusion, the ATTO647N dye on the vesicles mixes with suspended bilayer resulting in increase of the background ATTO647N fluorescence (right panel). (B) In the presence of excess inhibitory soluble cytoplasmic domain of the t-SNARE complex (CDT), the vesicles remain docked even after the removal of previously-bound Cpx by the buffer wash (middle panel). Subsequent addition of Ca2+ (1 mM) triggered rapid and synchronous fusion of these docked vesicles. Images corresponding to multiple 5 µm suspended bilayer is shown.

We used fluorescently-labeled Cpx to test and confirm the near-complete washout of Cpx from the clamped Syt1/Cpx vesicles following the extensive (40X) buffer wash (Figure 3—figure supplement 1). Without CDT, the docked vesicles proceeded to fuse spontaneously following the buffer wash (Figure 3A, Figure 3—figure supplement 2). This further confirmed that both Syt1 and Cpx are needed to produce a stably clamped state. In the presence of CDT, most of the vesicles remained docked even after the removal of previously-bound Cpx by the buffer wash (Figure 3B, Figure 3—figure supplement 2). Subsequent addition of Ca2+ (1 mM) triggered rapid and synchronous fusion of the docked vesicles (Figure 3B, Figure 3—figure supplement 2), with fusion kinetics similar to the control experiments (Figure 2C). This implied that there are at least two types of clamped SNAREpins under a docked vesicle – those clamped by Syt1 (which are shielded from CDT) and others arrested by Cpx (which become accessible to CDT following the buffer wash-out). It further indicated that even though both Syt1 and Cpx are required to produce a stably ‘clamped’ vesicle, the activation of the Syt1-associated SNAREpins is sufficient to elicit rapid, Ca2+-synchronized vesicular fusion.

Molecular mechanism of synaptotagmin clamp and Ca2+ activation of fusion

Considering these findings, we sought to establish the molecular determinants of the Syt1 clamp and its reversal by Ca2+. To focus on the Syt1 component of the clamp, we tested the effect of specific Syt1 mutations using low copy VAMP2 conditions, i.e. SUVs containing ~13 copies of VAMP2 and ~25 copies of Syt1 (wild type or mutants) in the absence of Cpx. Consistent with our earlier report (Ramakrishnan et al., 2019), wild-type Syt1 (Syt1WT) alone was sufficient to produce stably-clamped vesicles under these conditions (Figure 4A, Table 3, Table 4). Selective disruption of Syt1-SNARE ‘primary’ binding using the previously described (Zhou et al., 2015; Zhou et al., 2017) mutations in Syt1 C2B domain (R281A/E295A/Y338W/R398A/R399A; Syt1Q) and t-SNARE SNAP25 (K40A/D51A/E52A/E55A/D166A; SNAREQ) abolished the Syt1 clamp (Figure 4A, Table 3, Table 4,), with >99% of the docked Syt1Q vesicles fusing constitutively in the 10 min observation period (Note: From this point onwards, the ‘primary’ site mutation is simply referred as Syt1Q). On the other hand, Syt1 mutations (L387Q/L394Q; Syt1LLQQ) that disrupt the hydrophobic interaction that is an integral part of the SNARE-Cpx-Syt1 ‘tripartite’ interface (Zhou et al., 2017) had no effect on the Syt1 clamp (Figure 4A, Table 3, Table 4). Destabilization of the Syt1 C2B oligomers with a point-mutation (F349A)(Bello et al., 2018) also abrogated the Syt1 clamp, wherein ~ 85% of docked Syt1F349A vesicles proceeded to fuse spontaneously (Figure 4A, Table 3, Table 4). However, disrupting Ca2+ binding to the C2B domain (D309A, D363A, D365A; Syt1DA) (Shao et al., 1996) had no effect on Syt1 clamping function, with all vesicles remaining un-fused (Figure 4A, Table 3, Table 4). This meant that Syt1 ability to oligomerize and bind SNAREpins via the primary binding site is key to its clamping function.

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Molecular determinants of Syt1 clamp and its reversal by Ca2+.

To focus on the Syt1 component of the fusion clamp, all fusion analysis was carried out using vesicles containing low copy VAMP2 (~13 copies) with normal number (~25 copies) of Syt1 molecules (wild-type or targeted mutations) in the absence of Cpx. (A) Survival analysis shows that disrupting the Syt1-SNARE primary interface (Syt1Q, green curve) or destabilizing Syt1 oligomerization (Syt1F349A, purple curve) abrogates the Syt1 clamp, whilst the Syt1-Cpx-SNARE tripartite interface (Syt1LLQQ, yellow curve) and the Ca2+-binding motif on the Syt1 C2B domain (Syt1DA, red curve) are not involved in establishing the fusion clamp. (B) Addition of Ca2+ triggered rapid fusion of the majority (>90%) of Syt1LLQQ and the remainder (~15%) of the docked Syt1F349A vesicles, very similar to the behavior of the Syt1WT vesicles. Predictably, blocking Ca2+-binding to C2B domain rendered the vesicle Ca2+-insensitive, with the majority of Syt1DA remaining un-fused. We did not have sufficient number of docked Syt1Q vesicles to do a quantitative analysis, but qualitatively, the few that remained docked failed to fuse following Ca2+-addition. This implies that both C2B binding to Ca2+ and SNAREs are required for the Ca2+-activation, but the ability to form oligomers or Syt1-Cpx-SNARE tripartite interface are not crucial for the Ca2+-triggered reversal of the Syt1 clamp. The average values and standard deviations from four independent experiments are shown for each condition. In total,~250 vesicles were analyzed for each condition.

Table 3.

Survival probabilities at specific points in time post-docking (Kaplan Meier estimators) for the different Syt1 mutants under low-copy VAMP2 conditions in the absence of Cpx. The corresponding survival curves are shown in Figure 4A
Time (s)
(post-docking)		vSUVSyt1WTSyt1F349ASyt13DASyt1QSyt1LLQQ
0.5880.79250.99650.95970.99360.96120.9975
1.0290.46790.97990.89510.99310.83530.9963
2.0580.16970.95460.59920.98570.54030.9854
3.0870.08990.95460.38220.97940.31930.9525
4.1160.06620.93410.27450.97620.21630.9416
5.1450.06060.93410.22090.97220.17320.9416
7.4970.05490.92550.16630.95970.12530.9160
9.9960.04590.91890.15140.94750.11520.9124

Table 4.

The survival curves in Figure 4A were compared pair-wise using the log-rank test to determine the statistical significance (p-values) of the effects of the Syt1 mutants under low-copy VAMP2 conditions in the absence of Cpx.
vSUVSyt1WTSyt1F349ASyt13DASyt1QSyt1LLQQ
vSUVn/ap<0.0001p<0.004p<0.0001p<0.009p<0.0001
Syt1WTp<0.0001n/ap<0.0001p=0.912p<0.0001p=0.885
Syt1F349Ap<0.004p<0.0001n/ap<0.0001p=0.113p<0.0001
Syt13DAp<0.0001p=0.912p<0.0001n/ap<0.0001p=0.915
Syt1Qp<0.009p<0.0001p=0.113p<0.0001n/ap<0.0001
Syt1LLQQp<0.0001p=0.885p<0.0001p=0.915p<0.0001n/a

Addition of Ca2+ (1 mM) triggered rapid and synchronous fusion of all of the docked Syt1WT and Syt1LLQQ vesicles and the remaining minority fraction (~15%) of the ‘clamped’ Syt1F349A containing vesicles (Figure 4B). In fact, the docked Syt1F349A vesicles were indistinguishable in their behavior from Syt1WT vesicles, suggesting the Syt1 oligomerization is not critical for the Ca2+-activation mechanism. In contrast, docked vesicles containing the Ca2+-binding mutant (Syt1DA) never fused even after Ca2+-addition (Figure 4B). Similarly, Ca2+-influx failed to trigger the fusion of the residual (~1%) Syt1Q vesicles. However, the relatively small number of docked Syt1Q prior to Ca2+-influx precludes any meaningful quantitative analysis. Nonetheless, our data suggest that Ca2+-binding to the Syt1 C2B domain and its simultaneous interaction with the t-SNARE protein via the primary binding site is required for Ca2+-triggered reversal of the fusion clamp.

We also tested the effect of the Syt1 mutants using vesicles containing physiological VAMP2 and Syt1 copy numbers in the presence of 2 µM Cpx (Figure 5A, Table 5, Table 6). For all mutations tested, we observed immobile, stably docked vesicles (Figure 5A, Table 5, Table 6). Surprisingly, the Cpx-dependent clamp was observed even under conditions wherein the Syt1 clamp was absent (i.e. Syt1Q and Syt1F349A). Survival analysis revealed that both Syt1Q and Syt1F349A mutants introduce a meaningful delay in the overall fusion process albeit less than that observed with a stable Syt1 clamp (Figure 5—figure supplement 1). This implies that the kinetic delay introduced by Syt1, independent of its ability to clamp, is sufficient to enable Cpx to function as a fusion clamp, perhaps by providing time for Cpx to bind and block all SNAREpins.

An external file that holds a picture, illustration, etc. Object name is elife-54506-fig5.jpg
Syt1-clamped ‘central’ SNAREpins are required for Ca2+-evoked fusion.

The effect of targeted Syt1 mutations was assessed under physiologically-relevant SUV conditions (~74 copies of VAMP2 and ~25 copies of Syt1 wild type or mutant) in the presence of 2 μM Cpx. (A) Survival analysis shows neither the disruption of the Syt1-SNARE primary interface (Syt1Q, green curve) and the Syt1-Cpx-SNARE tripartite interface (Syt1LLQQ, yellow curve) nor destabilizing Syt1 oligomerization (Syt1F349A, purple curve) has any effect on the fusion clamp in the presence of Cpx, with stably docked vesicles observed under all conditions. (B) End-point analysis indicates that the Syt1Q and Syt1F349A vesicles are insensitive to Ca2+ and do not fuse upon addition of 1 mM Ca2+. In contrast, rapid and synchronous Ca2+-triggered fusion was observed with the majority of the docked Syt1WT and Syt1LLQQ vesicles. Notably, the LLQQ mutation does have a small but significant effect on Ca2+-triggered release. This implies that the Syt1-Cpx-SNARE tripartite interface is not necessary for clamp, but might be involved in the Ca2+-activation process. Overall, the data suggest that the Syt1 clamped SNAREpins are required for Ca2+-triggered exocytosis and in the absence of Syt1 clamp, Cpx irreversibly blocks vesicle fusion. The average values and standard deviations from three independent experiments are shown for each condition. In total,~200 vesicles were analyzed for each condition.

Figure 5—figure supplement 1.

An external file that holds a picture, illustration, etc. Object name is elife-54506-fig5-figsupp1.jpg
Syt1 non-clamping mutants introduce delay in fusion kinetics.

Survival analysis under physiologically-relevant SUVs (~74 copies of VAMP2 and ~25 copies of Syt1 wild type or mutant) in the absence of Cpx shows that Syt1 mutations that abrogate the fusion clamp i.e. primary interface (Syt1Q, green curve) or oligomerization (Syt1F349A, red curve) still introduce a meaningful (~2–3 s) delay in the fusion kinetics although less than observed with Syt1WT (blue curve). This delay might be sufficient for Cpx to bind and irreversibly block all SNARE driven fusion (Figure 5A). The average values and standard deviations from three independent experiments, from a total of ~150 vesicles each condition, is shown.

Figure 5—figure supplement 2.

An external file that holds a picture, illustration, etc. Object name is elife-54506-fig5-figsupp2.jpg
Cpx irreversibly blocks SNARE mediated fusion in the absence of Syt1 clamp.

(A) In situ removal of Cpx from the clamped Syt1Q vesicles by extensive (40X) buffer wash triggers spontaneous fusion of the docked vesicles (B) Inclusion of soluble cytoplasmic domain of t-SNAREs (CDT) blocked the spontaneous fusion events triggered by elimination of Cpx from the clamped Syt1Q vesicles. These CDT-treated vesicles are insensitive to Ca2+ and fail to fuse following the addition of 1 mM Ca2+. This shows in the Syt1Q vesicles, with no Syt1 clamp, all SNAREpins are arrested by Cpx alone and the SNARE-driven fusion is irreversibly blocked by CDT following the Cpx washout. Average and standard deviation from three independent experiments with at least 100 vesicles in total is shown.

Table 5.

Survival probabilities at specific points in time post-docking (Kaplan Meier estimators) for the different Syt1 mutants under normal VAMP2 conditions in the presence of 2 μM Cpx. The corresponding survival curves are shown in Figure 5A.
Time (s)
(post-docking)		vSUVSyt1WTSyt1F349ASyt1QSyt1LLQQ
0.5880.67140.99590.99360.99130.9878
1.0290.29810.99360.99100.98210.9810
2.0580.06170.99270.98570.97340.9688
3.0870.03910.99050.97940.96460.9566
4.1160.02670.98520.97620.95730.9471
5.1450.02430.98410.97220.94450.9444
7.4970.02150.98330.95970.93790.8943
9.9960.01950.98240.94750.93010.8808

Table 6.

The survival curves shown in Figure 5A were compared pair-wise using the log-rank test to determine the statistical significance (p-values) of the effects of the Syt1 mutants under normal VAMP2 conditions in the presence of 2 μM Cpx.
vSUVSyt1WTSyt1F349ASyt1QSyt1LLQQ
v-SUVn/ap<0.0001p<0.0001p<0.0001p<0.0001
Syt1WTp<0.0001n/ap=0.971p=0.912p=0.151
Syt1F349Ap<0.0001p=0.971n/ap=0.865p=0.906
Syt1Qp<0.0001p=0.912p=0.865n/ap=0.426
Syt1LLQQp<0.0001p=0.151p=0.906p=0.426n/a

However, the clamped Syt1Q and Syt1F349A vesicles were insensitive to Ca2+ and did not fuse following Ca2+ (1 mM) addition as opposed to the rapid and synchronous fusion observed with the majority of the Syt1WT and Syt1LLQQ vesicles (Figure 5B). Notably, a significant partial fraction (~25%) of the Syt1LLQQ vesicles remained un-fused even following Ca2+ addition (Figure 5B). This suggests that while the Syt1-Cpx-SNARE tripartite interface is not essential for establishing the fusion clamp, it is likely important in the Ca2+-triggering of fusion.

Taken together, our data indicate that (i) the Syt1 clamp and associated SNAREpins are critical for Ca2+-triggered fusion and (ii) Cpx, on its own, irreversibly blocks vesicle fusion. These conclusions are corroborated by Cpx washout (±CDT) experiments on the Syt1Q vesicles (Figure 5—figure supplement 2). Without CDT, Syt1Q vesicles fused spontaneously following the Cpx washout. In the presence of CDT, the Syt1Q vesicles remained docked following the buffer wash, but were insensitive to Ca2+ and failed to fuse following the addition of 1 mM Ca2+ (Figure 5—figure supplement 2). This denotes that in Syt1Q vesicles (and presumably in Syt1F349A vesicles), all SNAREpins are clamped by Cpx alone and become accessible to CDT block following the Cpx wash-out.

Discussion

Here we report that Syt1 and Cpx act concomitantly to clamp SNARE-driven constitutive fusion events (Figure 1). We find there are at least two distinct sets of clamped SNAREpins under every docked vesicle – a small population that is reversibly clamped by Syt1 oligomers and the remainder that is irreversibly blocked by Cpx. (Figure 3). These results taken together with the known structural and biochemical properties of Syt1 and Cpx prompts a novel ‘synergistic clamping’ mechanism.

We posit that the Syt1 C2B domain binds PIP2 (via the poly-lysine motif) on the PM and assembles into ring-like oligomeric structures (Bello et al., 2018; Wang et al., 2014; Zanetti et al., 2016). The Syt1 oligomers concurrently bind the t-SNAREs via the ‘primary’ interface (Zhou et al., 2015; Zhou et al., 2017; Grushin et al., 2019). The Syt1-t-SNARE interaction, which likely precedes the engagement of the v- and the t-SNAREs, positions the Syt1 such that it sterically blocks the full assembly of the associated SNAREpins (Grushin et al., 2019). The Syt1 oligomers in addition to creating a stable steric impediment could also radially restrain the assembling SNAREpins (Grushin et al., 2019; Rothman et al., 2017). It is worth noting in this configuration the helical extension of Syt1 C2B that forms the ‘tripartite’ interface with Cpx and SNAREs contacts the PM and is thus unavailable for tripartite binding (Rothman et al., 2017; Volynski and Krishnakumar, 2018; Grushin et al., 2019).

The Syt1 oligomers can bind and clamp only a small sub-set of SNAREpins (which we refer to as ‘central SNAREpins’) as the number of potential SNAREpins per SV far exceeds the Syt1 density (~70 copies of VAMP2 vs. ~20 copies of Syt1). We suggest that the remaining ‘peripheral’ SNAREs would then be unrestrained and can fully zipper to catalyze vesicle fusion, though at an impeded rate (Figure 1). It is tempting to speculate that the limited set of Syt1-clamped ‘central’ SNAREpins correspond physiologically to the six symmetrically organized protein densities that underlie each docked synaptic-like vesicle visualized by cryo-electron tomography analysis (Li et al., 2019). Indeed, this symmetrical organization depends on Syt1 oligomerization (Li et al., 2019).

Cpx potentially binds the SNARE complex only when the Syntaxin and VAMP2 are partially-assembled (Chen et al., 2002; Kümmel et al., 2011) and competitively blocks the complete zippering of the C-terminal portion of the VAMP2 SNARE motif (Kümmel et al., 2011; Li et al., 2011; Giraudo et al., 2009). It is well-established that the C-terminal portion of VAMP2 assembles with extraordinarily high energy and rate acting as the major power stroke to drive membrane fusion (Gao et al., 2012). Consequently, Cpx, on its own, is ineffective in clamping SNARE-driven vesicle fusion (Figure 1).

We envision that under physiological conditions, Syt1 sets the stage for Cpx to bind and clamp the ‘peripheral’ SNARE complexes. Syt1, in addition to fully-arresting the central SNAREpins, also kinetically hinders the assembly of the peripheral SNAREpins enabling Cpx to effectually arrest their complete zippering. In this manner, the sequential action of Syt1 and Cpx on different populations of SNAREpins would be involved in the generation of the stably docked, release-ready vesicles. Under each vesicle, Syt1 oligomers bind and clamp a small sub-set of available SNAREpins at the early stages of docking, which in turn enables Cpx to block the remainder of the SNAREpins. This molecular model would readily explain the importance of both Syt1 and Cpx in establishing the fusion clamp as evidenced in several genetic deletion studies (Bacaj et al., 2013; Geppert et al., 1994; Cho et al., 2014; Martin et al., 2011; Littleton et al., 1993; Yang et al., 2013).

We also report that the reversal of the Syt1 clamp is sufficient to drive rapid Ca2+-triggered fusion of the docked vesicles (Figure 2). It involves Syt1 C2B domain binding both Ca2+ and the SNARE complex (Figure 4). We have recently demonstrated that Ca2+-binding to Syt1 C2 domains induce a large-scale conformational rearrangement of the Syt1-SNARE complex on the lipid membrane surface, disrupting the pre-fusion clamped architecture (Grushin et al., 2019). This, in effect, reverses the Syt1 clamp, allowing the associated SNAREs to complete zippering and drive fusion. This implies that only a small fraction of available SNAREpins per vesicle (i.e. only those associated with Syt1) are involved in the Ca2+-activation process. This is consistent with the earlier reports that 2–3 SNARE complexes can be sufficient to facilitate Ca2+-evoked synchronous neurotransmitter release (Sinha et al., 2011; Mohrmann et al., 2010). Indeed, recent modeling studies considering the concept of mechanical coupling have predicted that an optimum of 4–6 SNAREpins is required to achieve sub-millisecond vesicular release (Manca et al., 2019).

There is a long-standing debate over the role of Cpx in establishing a fusion clamp (Yang et al., 2013; López-Murcia et al., 2019). We find that Cpx is an integral part of the overall clamping mechanism and Syt1 and Cpx play distinct roles in clamping different pools of SNAREpins. However, Cpx requires a kinetic delay (likely introduced by Syt1) to block vesicle fusion and this inhibition is not released by Ca2+. This raises the intriguing possibility that the Cpx clamp is not necessarily reversed during the Ca2+-activation process. The molecular details of the observed Cpx ‘clamp’ and its physiological relevance remains to be determined.

Overall, our data are in alignment with the emerging view that Syt1 plays a pivotal role in orchestrating Ca2+-regulated neurotransmitter release. Syt1 functions both as a fusion clamp and the principal Ca2+-sensor to establish Ca2+-regulation of vesicular fusion (Geppert et al., 1994; Littleton et al., 1993). Furthermore, Syt1, by virtue of self-assembling into oligomeric structures, also provides the molecular framework to organize the exocytic machinery into a co-operative structure to enable ultra-fast fusion (Li et al., 2019; Rothman et al., 2017; Volynski and Krishnakumar, 2018).

We have articulated the simplest hypothesis, considering discrete ‘central’ and ‘’peripheral’ SNAREpins associated with Syt1 and Cpx, respectively. However, it is easy to imagine that Cpx also binds the central SNAREpins. Indeed, recent X-ray crystal structure revealed that both Syt1 and Cpx can bind the same pre-fusion SNARE complex (Zhou et al., 2017). Cpx binding is also predicted to create a new Syt1 binding site (i.e. tripartite interface) on SNAREs. In our reconstituted setup, we find that this SNARE-Cpx-Syt1 ‘tripartite’ interface is not required to produce the fusion clamp but is likely involved in Ca2+ activation of fusion from the clamped state (Figure 5).

Since the SNARE-Syt1-Cpx tripartite interface is created only when the SNAREs are partially-assembled (i.e. when Cpx binds), it is possible that this interaction plays an auxiliary role in establishing the fusion clamp. As such, it can be bypassed in our biochemically-defined experimental setup. The tripartite interface might become more relevant in the pre-synaptic terminals (Zhou et al., 2017) where ~ 30% of Syt1 is present in the PM (Wienisch and Klingauf, 2006) and other Synaptotagmins could also participate in the tripartite interface (Rothman et al., 2017; Volynski and Krishnakumar, 2018; Zhou et al., 2017).

Indeed, we have postulated that such a ‘dual-clamp’ arrangement with Syt1 (from SV) occupying the ‘primary’ site and with Syt1 and Syt7 (from PM) competing for the ‘tripartite’ site (in conjunction with Cpx) could potentially explain the synergistic regulation of neurotransmitter release by different Syt isoforms (Rothman et al., 2017; Volynski and Krishnakumar, 2018). Such an arrangement could also potentially explain how Cpx could regulate Ca2+-evoked synchronous neurotransmitter release.

Physiologically, all three synaptic SNAREs (Syntaxin, SNAP-25 and VAMP2) are assembled into a SNAREpin in a concerted reaction involving chaperones Munc13 and Munc18 (Baker and Hughson, 2016; Rizo, 2018), the need for which is by-passed in our system by pre-assembled t-SNARE complexes. We imagine that these chaperones also function in a concerted manner with Syt1 and Cpx and have recently advanced a speculative model outlining how these proteins can co-operate to template SNAREpin assembly (Rothman et al., 2017). Moreover, lipid membranes may contribute to synergism or cooperativity between Syt1 and Cpx in both clamping the un-initiated fusion events and triggering rapid and synchronous fusion in response to Ca2+-influx.

As such, further studies with high temporal resolution involving detailed mutational and topological analysis of Syt1 and Cpx, along with the chaperones Munc13 and Munc18, is needed to establish pointillistic details of fast Ca2+-triggered SV fusion. Nonetheless, our data demonstrate that Syt1 and Cpx, along with SNARE proteins, form the minimal protein machinery that is necessary and sufficient to establish rapid Ca2+-regulated exocytosis.

Materials and methods

Materials

The following cDNA constructs, which have been previously described (Krishnakumar et al., 2013; Krishnakumar et al., 2011; Weber et al., 1998; Mahal et al., 2002), were used in this study: full-length VAMP2 (VAMP2-His6, residues 1–116); full-length VAMP24X (VAMP2-His6, residues 1–116 with L70D, A74R, A81D, L84D mutations), full-length t-SNARE complex (mouse His6-SNAP25B, residues 1–206 and rat Syntaxin1A, residues 1–288); soluble cytoplasmic domain of the t-SNAREs (CDT, mouse His6-SNAP25B, residues 1–206 and rat Syntaxin1A, residues 1–265); Synaptotagmin (rat Synaptotagmin1-His6, residues 57–421); and Complexin (human His6-Complexin 1, residues 1–134). All mutants including Syt1F349A (F349A); Syt1Q (R281A/E295A/Y338W/R398A/R399A); Syt1LLQQ (L387Q/L394Q); Syt1DA (D309A, D363A, D365A) and SNAREQ (SNAP25 K40A/D51A/E52A/E55A/D166A) were generated in the above described Syt1 and t-SNARE background respectively using the QuickChange mutagenesis kit (Agilent Technologies, Santa Clara, CA). Lipids, 1,2-dioleoyl -snglycero-3-phosphocholine (DOPC), 1,2-dioleoyl-sn-glycero-3- (phospho-L-serine) (DOPS), 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2–1,3-benzoxadiazol-4-yl) (NBD-DOPE), phosphatidylinositol 4, 5-bisphosphate (PIP2) were purchased from Avanti Polar Lipids (Alabaster, AL). ATTO647N-DOPE was purchased from ATTO-TEC, GmbH (Siegen, Germany) and lipophilic carbocyanine DiD (1,1'-Dioctadecyl-3,3,3',3'-Tetramethylindodicarbocyanine Perchlorate) was purchased from Thermofisher Scientific (Waltham, MA). Calcium Green conjugated to a lipophilic 24-carbon alkyl chain (Calcium Green C24) was custom synthesized by Marker Gene Technologies (Eugene, OR).

Protein expression and purification

All proteins (v- and t-SNAREs, Cpx, Syt1 wild type and mutants) were expressed and purified as described previously (Mahal et al., 2002; Krishnakumar et al., 2013; Krishnakumar et al., 2011; Weber et al., 1998). In brief, proteins were expressed in E. coli strain Rosetta2(DE3) (Novagen, Madison, WI) using 0.5 mM IPTG for 4 hr. Cells were pelleted and lysed using a cell disruptor (Avestin, Ottawa, Canada) in HEPES buffer (25 mM HEPES, 400 mM KCl, 4% Triton X-100, 10% glycerol, pH 7.4) containing 0.2 mM Tris(2-carboxyethyl) phosphinehydrochloride (TCEP), and 1 mM phenylmethylsulfonyl fluoride (PMSF). Samples were clarified using a 45Ti rotor (Beckman Coulter, Atlanta, GA) at 40 K RPM for 30 min and subsequently incubated with Ni-NTA resin (Thermofisher Scientific, Waltham, MA) overnight at 4°C. The resin was washed with HEPES buffer (for Cpx) or HEPES buffer supplemented with 1% octylglucoside (Syt1 and SNAREs). Protein was eluted using 350 mM Imidazole and the concentration was determined using a Bradford Assay (BioRad, Hercules, CA) with BSA as a standard. Syt1 was further treated with Benzonase (Millipore Sigma, Burlington, MA) at room temperature for 1 hr, followed by ion exchange (Mono S, AKTA purifier, GE) to remove DNA/RNA contamination. The purity was verified using SDS-PAGE analysis and all proteins were flash frozen and stored at −80°C with 10% glycerol without significant loss of function.

Liposome preparation

 t-SNAREs and VAMP2 (±Syt1) containing SUV were prepared using rapid detergent (1% Octylglucoside) dilution and dialysis method as described previously (Weber et al., 1998; Ji et al., 2010). VAMP2 (±Syt1) containing SUVs were subjected to additional purification on the discontinuous Nycodenz gradient. The lipid composition was 80 (mole)% DOPC, 15% DOPS, 3% PIP2% and 2% NBD-PE for t-SNARE SUV and 88% DOPC, 10% PS and 2% ATTO647-PE for VAMP2 (±Syt1) SUVs. To mimic physiological copy numbers of protein, we used an input of protein: lipid ratio as 1: 400 for t-SNARE, 1:100 for VAMP2 for physiological density, 1: 500 for VAMP2 at low copy number, and 1: 250 for Syt1. This was based on well-established parameters namely that the reconstitution efficiency for SNAREs and Syt1 is roughly 40–50% (densitometry analysis of the proteoliposomes) and only approximately 50–60% of the proteins are externally oriented (chymotrypsin protection analysis) (Ji et al., 2010; Ramakrishnan et al., 2019; Weber et al., 1998). Based on the densitometry analysis of Coomassie-stained SDS gels, we estimated vesicles at physiological density, contained 74 ± 4 and 26 ± 6 copies of outward-facing VAMP2 and Syt1 respectively (Figure 1—figure supplement 1) and vesicles at low copy number of VAMP2 contained 13 ± 2 and 26 ± 6 copies of outward-facing VAMP2 and Syt1 respectively.

Single vesicle fusion assay

All the single-vesicle fusion measurements were carried out with suspended lipid bilayers as previously described (Ramakrishnan et al., 2019; Ramakrishnan et al., 2018). Briefly, t-SNARE-containing giant unilamellar vesicles as prepared using the osmotic shock protocol (Motta et al., 2015) were busted on freshly plasma-cleaned Si/SiO2 chips containing 5 µm diameter holes in presence of HEPES buffer (25 mM HEPES, 140 mM KCl, 1 mM DTT) supplemented with 5 mM MgCl2. The bilayers were extensively washed with HEPES buffer containing 1 mM MgCl2 and the fluidity of the t-SNARE containing bilayers was verified using fluorescence recovery after photo-bleaching using the NBD fluorescence. In some experiments, we labeled the t-SNAREs with Alexa-488 and confirmed protein mobility as described previously (Ramakrishnan et al., 2018).

Vesicles (100 nM lipids) were added from the top using a pipette and allowed to interact with the bilayer for 10 min. We used the ATTO647-PE fluorescence to track vesicle docking, post-docking diffusion, docking-to-fusion delays and spontaneous fusion events. Fusion was attested by a burst and then rapid decrease in fluorescence intensity as the fluorescent PE from the vesicle diffuses away. The time between docking and fusion corresponded to the fusion clamp and was quantified using a ‘survival curve’ whereby delays are pooled together, and their distribution is plotted in the form of a survival function (Figure 1). After the initial 10 min interaction phase, the excess vesicles in the chamber were removed by buffer exchange (3x buffer wash) and 1 mM CaCl2 was added from the top to monitor the effect of Ca2+ on the docked vesicles. The number of fused (and the remaining un-fused) vesicles was estimated (end-point analysis)~1 min after Ca2+-addition.

All experiments were carried out at 37°C using an inverted laser scanning confocal microscope (Leica-SP5) equipped with a multi-wavelength argon laser including 488, diode lasers (532 and 641 nm), and a long-working distance 40X water immersion objective (NA 1.1). The emission light was spectrally separated and collected by photomultiplier tubes. To cover large areas of the planar bilayer and simultaneously record large ensembles of vesicles, the movies were acquired at a speed of 150 ms per frame. Accurate quantification and fate of each vesicles were analyzed using our custom written MATLAB script described previously (Ramakrishnan et al., 2018). The files can be downloaded from MATLAB Central at the following website: https://www.mathworks.com/matlabcentral/fileexchange/66521-fusion-analyzer-fas. Note: We excluded vesicles bound to the edge of the holes as they may not be representative of vesicles bound to the free-floating membrane. We thus used only the centrally-docked vesicles for analysis. We did not observe any change in ATTO-647-PE fluorescence for the vesicles that remain docked and un-fused during the observation period or post Ca2+-addition. Thus, we can rule out hemi-fusion diaphragm formation as a possible explanation for the observed ‘clamped’ or ‘un-fused’ state.

Single-vesicle docking analysis

To get an accurate count of the docked vesicles, we used VAMP2 protein with mutations in the C-terminal half (L70D, A74R, A81D and L84D; VAMP24X) that eliminates fusion without impeding the docking process (Krishnakumar et al., 2013). For the docking analysis, VAMP24X containing SUVs (vSUV4X) were introduced into the chamber and allowed to interact with the t-SNARE bilayer. After a 10 min incubation, the bilayer was thoroughly washed with running buffer (3x minimum) and the number of docked vesicles were counted. For an unbiased particle count, we employed a custom-written algorithm to count particles from top-left to bottom-right that ensures every spot is counted only once.

Calcium dynamics 

To quantify the kinetics of Ca2+-triggered vesicle fusion, we used a Ca2+-sensor dye, Calcium Green conjugated to a lipophilic 24-carbon alkyl chain (Calcium Green C24) introduced in the suspended bilayer to directly monitor the arrival of Ca2+. Calcium green is a high-affinity Ca2+-sensor (Kd of ~75 nM) and exhibits a large fluorescent increase (at 532 nm) upon binding Ca2+. To accurately estimate the arrival of Ca2+ to/near the vesicles docked on the bilayer, we used confocal microscopy equipped with resonant scanner focused at or near the bilayer membrane and acquired movies at a speed of up to 36 msec per frame. We typically observed the fluorescence signal increase at the bilayer surface between about three frames (~100 msec) after Ca2+ addition (Figure 2—figure supplement 1). We therefore used 100 msec as the benchmark to accurately estimate the time-constants for the Ca2+-triggered fusion reaction.

Acknowledgements

We thank Dr. Frederic Pincet for critical inputs during the design and development of this project. This work was supported by National Institute of Health (NIH) grant DK027044 to JER.

Funding Statement

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

Contributor Information

Axel T Brunger, Stanford University, United States.

Vivek Malhotra, The Barcelona Institute of Science and Technology, Spain.

Funding Information

This paper was supported by the following grant:

  • National Institute of Diabetes and Digestive and Kidney Diseases 027044 to James E Rothman.

Additional information

Competing interests

No competing interests declared.

Author contributions

Conceptualization, Data curation, Software, Formal analysis, Investigation, Methodology, Writing - review and editing.

Data curation, Formal analysis, Investigation, Methodology, Writing - review and editing.

Resources, Investigation, Methodology, Writing - review and editing.

Conceptualization, Supervision, Funding acquisition, Methodology, Writing - original draft, Project administration, Writing - review and editing.

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

Additional files

Source data 1.

Ramakrishan et al_2020_Source Data.

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Data availability

All data generated or analysed during this study are included in the manuscript and supporting files, including Source data 1.

References

  • Bacaj T, Wu D, Yang X, Morishita W, Zhou P, Xu W, Malenka RC, Südhof TC. Synaptotagmin-1 and synaptotagmin-7 trigger synchronous and asynchronous phases of neurotransmitter release. Neuron. 2013;80:947–959. doi: 10.1016/j.neuron.2013.10.026. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Baker RW, Hughson FM. Chaperoning SNARE assembly and disassembly. Nature Reviews Molecular Cell Biology. 2016;17:465–479. doi: 10.1038/nrm.2016.65. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Bello OD, Jouannot O, Chaudhuri A, Stroeva E, Coleman J, Volynski KE, Rothman JE, Krishnakumar SS. Synaptotagmin oligomerization is essential for calcium control of regulated exocytosis. PNAS. 2018;115:E7624–E7631. doi: 10.1073/pnas.1808792115. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Brose N, Petrenko AG, Südhof TC, Jahn R. Synaptotagmin: a calcium sensor on the synaptic vesicle surface. Science. 1992;256:1021–1025. doi: 10.1126/science.1589771. [PubMed] [CrossRef] [Google Scholar]
  • Brunger AT, Choi UB, Lai Y, Leitz J, Zhou Q. Molecular mechanisms of fast neurotransmitter release. Annual Review of Biophysics. 2018;47:469–497. doi: 10.1146/annurev-biophys-070816-034117. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Chen X, Tomchick DR, Kovrigin E, Araç D, Machius M, Südhof TC, Rizo J. Three-dimensional structure of the complexin/SNARE complex. Neuron. 2002;33:397–409. doi: 10.1016/S0896-6273(02)00583-4. [PubMed] [CrossRef] [Google Scholar]
  • Cho RW, Kümmel D, Li F, Baguley SW, Coleman J, Rothman JE, Littleton JT. Genetic analysis of the complexin trans-clamping model for cross-linking SNARE complexes in vivo. PNAS. 2014;111:10317–10322. doi: 10.1073/pnas.1409311111. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Coleman J, Jouannot O, Ramakrishnan SK, Zanetti MN, Wang J, Salpietro V, Houlden H, Rothman JE, Krishnakumar SS. PRRT2 regulates synaptic fusion by directly modulating SNARE complex assembly. Cell Reports. 2018;22:820–831. doi: 10.1016/j.celrep.2017.12.056. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Courtney NA, Bao H, Briguglio JS, Chapman ER. Synaptotagmin 1 clamps synaptic vesicle fusion in mammalian neurons independent of complexin. Nature Communications. 2019;10:4076. doi: 10.1038/s41467-019-12015-w. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Fuson KL, Montes M, Robert JJ, Sutton RB. Structure of human synaptotagmin 1 C2AB in the absence of Ca2+ reveals a novel domain association. Biochemistry. 2007;46:13041–13048. doi: 10.1021/bi701651k. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Gao Y, Zorman S, Gundersen G, Xi Z, Ma L, Sirinakis G, Rothman JE, Zhang Y. Single reconstituted neuronal SNARE complexes zipper in three distinct stages. Science. 2012;337:1340–1343. doi: 10.1126/science.1224492. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Geppert M, Goda Y, Hammer RE, Li C, Rosahl TW, Stevens CF, Südhof TC. Synaptotagmin I: a major Ca2+ sensor for transmitter release at a central synapse. Cell. 1994;79:717–727. doi: 10.1016/0092-8674(94)90556-8. [PubMed] [CrossRef] [Google Scholar]
  • Giraudo CG, Garcia-Diaz A, Eng WS, Chen Y, Hendrickson WA, Melia TJ, Rothman JE. Alternative zippering as an on-off switch for SNARE-mediated fusion. Science. 2009;323:512–516. doi: 10.1126/science.1166500. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Grushin K, Wang J, Coleman J, Rothman JE, Sindelar CV, Krishnakumar SS. Structural basis for the clamping and Ca2+ activation of SNARE-mediated fusion by synaptotagmin. Nature Communications. 2019;10:2413. doi: 10.1038/s41467-019-10391-x. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Honigmann A, van den Bogaart G, Iraheta E, Risselada HJ, Milovanovic D, Mueller V, Müllar S, Diederichsen U, Fasshauer D, Grubmüller H, Hell SW, Eggeling C, Kühnel K, Jahn R. Phosphatidylinositol 4,5-bisphosphate clusters act as molecular beacons for vesicle recruitment. Nature Structural & Molecular Biology. 2013;20:679–686. doi: 10.1038/nsmb.2570. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Huntwork S, Littleton JT. A complexin fusion clamp regulates spontaneous neurotransmitter release and synaptic growth. Nature Neuroscience. 2007;10:1235–1237. doi: 10.1038/nn1980. [PubMed] [CrossRef] [Google Scholar]
  • Ji H, Coleman J, Yang R, Melia TJ, Rothman JE, Tareste D. Protein determinants of SNARE-mediated lipid mixing. Biophysical Journal. 2010;99:553–560. doi: 10.1016/j.bpj.2010.04.060. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Kaeser PS, Regehr WG. Molecular mechanisms for synchronous, asynchronous, and spontaneous neurotransmitter release. Annual Review of Physiology. 2014;76:333–363. doi: 10.1146/annurev-physiol-021113-170338. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Krishnakumar SS, Radoff DT, Kümmel D, Giraudo CG, Li F, Khandan L, Baguley SW, Coleman J, Reinisch KM, Pincet F, Rothman JE. A conformational switch in complexin is required for synaptotagmin to trigger synaptic fusion. Nature Structural & Molecular Biology. 2011;18:934–940. doi: 10.1038/nsmb.2103. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Krishnakumar SS, Kümmel D, Jones SJ, Radoff DT, Reinisch KM, Rothman JE. Conformational dynamics of calcium-triggered activation of fusion by synaptotagmin. Biophysical Journal. 2013;105:2507–2516. doi: 10.1016/j.bpj.2013.10.029. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Kümmel D, Krishnakumar SS, Radoff DT, Li F, Giraudo CG, Pincet F, Rothman JE, Reinisch KM. Complexin cross-links prefusion SNAREs into a zigzag array. Nature Structural & Molecular Biology. 2011;18:927–933. doi: 10.1038/nsmb.2101. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Li F, Pincet F, Perez E, Giraudo CG, Tareste D, Rothman JE. Complexin activates and clamps SNAREpins by a common mechanism involving an intermediate energetic state. Nature Structural & Molecular Biology. 2011;18:941–946. doi: 10.1038/nsmb.2102. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Li F, Tiwari N, Rothman JE, Pincet F. Kinetic barriers to SNAREpin assembly in the regulation of membrane docking/priming and fusion. PNAS. 2016;113:10536–10541. doi: 10.1073/pnas.1604000113. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Li X, Radhakrishnan A, Grushin K, Kasula R, Chaudhuri A, Gomathinayagam S, Krishnakumar SS, Liu J, Rothman JE. Symmetrical organization of proteins under docked synaptic vesicles. FEBS Letters. 2019;593:144–153. doi: 10.1002/1873-3468.13316. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Littleton JT, Stern M, Schulze K, Perin M, Bellen HJ. Mutational analysis of Drosophila synaptotagmin demonstrates its essential role in ca(2+)-activated neurotransmitter release. Cell. 1993;74:1125–1134. doi: 10.1016/0092-8674(93)90733-7. [PubMed] [CrossRef] [Google Scholar]
  • López-Murcia FJ, Reim K, Jahn O, Taschenberger H, Brose N. Acute complexin knockout abates spontaneous and evoked transmitter release. Cell Reports. 2019;26:2521–2530. doi: 10.1016/j.celrep.2019.02.030. [PubMed] [CrossRef] [Google Scholar]
  • Mahal LK, Sequeira SM, Gureasko JM, Söllner TH. Calcium-independent stimulation of membrane fusion and SNAREpin formation by synaptotagmin I. Journal of Cell Biology. 2002;158:273–282. doi: 10.1083/jcb.200203135. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Manca F, Pincet F, Truskinovsky L, Rothman JE, Foret L, Caruel M. SNARE machinery is optimized for ultrafast fusion. PNAS. 2019;116:2435–2442. doi: 10.1073/pnas.1820394116. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Martin JA, Hu Z, Fenz KM, Fernandez J, Dittman JS. Complexin has opposite effects on two modes of synaptic vesicle fusion. Current Biology. 2011;21:97–105. doi: 10.1016/j.cub.2010.12.014. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Mohrmann R, de Wit H, Verhage M, Neher E, Sørensen JB. Fast vesicle fusion in living cells requires at least three SNARE complexes. Science. 2010;330:502–505. doi: 10.1126/science.1193134. [PubMed] [CrossRef] [Google Scholar]
  • Motta I, Gohlke A, Adrien V, Li F, Gardavot H, Rothman JE, Pincet F. Formation of giant unilamellar Proteo-Liposomes by osmotic shock. Langmuir. 2015;31:7091–7099. doi: 10.1021/acs.langmuir.5b01173. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Parisotto D, Malsam J, Scheutzow A, Krause JM, Söllner TH. SNAREpin assembly by Munc18-1 requires previous vesicle docking by synaptotagmin 1. Journal of Biological Chemistry. 2012;287:31041–31049. doi: 10.1074/jbc.M112.386805. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Park Y, Hernandez JM, van den Bogaart G, Ahmed S, Holt M, Riedel D, Jahn R. Controlling synaptotagmin activity by electrostatic screening. Nature Structural & Molecular Biology. 2012;19:991–997. doi: 10.1038/nsmb.2375. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Ramakrishnan S, Gohlke A, Li F, Coleman J, Xu W, Rothman JE, Pincet F. High-Throughput monitoring of single vesicle fusion using freestanding membranes and automated analysis. Langmuir. 2018;34:5849–5859. doi: 10.1021/acs.langmuir.8b00116. [PubMed] [CrossRef] [Google Scholar]
  • Ramakrishnan S, Bera M, Coleman J, Krishnakumar SS, Pincet F, Rothman JE. Synaptotagmin oligomers are necessary and can be sufficient to form a Ca2+ -sensitive fusion clamp. FEBS Letters. 2019;593:154–162. doi: 10.1002/1873-3468.13317. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Rizo J. Mechanism of neurotransmitter release coming into focus. Protein Science. 2018;27:1364–1391. doi: 10.1002/pro.3445. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Rothman JE, Krishnakumar SS, Grushin K, Pincet F. Hypothesis - buttressed rings assemble, clamp, and release SNAREpins for synaptic transmission. FEBS Letters. 2017;591:3459–3480. doi: 10.1002/1873-3468.12874. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Shao X, Davletov BA, Sutton RB, Südhof TC, Rizo J. Bipartite Ca2+-binding motif in C2 domains of synaptotagmin and protein kinase C. Science. 1996;273:248–251. doi: 10.1126/science.273.5272.248. [PubMed] [CrossRef] [Google Scholar]
  • Sinha R, Ahmed S, Jahn R, Klingauf J. Two synaptobrevin molecules are sufficient for vesicle fusion in central nervous system synapses. PNAS. 2011;108:14318–14323. doi: 10.1073/pnas.1101818108. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Söllner T, Whiteheart SW, Brunner M, Erdjument-Bromage H, Geromanos S, Tempst P, Rothman JE. SNAP receptors implicated in vesicle targeting and fusion. Nature. 1993;362:318–324. doi: 10.1038/362318a0. [PubMed] [CrossRef] [Google Scholar]
  • Südhof TC. Neurotransmitter release: the last millisecond in the life of a synaptic vesicle. Neuron. 2013;80:675–690. doi: 10.1016/j.neuron.2013.10.022. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Südhof TC, Rothman JE. Membrane fusion: grappling with SNARE and SM proteins. Science. 2009;323:474–477. doi: 10.1126/science.1161748. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Sutton RB, Davletov BA, Berghuis AM, Südhof TC, Sprang SR. Structure of the first C2 domain of synaptotagmin I: a novel Ca2+/phospholipid-binding fold. Cell. 1995;80:929–938. doi: 10.1016/0092-8674(95)90296-1. [PubMed] [CrossRef] [Google Scholar]
  • Tagliatti E, Bello O, Mendonca PRF, Kotzadimitriou D, Nicholson E, Coleman J, Timofeeva Y, Rothman JE, Krishnakumar SS, Volynski KE. Synaptotagmin oligomers clamp and regulate different modes of neurotransmitter release. bioRxiv. 2019 doi: 10.1101/594051. [PMC free article] [PubMed] [CrossRef]
  • Volynski KE, Krishnakumar SS. Synergistic control of neurotransmitter release by different members of the synaptotagmin family. Current Opinion in Neurobiology. 2018;51:154–162. doi: 10.1016/j.conb.2018.05.006. [PubMed] [CrossRef] [Google Scholar]
  • Wang J, Bello O, Auclair SM, Wang J, Coleman J, Pincet F, Krishnakumar SS, Sindelar CV, Rothman JE. Calcium sensitive ring-like oligomers formed by synaptotagmin. PNAS. 2014;111:13966–13971. doi: 10.1073/pnas.1415849111. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Weber T, Zemelman BV, McNew JA, Westermann B, Gmachl M, Parlati F, Söllner TH, Rothman JE. SNAREpins: minimal machinery for membrane fusion. Cell. 1998;92:759–772. doi: 10.1016/S0092-8674(00)81404-X. [PubMed] [CrossRef] [Google Scholar]
  • Wienisch M, Klingauf J. Vesicular proteins exocytosed and subsequently retrieved by compensatory endocytosis are nonidentical. Nature Neuroscience. 2006;9:1019–1027. doi: 10.1038/nn1739. [PubMed] [CrossRef] [Google Scholar]
  • Wragg RT, Snead D, Dong Y, Ramlall TF, Menon I, Bai J, Eliezer D, Dittman JS. Synaptic vesicles position complexin to block spontaneous fusion. Neuron. 2013;77:323–334. doi: 10.1016/j.neuron.2012.11.005. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Xu J, Mashimo T, Südhof TC. Synaptotagmin-1, -2, and -9: ca(2+) sensors for fast release that specify distinct presynaptic properties in subsets of neurons. Neuron. 2007;54:567–581. doi: 10.1016/j.neuron.2007.05.004. [PubMed] [CrossRef] [Google Scholar]
  • Xu J, Pang ZP, Shin OH, Südhof TC. Synaptotagmin-1 functions as a Ca2+ sensor for spontaneous release. Nature Neuroscience. 2009;12:759–766. doi: 10.1038/nn.2320. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Xu W, Nathwani B, Lin C, Wang J, Karatekin E, Pincet F, Shih W, Rothman JE. A programmable DNA origami platform to organize SNAREs for membrane fusion. Journal of the American Chemical Society. 2016;138:4439–4447. doi: 10.1021/jacs.5b13107. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Yang X, Cao P, Südhof TC. Deconstructing complexin function in activating and clamping Ca2+-triggered exocytosis by comparing knockout and knockdown phenotypes. PNAS. 2013;110:20777–20782. doi: 10.1073/pnas.1321367110. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Yang X, Pei J, Kaeser-Woo YJ, Bacaj T, Grishin NV, Südhof TC. Evolutionary conservation of complexins: from choanoflagellates to mice. EMBO Reports. 2015;16:1308–1317. doi: 10.15252/embr.201540305. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Zanetti MN, Bello OD, Wang J, Coleman J, Cai Y, Sindelar CV, Rothman JE, Krishnakumar SS. Ring-like oligomers of synaptotagmins and related C2 domain proteins. eLife. 2016;5:e17262. doi: 10.7554/eLife.17262. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Zhou Q, Lai Y, Bacaj T, Zhao M, Lyubimov AY, Uervirojnangkoorn M, Zeldin OB, Brewster AS, Sauter NK, Cohen AE, Soltis SM, Alonso-Mori R, Chollet M, Lemke HT, Pfuetzner RA, Choi UB, Weis WI, Diao J, Südhof TC, Brunger AT. Architecture of the synaptotagmin-SNARE machinery for neuronal exocytosis. Nature. 2015;525:62–67. doi: 10.1038/nature14975. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
  • Zhou Q, Zhou P, Wang AL, Wu D, Zhao M, Südhof TC, Brunger AT. The primed SNARE-complexin-synaptotagmin complex for neuronal exocytosis. Nature. 2017;548:420–425. doi: 10.1038/nature23484. [PMC free article] [PubMed] [CrossRef] [Google Scholar]
2020; 9: e54506.
Published online 2020 May 13. doi: 10.7554/eLife.54506.sa1

Decision letter

Axel T Brunger, Reviewing Editor
Axel T Brunger, Stanford University, United States;
Copyright and License information PMC Disclaimer

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

Acceptance summary:

One of the most compelling new results reported in this work shows that the combination of complexin and synaptotagmin clamps much more than the simple sum of both their clamping functions. This observation may suggest that the two molecules (complexin and synaptotagmin) act cooperatively in their clamping function.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "Independent Yet Synergistic Roles of Synaptotagmin-1 and Complexin in Calcium Regulated Neuronal Exocytosis" for consideration by eLife. Your article has been reviewed by a Senior Editor, a Reviewing Editor, and three reviewers. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that current manuscript is not acceptable, but we encourage you to consider a resubmission after addressing the concerns raised by the reviewers.

While the reviewers and editors found the new reconstitution method and the data presented very interesting, there appears to be a large gap between the experiments and the model/interpretation presented. Further experiments and a substantial revision of the paper could improve the work and increase its impact.

One of the most compelling new results suggests that combination of complexin and synaptotagmin clamps much more than the simple sum of both their clamping functions. This observation may suggest that the two molecules (complexin and synaptotagmin) act cooperatively in their clamping function. For example, the tripartite interface observed in the crystal structures by Zhou et al., 2017, may produce this cooperativity, but there could be other explanations. Additional mutations should be tested to correlate the various interactions (observed in structures and by the author's previous experiments) with functional studies with their reconstituted assay. We note that while the DA and Q mutants have been tested extensively in different contexts (these mutants primarily interfere with calcium binding and formation of the primary interface, respectively) the F349A mutation is less clearly defined since it may also affect (positively or negatively) the tripartite interface (it is an integral part of largely hydrophobic interactions between Syt1 C2B and the SNARE complex through the tripartite interaction, please consult the crystal structure of this complex). Mutations should be designed that more precisely test the oligomeric Syt1 interactions and the tripartite interface.

Reviewer #1:The authors have used a pore-spanning lipid bilayer setup to study the functional interplay between small unilamellar vesicles (SUVs) bearing both the v-SNARE VAMP2 and synaptotagmin (Syt1) and target membranes bearing pre-formed t-SNAREs, with or without complexin (Cpx). Previously (Ramakrishnan, 2019), the authors showed that the presence of Syt1 on the VAMP2-bearing SUVs led to 'clamped' vesicles – that is, vesicles that docked and remained immobile, without fusing, for at least ten minutes. In Figure 4 of this manuscript, they reproduce the experiments shown in Figure 3 of the earlier manuscript, albeit with two additional Syt1 mutants. Most of the current manuscript, however, is founded on the new observation that using a five-fold higher – and more physiologically realistic – number of VAMP2 per SUV dramatically alters the results. Instead of Syt1 being sufficient for clamping, its presence now merely delays fusion modestly. Strikingly the addition of Cpx, which in the absence of Syt1 has no effect, leads to an increase in the number of docked vesicles, and >90% of them (as opposed to 20% without Cpx) are securely clamped. The clamp so formed is efficiently reversed by Ca2+.

This leads to a model in which some SNARE complexes are clamped by Syt1 and others are clamped by Cpx. Since, in these experiments, Syt1 is outnumbered by VAMP2 by about 3:1, it stands to reason that it can't do all of the needed clamping on its own. (In Ramakrishnan, 2019, there was twice as much Syt1 as VAMP2). When the authors washed out Cpx the vesicles, presumably using the excess SNAREs, fused; when the Cpx was washed out in the presence of a reagent that trapped the liberated SNAREs in an inactive state, fusion was not observed until the Syt1 clamp was released by Ca2+ addition.

The authors' results imply that Syt1 is somehow required for Cpx to function as a fusion clamp, since Cpx alone shows no discernable clamping activity. Their model to explain this is that Syt1 forms a ring-like oligomer that impedes full zippering of the other 'peripheral' SNARE complexes, giving Cpx the opportunity to bind and clamp them. The problem with this model is that the only experiment that might be considered to be a critical test – using the F349A mutant to destabilize the Syt1 oligomers – not only doesn't abrogate Cpx-mediated clamping, but renders Cpx clamping irreversible. I don't know how to make sense of this. More generally, since Syt1:Cpx 'synergy' is quite central to the manuscript, one could argue that the authors need to investigate further; for example, by testing Syt1 mutations that disrupt the tripartite complex.Reviewer #2:Ramakrishnan and colleagues follow up on a study of theirs published earlier this year (FEBS Letters 2019) examining synaptic vesicle (SV) clamping mechanisms using a minimal reconstituted liposome fusion system comprising SNAREs, Syt1, and Cpx1. The current results indicate separate clamping functions for Syt1 and Cpx1, and both are required to produce calcium-sensitive stably docked vesicles in the context of physiological VAMP2 and Syt1 copy numbers. Disruption of a binding interface between the tSNAREs and C2B prevents Syt1-mediated clamping but also reveals an apparently irreversible clamping function of Cpx1. I am generally enthusiastic about the work, but I do have some questions on data interpretation and the proposed model.

From a technical perspective, this is a nice study that further develops a powerful in vitro docking/fusion model sufficient for detailed exploration of Syt1/Cpx1/SNARE mechanisms. Several of the interpretations put forward by the authors could use some clarification. In particular, the conclusion that Cpx1 acts independently of Syt1 on a distinct set of peripheral SNAREpins to clamp fusion is emphasized throughout the manuscript but not strongly supported by the data. On its own, Cpx1 does not provide a stable (ie 10's of seconds to minutes time scale) clamping function in this assay. But mutations in the 'primary' C2B-tSNARE interface eliminate Syt1 clamping function while somehow permitting an unexpected and potent Cpx1 stable clamping function. Also, previous work has suggested two distinct binding interfaces between C2B and the tSNAREs: the primary and tripartite interfaces (Zhou et al., 2017). The tripartite interface involves Cpx1 and may provide an explanation for the stable clamping by Cpx1 when the primary C2B interface is disrupted. Since C2B domains may interact with SNAREpins in these two distinct interfaces, changing the VAMP2:Syt1 ratio may alter the relative fraction of primary vs tripartite binding interactions and provide another explanation for the dose-dependence of VAMP2 on sufficiency of Syt1 for clamping versus the requirement for Cpx1 in the face of excess VAMP2. These scenarios may not be consistent with the picture of a central set of SNAREpins harboring Syt1 surrounded by Cpx1-bound SNAREpins as promoted by the authors. And no direct evidence for these central/peripheral SNAREpin arrangements is provided here. Thus, such a strong emphasis on independence of Syt1 and Cpx1 mechanisms in the Abstract, Results section and Discussion section is not warranted in my opinion.

In the high VAMP2:Syt1 experiments, the authors elegantly demonstrated the reversible nature of the Cpx1 clamping function by washing out Cpx1 and competing away excess SNAREpins with soluble tSNARE complexes. Did the authors attempt to wash out Cpx1 in the Syt1(Q)+tSNARE(Q) experiments where Cpx1 appeared to drive an irreversible clamp? And given the possible contribution of the tripartite C2B-SNARE interaction, did the authors attempt to disrupt this interface with mutations published in the Zhou et al., 2017 study? I do not think additional experiments would be required for publication of the current study, but could help bolster the proposed model.

The authors emphasize the importance of physiological copy number for the VAMP2 and Syt1 used in this study. How important is it to have 25 copies of (outwardly facing) Syt1 per vesicle? Work from Jahn put the number at around 15 copies per glutamatergic SV and possibily even fewer for GABAergic SVs. Does the clamping efficiency gradually go down as the copy numbers are reduced or is there a minimal requirement for any clamping to be observed? Perhaps other sources of C2 domains could contribute in synapses, but I was anticipating some comment on this 25 copy number given the Jahn work.

Reviewer #3:This manuscript explores the functional interactions between the Ca2+ sensor, Synaptotagmin1 (Syt1) and Complexin (Cpx) in clamping of neuronal SNARE complexes preceding Ca2+ influx and rapid fusion. They use a version of an in vitro single molecule fusion assay, which uses v-SNARE containing SUVs, with Si/SiO2 chips with t-SNARE containing membranes spread across the holes. Here, the authors examine membrane fusion between VAMP2-SUVs w/ and w/o Syt1, +/- Cpx, with t-SNARE complex membranes. Using a series of different protein combinations and concentrations, and various Syt1 mutant proteins, the authors suggest that two types of clamped SUVs occur: a small central "core" that is clamped by Syt1, and the rest that are clamped by Cpx, but only when facilitated by Syt1. Furthermore, they suggest that addition of Ca2+ facilitates fusion of only the Syt1-clamped SUVs, and that Cpx-clamped SUVs are blocked from fusion. It is an interesting hypothesis, but not well supported by the data presented.

A substantial amount of the data in the paper is basically validating their assay under "physiological conditions" of ~70 VAMPs and ~25 Syt1s per SUV, +/- Cpx (2uM), with the number of t-SNARE complexes unknown. The authors build on previous data from their lab, as well as others, related to the idea that Syt1 and Cpx work both independently as well as synergistically. The logic of the conclusions is based on a number of assumptions based on previous work, especially with regard to oligomerization of Syt1, interactions with Cpx and SNAREs, and whether washing 40x only serves to displace Cpx. This is particularly an issue in assays that are quite sensitive to the amount of protein used, but the extent of complex formation is only inferred indirectly. The authors present a hypothesis to try and explain the complicated findings, yet it is unclear whether the data fully support such a hypothesis. One particular example is the finding in Figure 1 that SUVs with Cpx alone fuse relatively quickly, yet the conclusion from Figure 4 is that "in the absence of the Syt1 clamp, Cpx blocks SNARE assembly irreversibly".

For most experiments, only a representative curve is shown, or the data (late time point?) summarized in bar graphs, with little detail of kinetics or binding shown. In many cases, data are "normalized," but it is unclear what the data are normalized to. It is also unclear if contents mixing experiments are performed for each of the different assay conditions, or only a representative experiment was done. These issues make many of the experiments tricky to interpret, and it is quite possible that alternative hypotheses could explain the data.

Furthermore, there is a lack of sufficient discussion (and referencing) of a number of other studies in the field. The authors need to try to reconcile their data with that of other groups, in order for their new hypothesis of Syt1-Cpx to be at all convincing.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for submitting your article "Independent Yet Synergistic Roles of Synaptotagmin-1 and Complexin in Calcium Regulated Neuronal Exocytosis" for consideration by eLife. Your article has been reviewed by Vivek Malhotra as the Senior Editor, a Reviewing Editor (Axel Brunger), and three reviewers. The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission. Please aim to submit the revised version within two months, but we are happy to extend this timeframe if needed. We recognize that we live in difficult and unprecedented times, so we are prepared to hear from you about a longer than usual time period for revisions.

Summary:

We thank the authors for responding to our previous concerns and including additional experiments. The hypothesis that there are distinct roles for Cpx and Syt1 in clamping two populations of SNAREs on a single vesicle is interesting and novel. However, this revised manuscript will benefit from substantial further revisions, clarifications, and discussion of other models and explanations. Moreover, rigorous statistical analyses of the survival times are requested. Our chief concerns are centered on the proposed model, interpretations, and analysis of the data, and no new experiments are requested. Below we detail several revisions required by the reviewers.

Essential revisions:

1) We are not convinced that the interpretation of their data is consistent with regard to the 'independent' roles of Cpx and Syt1 as stated in the title. Cpx on its own cannot clamp vesicles whereas in the presence of even a defective version of Syt1 (either the oligomer mutation or the quintuple Syt1+quintuple SNAP-25 mutation), Cpx now permanently inhibits vesicles from fusing. Thus, Cpx still clearly depends on Syt1 in a way that is not consistent with either the oligomer assembly or the primary Syt1-SNARE interface. One is left with the puzzle of how precisely synaptotagmin is creating this delay. Whatever this kinetic effect is, Cpx is not functioning independently of Syt1. Please revise the Results section and Discussion section to clarify your hypothesis and to better define independence in this context.

2) The notion of "clamping" suggests that an underlying molecular mechanism involving the synaptic proteins that have been included in this study. Yet, the measurements are assessing the survival of docked vesicles which is not a direct measure of molecular clamping. While molecular clamping could indeed be a possible explanation for the observations, there could be other explanations as well that involve the interplay between molecules and membranes. For example, it is possible that synaptotagmin or SNARE-induced hemifusion diaphragm formation (a long-lived metastable state) could affect the time of survival. Another possibility is that the membrane itself is the conduit for synergism. For example, it is known that synaptotagmin bends membranes and complexin preferentially binds to curved membranes, and thus, the membrane may introduce apparent synergism or cooperativity between the proteins. Please revise the Abstract, Results section and Discussion section accordingly.

3) The discussion in the text leaves one with the impression that the Syt1 C2B LLQQ mutant has no effect in their assay. Actually, it does have a significant effect, see Figure 4—figure supplement 1 panel B – the effect is somewhere between the quintuple mutant and wildtype after calcium addition. Please discuss.

4) The "survival percentage" plots are not quantitative. Please provide bar charts with error bars (along with significance tests) of the survival percentages after some defined time period(s). Another suggestion is to provide survival statistics (such as Kaplan Meier estimators).

5) Please promote some of the supplemental figures (especially the supplements to Figure 4) to primary figures, to in order to avoid 'burying' some of the most important results.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Synergistic Roles of Synaptotagmin-1 and Complexin in Calcium Regulated Neuronal Exocytosis" for further consideration by eLife. Your revised article has been evaluated by Vivek Malhotra (Senior Editor) and a Reviewing Editor.

We thank the authors for addressing the concerns of the reviewers and reviewing editor. The manuscript has much improved. However, there is a remaining point on statistical analysis that needs to be addressed before final acceptance, as outlined below:

The authors still do not provide some statistical statement of significance in the data presented in Figure 3, Figure 4, and Figure 5. They do provide info on the Kaplan Meier estimator of survival probability, but the reader is left to assume that all the differences shown are significant. They are very likely to be significant because the effects are quite large, but one usually expects a statistical test with either a p value or confidence interval to bolster the claim that an experimental manipulation did or did not have an effect.

2020; 9: e54506.
Published online 2020 May 13. doi: 10.7554/eLife.54506.sa2

Author response

Copyright and License information PMC Disclaimer

[Editors’ note: The authors appealed the original decision. What follows is the authors’ response to the first round of review.]

While the reviewers and editors found the new reconstitution method and the data presented very interesting, there appears to be a large gap between the experiments and the model/interpretation presented. Further experiments and a substantial revision of the paper could improve the work and increase its impact.

One of the most compelling new results suggests that combination of complexin and synaptotagmin clamps much more than the simple sum of both their clamping functions. […] Mutations should be designed that more precisely test the oligomeric Syt1 interactions and the tripartite interface.

To assess if the SNARE-Cpx-Syt1 ‘tripartite’ interaction1 could explain the observed cooperative function of Syt1 and Cpx, we tested the effect of the previously described Syt1 mutations (L387Q/L394Q) that disrupt this tripartite interface1 in our reconstituted assay. The LLQQ mutation had very little to no effect on the Syt1 clamp or the Ca2+-triggered fusion under both the low-copy VAMP2 (without Cpx) and the normal VAMP2 (with 2 µM Cpx) conditions. This suggests that the tripartite interface likely plays an auxiliary role in establishing the Syt1 clamp and cannot explain the observed synergistic effect of Syt1 and Cpx in clamping SNARE-mediated fusion.

These data are now included in Figure 4 and Figure 4—figure supplement 1 of the revised manuscript. Additionally, we have now expanded the Discussion section as follows: “We find that the SNARE-Cpx-Syt1 ‘tripartite’ interface, which has been shown to be physiologically-relevant 1, is not absolutely required to produce the Syt1 clamp or for the Ca2+ activation process under reconstituted conditions (Figure 4). […] Nonetheless, our data demonstrates that Syt1 and Cpx, along with SNARE proteins, form the minimal protein machinery that is necessary and sufficient to establish rapid Ca2+-regulated exocytosis”.

With regards to the F349A mutation, we have previously shown that this mutation specifically disrupts Syt1 oligomerization without affecting other molecular properties, including overall SNARE binding7. However, its effect on the ‘primary’ or the ‘tripartite’ SNARE binding sites has not been resolved. Nonetheless, considering that the LLQQ mutation has no effect on Syt1 clamping or activation function, it is unlikely that the F349A mutation exerts its effect via the tripartite interface. We thus conclude that Syt1 oligomerization is a key element of the Syt1 clamp.

Reviewer #1:

The authors' results imply that Syt1 is somehow required for Cpx to function as a fusion clamp, since Cpx alone shows no discernable clamping activity. Their model to explain this is that Syt1 forms a ring-like oligomer that impedes full zippering of the other 'peripheral' SNARE complexes, giving Cpx the opportunity to bind and clamp them. The problem with this model is that the only experiment that might be considered to be a critical test – using the F349A mutant to destabilize the Syt1 oligomers – not only doesn't abrogate Cpx-mediated clamping, but renders Cpx clamping irreversible I don't know how to make sense of this.

Survival analysis shows that the Syt1Q (SNARE ‘primary’ interface) and Syt1F349A (oligomerization) mutants destabilize the Syt1 clamp and abrogate the formation of central SNAREpins but still introduce a meaningful delay in the overall fusion process. This kinetic delay is sufficient for Cpx to arrest the assembly of all available SNAREs. Thus, we observe an irreversible clamped state under these conditions. This implies that the kinetic delay introduced by Syt1, irrespective of the Syt1 clamp, is adequate for Cpx clamping function.

These data are now included in Figure 4—figure supplement 2 and we have revised the relevant Results section as follows: “We also tested the effect of the Syt1 mutants using vesicles containing physiological VAMP2 and Syt1 copy numbers in the presence of 2 µM Cpx. […] This denotes that in Syt1Q vesicles (and presumably in Syt1F349A vesicles), all SNAREpins are clamped by Cpx alone and become accessible to CDT block following the Cpx wash-out”

More generally, since Syt1:Cpx 'synergy' is quite central to the manuscript, one could argue that the authors need to investigate further; for example, by testing Syt1 mutations that disrupt the tripartite complex.

We have now tested and confirmed that the tripartite interaction cannot explain the Syt1/Cpx synergy. See question #1 for detailed response.

Reviewer #2:

On its own, Cpx1 does not provide a stable (ie 10's of seconds to minutes time scale) clamping function in this assay. But mutations in the 'primary' C2B-tSNARE interface eliminate Syt1 clamping function while somehow permitting an unexpected and potent Cpx1 stable clamping function.

We find that Syt1 even in the absence of a stable clamp (i.e. Syt1Q and Syt1F349A) introduces a meaningful delay in the SNARE-mediated fusion and this kinetic delay is sufficient/required for Cpx to act as a fusion clamp. See question #2 for detailed response.

The tripartite interface involves Cpx1 and may provide an explanation for the stable clamping by Cpx1 when the primary C2B interface is disrupted. Since C2B domains may interact with SNAREpins in these two distinct interfaces, changing the VAMP2:Syt1 ratio may alter the relative fraction of primary vs tripartite binding interactions and provide another explanation for the dose-dependence of VAMP2 on sufficiency of Syt1 for clamping versus the requirement for Cpx1 in the face of excess VAMP2. Results section, Discussion section.

In the high VAMP2:Syt1 experiments, the authors elegantly demonstrated the reversible nature of the Cpx1 clamping function by washing out Cpx1 and competing away excess SNAREpins with soluble tSNARE complexes. And given the possible contribution of the tripartite C2B-SNARE interaction, did the authors attempt to disrupt this interface with mutations published in the Zhou et al., 2017 study?

We have now tested and confirmed that the tripartite interface is not involved in establishing the Syt1 clamp or Ca2+-activation mechanisms. Hence, this cannot explain the observed co-operative function of Syt1 and Cpx. See question #1 for detailed response.

These scenarios may not be consistent with the picture of a central set of SNAREpins harboring Syt1 surrounded by Cpx1-bound SNAREpins as promoted by the authors. And no direct evidence for these central/peripheral SNAREpin arrangements is provided here. Thus, such a strong emphasis on independence of Syt1 and Cpx1 mechanisms in the Abstract, Results section, and Discussion section is not warranted in my opinion.

We find that (i) Syt1 and Cpx act synergistically to block spontaneous fusion (ii) Syt1 and Cpx likely act independently and produce molecularly distinct clamped SNAREpins; (iii) the kinetic delay introduced by Syt1, independent of that Syt1 clamp, is need for Cpx to irreversibly arrest SNARE assembly; and (iv) SNARE-Syt1-Cpx tripartite interface cannot explain the observed co-operative function of Syt1 and Cpx (Figure 4). We believe that the outlined central/peripheral SNAREpins associated with Syt1 and Cpx offers the best explanation for all observed data.

Did the authors attempt to wash out Cpx1 in the Syt1(Q)+tSNARE(Q) experiments where Cpx1 appeared to drive an irreversible clamp?

In light of reviewer’s comment, we have carried the Cpx washout experiments under Syt1Q + SNAREQ. We find that in the absence of CDT, the Syt1Q vesicle proceed to fuse spontaneously following the buffer wash. Inclusion of CDT (with Cpx washout) irreversibly blocks all fusion events, even in the presence of 1 mM Ca2+. This, taken together with data shown in Figure 4, indicates that (i) the Syt1 clamp and the associated central SNAREpins are needed for Ca2+-triggered fusion and (ii) Cpx irreversibly blocks SNARE-mediated fusion. These data are included as Figure 4—figure supplement 3 in the revised manuscript and discussed in the Results section.

8) The authors emphasize the importance of physiological copy number for the VAMP2 and Syt1 used in this study. How important is it to have 25 copies of (outwardly facing) Syt1 per vesicle? Work from Jahn put the number at around 15 copies per glutamatergic SV and possibily even fewer for GABAergic SVs. Does the clamping efficiency gradually go down as the copy numbers are reduced or is there a minimal requirement for any clamping to be observed? Perhaps other sources of C2 domains could contribute in synapses, but I was anticipating some comment on this 25 copy number given the Jahn work.

Based on the available literature (Takamori et al., 2014; Wilhelm et al., 2014), there are ~16-22 copies of Syt1 per synaptic vesicle. We chose reconstitution conditions to produce 20-25 copies of outward facing Syt1 as a reasonable approximation. We are currently systematically testing the importance of the Syt1 and VAMP2 copy number. Preliminary analysis indicates that Syt1 clamping and Ca2+-triggered fusion is related to the Syt1 copy number and the Syt1:VAMP2 ratio. This will be focus of a future manuscript and as such, we chose not to address this in the current report.

Reviewer #3:

[…] A substantial amount of the data in the paper is basically validating their assay under "physiological conditions" of ~70 VAMPs and ~25 Syt1s per SUV, +/- Cpx (2uM), with the number of t-SNARE complexes unknown. The authors build on previous data from their lab, as well as others, related to the idea that Syt1 and Cpx work both independently as well as synergistically. The logic of the conclusions is based on a number of assumptions based on previous work, especially with regard to oligomerization of Syt1, interactions with Cpx and SNAREs, and whether washing 40x only serves to displace Cpx. This is particularly an issue in assays that are quite sensitive to the amount of protein used, but the extent of complex formation is only inferred indirectly.

We made no a priori assumptions and systematically tested the effect of Cpx and Syt1 on SNAREmediated fusion. We have examined the role of well-known Syt1 molecular properties, namely ‘primary’ and ‘tripartite’ SNARE binding, oligomerization and Ca2+-binding, in relation to the Syt1 clamp and its reversal by Ca2+. We advance the ‘synergistic clamping’ model as it best explains all available data (both in vitro and physiology) and is consistent with well-established structural and biochemical properties of Syt1, Cpx and SNAREs. As we have noted, we have put forward the simplest hypothesis to explain our findings and are aware of possible variations/alterations of our model. Nonetheless, we are confident of our central findings that (i) Syt1 and Cpx act independently and produce molecularly-distinct clamped SNAREpins under a single vesicle and (ii) the Syt1-associated ‘central’ SNAREpins are critical and likely sufficient for Ca2+-evoked fast vesicular release.

Based on our reconstitution conditions, we estimate that there are ~15,000 freely diffusing t-SNARE complex per 5 µm suspended bilayer. Thus, t-SNARE concentration is not a limiting factor in these experiments. It is worth noting that the t-SNARE concentration on the pre-synaptic membrane is not well-defined and is predicted to be in excess and thus, freely available. As noted, in most experiments, we use physiological copy numbers of Syt1and VAMP2 per vesicles. We chose 2 µM of soluble Cpx in our assay, considering its affinity to pre-fusion SNARE complexes (Kd ~ 0.5 µM) and the typical concentration (2-5 µM) used in previous studies (Malsam et al., 2012; Diao et al., 2012; Lai et al., 2014). Hence, we strongly believe our reconstitution conditions are a reasonable approximation of physiological-conditions.

The authors present a hypothesis to try and explain the complicated findings, yet it is unclear whether the data fully support such a hypothesis. One particular example is the finding in Figure 1 that SUVs with Cpx alone fuse relatively quickly, yet the conclusion from Figure 4 is that "in the absence of the Syt1 clamp, Cpx blocks SNARE assembly irreversibly".

Closer inspection of our data reveals that the delay introduced by Syt1, independent of the Syt1 clamp, is required for Cpx to function as a fusion clamp. We have revised our results and discussion accordingly. See question #2 for detailed response.

For most experiments, only a representative curve is shown, or the data (late time point?) summarized in bar graphs, with little detail of kinetics or binding shown. In many cases, data are "normalized," but it is unclear what the data are normalized to. It is also unclear if contents mixing experiments are performed for each of the different assay conditions, or only a representative experiment was done.

All data shown, including the curves, correspond to the average data (± standard deviations) and not representative data. The number of independent trials and total vesicles imaged is included in every figure legend and we will include relevant raw data in the source file. In addition, we have included representative images and video for selected data for clarity and transparency.

We indeed show vesicle fusion kinetics under both basal and Ca2+ conditions (Figure 1B and 2C). We do not have single-molecule resolution in our current setup to track protein assembly kinetics. It is worth noting that the protein concentrations used in the reconstituted experiments are comparable to those used by other labs. Considering the binding affinities and the temporal resolution of the experiments, we believe that the protein assembly kinetics should little to no effect on the clamping and fusion properties measured in this study.

We use normalization, with the number of docked vesicles set to 100%, for easy comparison between different assay conditions (Figure 3) and Syt1 mutants (Figure 4). For example, as the number of docked vesicles vary between different mutants, normalization allows us to best illustrate the effect of a given mutation(s) on spontaneous fusion and Ca2+-triggered fusion. We will include raw data used in these calculations in the source file.

We carried the content release experiments for the Syt1WT and few chosen mutations to test and qualitatively verify the findings from the lipid mixing analysis. We used lipid-mixing data for all statistical analysis as the fluorescence properties of the Alexa647, which is far superior compared to Sulphorhodamine B, is best suited for our automated analysis. We thus chose to only include a representative content release data for illustrative purposes.

[Editors’ note: what follows is the authors’ response to the second round of review.]

Essential revisions:

1) We are not convinced that the interpretation of their data is consistent with regard to the 'independent' roles of Cpx and Syt1 as stated in the title. Cpx on its own cannot clamp vesicles whereas in the presence of even a defective version of Syt1 (either the oligomer mutation or the quintuple Syt1+quintuple SNAP-25 mutation), Cpx now permanently inhibits vesicles from fusing. Thus, Cpx still clearly depends on Syt1 in a way that is not consistent with either the oligomer assembly or the primary Syt1-SNARE interface. One is left with the puzzle of how precisely synaptotagmin is creating this delay. Whatever this kinetic effect is, Cpx is not functioning independently of Syt1. Please revise the Results section and Discussion section to clarify your hypothesis and to better define independence in this context.

The intent was to convey our central finding that Syt1 and Cpx play distinct roles in clamping different pools of SNAREpins. We recognize that the original title might be misleading. As such, we have removed the reference to the ‘independent’ role from the title and revised it to read: Synergistic Roles of Synaptotagmin-1 and Complexin in Calcium Regulated Neuronal Exocytosis. In addition, we have made minor edits to the Abstract and in the Results section Discussion section to clearly explicate our findings.

2) The notion of "clamping" suggests that an underlying molecular mechanism involving the synaptic proteins that have been included in this study. Yet, the measurements are assessing the survival of docked vesicles which is not a direct measure of molecular clamping. While molecular clamping could indeed be a possible explanation for the observations, there could be other explanations as well that involve the interplay between molecules and membranes. For example, it is possible that synaptotagmin or SNARE-induced hemifusion diaphragm formation (a long-lived metastable state) could affect the time of survival. Another possibility is that the membrane itself is the conduit for synergism. For example, it is known that synaptotagmin bends membranes and complexin preferentially binds to curved membranes, and thus, the membrane may introduce apparent synergism or cooperativity between the proteins. Please revise the Abstract, Results section and Discussion section accordingly.

In this study, we define ‘clamping as the survival of docked vesicles beyond the initial observation period. Nonetheless, we find compelling evidence towards the existence of a ‘molecular clamp’. For example, we observe the fusion clamp only in the presence of Syt1 and Cpx and the fusion clamp is altered by disruption of specific molecular properties of Syt1 (i.e. SNARE primary binding, oligomerization). It is quite possible other factors, including membranes, play a role in augmenting the observed synergistic effect of Syt1 and Cpx. This is a very interesting possibility, with wideranging implication and will be focus of our future work. As such, this is beyond the scope of the current work. We have now highlighted this possibility in the Discussion section and included the following statement: Moreover, lipid membranes may contribute to synergism or cooperativity between Syt1 and Cpx in both clamping the un-initiated fusion events and triggering rapid and synchronous fusion in response to Ca2+-influx.

Nevertheless, we wish to point out that we do not believe that synaptotagmin or SNARE-induced hemi-fusion diaphragm formation is a likely explanation for the observed clamped state. Hemi-fusion would result in partial-lipid mixing and this can be readily verified by reduction of the vesicle fluorescence (ATTO647-DOPE) and spreading of fluorescence in the outer ROI. We do NOT observe any change in vesicle fluorescence signal for the docked/un-fused vesicles. Thus, we can rule out this possibility. We have explicated this in the revised Materials and methods section as follows: We did not observe any change in ATTO-647-PE fluorescence for the vesicles that remain docked and un-fused during the observation period or post Ca2+-addition. Thus, we can rule out hemi-fusion diaphragm formation as a possible explanation for the observed ‘clamped’ or ‘un-fused’ state.

3) The discussion in the text leaves one with the impression that the Syt1 C2B LLQQ mutant has no effect in their assay. Actually, it does have a significant effect, see Figure 4—figure supplement 1 panel B – the effect is somewhere between the quintuple mutant and wildtype after calcium addition. Please discuss.

With regards to the LLQQ mutational analysis, we observe very little to no effect of LLQQ mutation on the fusion clamp under both the low-copy VAMP2 (without Cpx) and the normal VAMP2 (with 2 µM Cpx) conditions. Thus, we conclude that the tripartite site is not essential for the fusion clamp.

The Syt1 LLQQ mutant does have a small, but significant effect in calcium triggering in the presence of Cpx. However, this effect was observed only under physiological VAMP2 conditions (Figure 5), but not under low copy VAMP2 conditions (Data not shown). This does suggest that the Syt1-Cpx-SNARE tripartite interaction might be important for calcium triggering mechanism, but it is not conclusive. Indeed, we are investigating this in further detail, including testing the effect of Syt1LLQQ and Syt1 quintuple mutation under varying calcium concentrations and VAMP2/Syt1 ratio with higher temporal resolution.

We do agree that we had not adequately highlighted/discussed the effect of LLQQ mutation on calcium triggering of fusion in the current manuscript. In addition to moving the relevant data into the primary figures (Figure 5), we have revised the Results section as follows: “However, the clamped Syt1Q and Syt1F349A vesicles were insensitive to Ca2+ and did not fuse following Ca2+ (1 mM) addition as opposed to the rapid and synchronous fusion observed with the majority of the Syt1WT and Syt1LLQQ vesicles (Figure 5). Notably, a significant partial fraction (~25%) of the Syt1LLQQ vesicles remained un-fused even following Ca2+ addition (Figure 5). This suggests that while the Syt1- Cpx-SNARE tripartite interface is not essential for establishing the fusion clamp, it is likely important in the Ca2+-activation mechanism”.

In addition, we have updated the relevant section in the Discussion section as follows: “We have articulated the simplest hypothesis, considering discrete ‘central’ and ‘’peripheral’ SNAREpin associated with Syt1 and Cpx, respectively. […] The tripartite interface might become more relevant in the pre-synaptic terminals (Zhou et al. 2017) where ~ 30% of Syt1 is present in the plasma membrane (Wienisch and Klingauf 2006) and other Synaptotagmins could also participate in the tripartite interface (Rothman et al. 2017; Volynski and Krishnakumar 2018; Zhou et al. 2017)”.

4) The "survival percentage" plots are not quantitative. Please provide bar charts with error bars (along with significance tests) of the survival percentages after some defined time period(s). Another suggestion is to provide survival statistics (such as Kaplan Meier estimators).

We wish to clarify that all survival curves shown in the manuscript are based on Kaplan Meier estimate calculations and plotted as ‘survival percentage’. We have now made this explicit in the Materials and methods section of the revised manuscript.

As suggested, we have also included the survival statistics (i.e. Kaplan Meier estimators) at defined time-period periods post-docking. These are shown in Table 1, Table 2 and Table 3 in the revised manuscript.

In addition, we have now included expanded (1h) survival plots for vSUVs clamped by Syt1 and Cpx. This is shown in Figure 1—figure supplement 4 of the revised manuscript.

5) Please promote some of the supplemental figures (especially the supplements to Figure 4) to primary figures, to in order to avoid 'burying' some of the most important results.

As recommended, we have promoted Figure 4—figure supplement 1 to the primary Figure 5.

[Editors’ note: what follows is the authors’ response to the second round of review.]

The authors still do not provide some statistical statement of significance in the data presented in Figure 3, Figure 4, and Figure 5. They do provide info on the Kaplan Meier estimator of survival probability, but the reader is left to assume that all the differences shown are significant. They are very likely to be significant because the effects are quite large, but one usually expects a statistical test with either a p value or confidence interval to bolster the claim that an experimental manipulation did or did not have an effect.

We now submit a revised manuscript incorporating the statistical test (log-rank test) of the survival curves shown in Figure 1B, Figure 4A and Figure 5A as recommended by the reviewer.

The statistical significance (p-values) for the pairwise comparison of the survival curves are shown in Table 2, Table 4 and Table 6 of the revised manuscript.

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