Synaptotagmin-1 and complexin inhibit spontaneous vesicle fusion by masking PIP2, not by clamping SNARE assembly

Synaptotagmin-1 and complexin inhibit spontaneous vesicle fusion by masking PIP2, not by clamping SNARE assembly | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Synaptotagmin-1 and complexin inhibit spontaneous vesicle fusion by masking PIP2, not by clamping SNARE assembly Houda Yasmine Ali Moussa , Kyung Chul Shin , Yongsoo Park doi: https://doi.org/10.1101/2025.03.24.644869 Houda Yasmine Ali Moussa 1 Neurological Disorders Research Center, Qatar Biomedical Research Institute (QBRI), Hamad Bin Khalifa University (HBKU), Qatar Foundation , Doha, Qatar Find this author on Google Scholar Find this author on PubMed Search for this author on this site Kyung Chul Shin 1 Neurological Disorders Research Center, Qatar Biomedical Research Institute (QBRI), Hamad Bin Khalifa University (HBKU), Qatar Foundation , Doha, Qatar Find this author on Google Scholar Find this author on PubMed Search for this author on this site Yongsoo Park 1 Neurological Disorders Research Center, Qatar Biomedical Research Institute (QBRI), Hamad Bin Khalifa University (HBKU), Qatar Foundation , Doha, Qatar 2 College of Health and Life Sciences (CHLS), Hamad Bin Khalifa University (HBKU), Qatar Foundation , Doha, Qatar Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: ypark{at}hbku.edu.qa Abstract Full Text Info/History Metrics Preview PDF Abstract Vesicle fusion underlies neurotransmitter release, enabling cellular communication. The rapid kinetics of vesicle fusion, tight docking to the plasma membrane, and coordination by multiple SNARE complexes suggest that SNARE complexes are already pre-assembled before fusion. Synaptotagmin-1 (Syt-1) and complexin (CPLX) have been proposed to clamp SNARE assembly and arrest fusion. However, these models are largely based on studies conducted under low ionic strength and structural analyses utilizing SNARE–Syt-1 chimera conjugates. We propose phosphatidylinositol 4,5-bisphosphate (PIP 2 ) as a critical lipid catalyst that facilitates fusion through electrostatic dehydration. Here we show that neither the C2AB domain of Syt-1 nor CPLX-2 has clamping or inhibitory effect on SNARE assembly. Instead, the C2AB domain and CPLX-2 inhibit Ca 2+ -independent vesicle fusion by masking PIP 2 . Our data resolve the long-standing question of increased spontaneous neurotransmitter release in Syt-1 and CPLX knockout neurons, emphasizing PIP 2 as an electrostatic lipid catalyst for fusion. One Sentence Summary Syt-1 and complexin inhibit spontaneous fusion by masking PIP 2 . Introduction Vesicle fusion is a fundamental process to release neurotransmitters and hormones in neurons and neuroendocrine cells 1 . Different types of vesicles are specialized to store specific neurotransmitters: synaptic vesicles (SVs) predominantly contain classical neurotransmitters, whereas large dense-core vesicles (LDCVs) mainly store amines, neuropeptides, and hormones 2 , 3 . Vesicle fusion is orchestrated by soluble N -ethylmaleimide-sensitive factor attachment protein receptor (SNARE) proteins 4 , 5 and synaptotagmin-1 (Syt-1) serving as a Ca 2+ sensor to mediate Ca²⁺-dependent vesicle fusion 6 , 7 . Given the ultrafast timescale of vesicle fusion 8 , 9 , the tight docking of vesicles at a distance of 1∼5 nm from the plasma membrane 10 , 11 , and multiple SNARE complexes to coordinate fusion 12 , 13 , it is likely that the SNARE complex is partially pre-assembled before fusion in a vesicle docking state. While several models have been proposed to explain SNARE clamping mechanisms involving Syt-1 and complexin (CPLX), most models rely on data obtained under low ionic strength conditions and structural studies using SNARE–Syt-1 chimera conjugates, where the C2AB domain is artificially linked to SNAP-25 via a short linker 14 , 15 , thereby making the mechanisms of SNARE clamping controversial 6 . Furthermore, although SNARE assembly provides the energy required to overcome the fusion energy barrier, the fact that SNARE complexes are already pre-assembled in a tightly docked state raises important questions about how the energy barrier is effectively overcome to initiate vesicle fusion. We recently proposed a paradigm shift in understanding vesicle fusion mechanisms, highlighting phosphatidylinositol 4,5-bisphosphate (PIP 2 ) as a critical lipid catalyst for vesicle fusion 16 , 17 . PIP 2 facilitates fusion by lowering the hydration energy barrier when vesicles are tightly docked through partial SNARE assembly 16 , 17 . Masking of PIP 2 by the effector domain (ED) of myristoylated alanine-rich C-kinase substrate (MARCKS) arrests vesicle fusion, stabilizing the vesicles in a state of partial SNARE assembly and tight docking 16 . The vesicle fusion arrest can be reversed by Ca²⁺/calmodulin (CaM) and protein kinase C (PKC), which dissociate the MARCKS ED from membranes, thereby unmasking PIP 2 17 . Syt-1 interacts with PIP 2 primarily through the polybasic region 18 . CPLX also binds to PIP 2 -containing membranes, competing with Syt-1 for PIP 2 binding 19 . Here, we provide evidence that neither the C2AB domain of Syt-1 nor CPLX-2 has clamping or inhibitory effect on SNARE assembly. Notably, no interaction between the SNARE complex and the C2AB domain is observed under physiological ionic conditions. Instead, the C2AB domain and CPLX-2 inhibit Ca 2+ -independent vesicle fusion by masking PIP 2 . Our data address the longstanding question of why knockout of Syt-1 and CPLX leads to an increase in spontaneous neurotransmitter release, highlighting the role of PIP 2 as a lipid catalyst for fusion through electrostatic dehydration. Results No interaction between the SNARE complex and Syt-1 in physiological ionic conditions Interaction between the SNARE complex and Syt-1 is observed only under low ionic strength and is entirely disrupted under physiological ionic conditions, including the presence of Mg 2+ /ATP 20 . Electron paramagnetic resonance (EPR) spectroscopy demonstrates no detectable interaction between the SNARE complex and Syt-1 in the presence of ATP 21 . Similarly, experiments with PIP 2 -containing nanodiscs using FRET confirm that ATP-induced charge shielding significantly reduces this interaction 22 . However, potential unconventional interaction modes may remain undetected with labeling strategies used in EPR and FRET measurement. To further confirm no interaction between the SNARE complex and Syt-1 under physiological ionic conditions, we performed fluorescence anisotropy measurement. Anisotropy, which monitors the conformational flexibility of labeled proteins, provides a sensitive measure of interactions regardless of diverse binding modes 23 . The stabilized SNARE complex, comprising syntaxin-1A (residues 183–288), SNAP-25A (cysteine-free), and VAMP-2 fragment (residues 49–96), was reconstituted into liposomes (Lip.) composed of 60% PC, 15% PE, and 25% cholesterol; no anionic phospholipids included. Alexa Fluor 488-labeled VAMP-2 (residues 49–96) served as the fluorescent probe ( Fig. 1a ). Download figure Open in new tab Figure 1 No interaction between the SNARE complex and Syt-1. ( a,b ) Interaction of the SNARE complex with the C2AB domain of Syt-1 was monitored using anisotropy measurement. The stabilized SNARE complex, incorporating VAMP-2 (residues 49–96) labeled with Alexa Fluor 488 (Alexa 488-labeled VAMP-2), was reconstituted into liposomes (Lip.) that contained no PS/PIP 2 : 60% PC, 15% PE, and 25% Chol. The C2AB domain (5 µM) was added as indicated by the arrow. No interaction between the SNARE complex and the C2AB domain was detected under physiological ionic conditions: 1 mM MgCl 2 /3 mM ATP in 150 mM KCl. ( c,d ) Instead of the SNARE complex, full-length syntaxin-1A (residues 1–288) labeled with Alexa Fluor 594 at T197C (magenta) was incorporated into liposomes. ( e-g ) No C2AB domain−SNARE interaction even in the presence of complexin-2 (CPLX-2). Sequential addition of CPLX-2 and the C2AB domain was performed to monitor the interaction with the SNARE complex as described in a . Physiological ionic strength with 1 mM MgCl 2 /3 mM ATP in 150 mM KCl was used in e-g . Anisotropy was normalized as A/A 0 , where A 0 is the initial value of anisotropy. Data in b,d,g represent means ± SD of three to six independent experiments. The addition of the C2AB domain of Syt-1 increased anisotropy under low ionic strength conditions (50 mM KCl), confirming SNARE–Syt-1 interaction. However, this interaction was abolished in the presence of 1 mM MgCl 2 /3 mM ATP with 150 mM KCl, demonstrating that physiological ionic conditions disrupt the SNARE–Syt-1 interaction regardless of the different binding modes ( Fig. 1a,b ). Additionally, full-length syntaxin-1A (residues 1–288), labeled with Alexa Fluor 594 at T197C, was tested. The C2AB domain interacted with syntaxin-1A incorporated in liposomes under low ionic strength conditions (50 mM KCl), but this interaction was entirely disrupted under physiological ionic conditions (1 mM MgCl 2 /3 mM ATP with 150 mM KCl)( Fig. 1c,d ). Given that CPLX-2 binds the SNARE complex, we further examined the interaction between Syt-1 and the SNARE complex in the presence of CPLX-2. CPLX-2 robustly interacted with the SNARE complex even under physiological ionic conditions (1 mM MgCl 2 /3 mM ATP with 150 mM KCl). However, the C2AB domain showed no additional interaction with the SNARE complex in the presence of CPLX-2 ( Fig. 1e-g ), consistent with previous FRET-based findings 20 . Furthermore, CPLX-2 showed no binding affinity to full-length syntaxin-1A ( Supplementary Fig. 1 ). Altogether, we confirmed no interaction between the SNARE complex and Syt-1 in physiological ionic conditions, regardless of the binding modes. Download figure Open in new tab Supplementary Figure 1 No interaction of CPLX-2 with syntaxin-1A. Interaction between CPLX-2 and syntaxin-1A was monitored using fluorescence anisotropy measurement as described in Fig. 1c . Full-length syntaxin-1A (residues 1–288) labeled with Alexa Fluor 594 at T197C (magenta) was incorporated into liposomes that contained no PS/PIP 2 : 60% PC, 15% PE, and 25% Chol. CPLX-2 (5 µM) was added as indicated by the arrow. No interaction between CPLX-2 and syntaxin-1A was detected. 1 mM MgCl 2 /3 mM ATP in 150 mM KCl. Anisotropy was normalized as A/A 0 , where A 0 is the initial value of anisotropy. No clamping effect of Syt-1 and CPLX on SNARE assembly and fusion To investigate whether Syt-1 or CPLX-2 has a clamping effect on SNARE assembly, we used FRET measurements as previously described 24 . The cytoplasmic domain (CD) of VAMP-2 (residues 1–96) was labeled with Oregon Green as a donor dye, and SNAP-25A was labeled with Texas Red within a stabilized Q-SNARE complex as the acceptor dye ( Fig. 2a ). SNARE assembly was monitored by a decrease in donor fluorescence, which occurs as soluble VAMP-2 CD labeled with Oregon Green assembles with the Q-SNARE complex containing Texas Red-labeled SNAP-25A 25 . The Q-SNARE complex was incorporated in liposomes. As a control, preincubation of unlabeled VAMP-2 CD with the Q-SNARE complex effectively blocked SNARE assembly mediated by labeled VAMP-2 CD ( Fig. 2a ). The C2AB domain did not inhibit SNARE assembly, even under low ionic strength conditions where the C2AB domain interacts with SNAREs ( Fig. 2a, b ). Neither the C2AB domain nor CPLX-2 exhibited any clamping effect on SNARE assembly ( Fig. 2a, b ), despite their interaction with the SNARE complex. Download figure Open in new tab Figure 2 Syt-1 and CPLX-2 show no clamping effect on SNARE assembly and fusion. ( a,b ) The effect of Syt-1 and CPLX-2 on SNARE assembly was investigated using FRET. SNARE assembly was monitored by the cytoplasmic domain (CD) of VAMP-2 (residues 1–96) labeled with Oregon Green and SNAP-25A in a stabilized Q-SNARE complex labeled with Texas Red (refer to Methods for details). The VAMP-2 fragment in the SNARE complex was omitted in the diagram for clarity. The reaction was initiated by adding 40 nM labeled soluble VAMP-2 CD to liposomes containing labeled SNAP-25A. SNARE complex assembly led to quenching of donor fluorescence. Liposome lipid composition: 45% PC, 15% PE, 10% PS, 25% Chol, 4% PI, and 1% PIP 2 . Preincubation was performed for 5 min with either 5 µM C2AB domain, 5 µM CPLX-2, or 1 µM unlabeled VAMP-2 CD. Low ionic strength conditions (50 mM KCl, without Mg 2+ /ATP) were used exclusively for the C2AB experiments. FRET was normalized as F/F 0 , where F 0 represents the initial value of the donor fluorescence intensity. ( c-f ) In-vitro reconstitution of vesicle fusion using a lipid-mixing assay in the presence of C2AB ( c,d ) or CPLX-2 ( e,f ). Neither C2AB nor CPLX-2 has the clamping effect on vesicle fusion. Purified native LDCVs were fused with PM-liposomes that incorporated the Q-SNARE complex consisting of the full-length syntaxin-1A (residues 1–288) and SNAP-25A (no cysteine, cysteines are replaced by alanines). The lipid composition of liposomes was consistent with fig. 2a except the addition of labeled PE. Preincubation of PM-liposomes with soluble VAMP-2 CD specifically disrupted SNARE-dependent vesicle fusion. Physiological ionic strength with 1 mM MgCl 2 /3 mM ATP was used in all experiments, unless stated otherwise. Note that ATP removes contaminating Ca 2+ as a Ca 2+ chelator 30 and prevents nonspecific weak interactions between SNAREs and synaptotagmin-1 20 . Data in b,d,f are means ± SD from three independent experiments. Next, we tested whether Syt-1 and CPLX-2 inhibit basal vesicle fusion using purified native vesicles. Physiological vesicle fusion was reconstituted in vitro using native vesicles such as LDCVs and SVs under physiological ionic strength (1 mM MgCl 2 /3 mM ATP) 16 , 17 , 20 , 24 , 26 – 29 . ATP removes contaminating Ca 2+ as a Ca 2+ chelator 30 and disrupts nonspecific weak interactions between SNAREs and Syt-1 20 , as well as cis -interactions of Syt-1 with vesicle membranes 26 , 30 . Physiological ionic strength with 1 mM MgCl 2 /3 mM ATP is critical for reconstitution of vesicle fusion in a physiological context and therefore, MgCl 2 /ATP were used in all experiments, unless stated otherwise. Native vesicles readily fused with plasma membrane-mimicking liposomes (PM-liposomes, 1% PIP 2 included) containing the Q-SNARE 31 ( Fig. 2c -f ). As expected, neither the C2AB domain ( Fig. 2c,d ) nor CPLX-2 ( Fig. 2e,f ) showed the inhibitory effect on basal LDCV fusion, indicating that Syt-1 and CPLX-2 do not clamp or inhibit SNARE assembly and basal vesicle fusion. Masking PIP 2 by the C2AB domain of Syt-1 The data presented above demonstrate that neither Syt-1 nor CPLX-2 shows a clamping effect on SNARE assembly. Given that vesicle fusion can be arrested by masking PIP 2 in a docking state with partial SNARE assembly 16 , we investigated whether the C2AB domain of Syt-1 inhibits basal fusion by masking PIP 2 . The binding of the C2AB domain to PIP 2 -containing membrane was analyzed using FRET measurement ( Fig. 3 ). The C2AB domain (residues 97–421), labeled with Alexa Fluor 488 at residue S342C (donor dye), interacted with liposomes containing Rhodamine-PE (acceptor dye): 45% PC, 13.5% PE, 1.5% Rho-PE, 10% PS, 25% cholesterol, 4% PI, and 1% PIP 2 . Download figure Open in new tab Figure 3 C2AB domain masks PIP 2 in a dose-dependent manner. ( a-d ) The membrane binding of the C2AB domain was analyzed using FRET measurement. The C2AB domain of Syt-1 (residues 97–421) was labeled with Alexa Fluor 488 at S342C (donor dye, green dot), while liposomes incorporated Rhodamine (Rho)-PE (acceptor dye, red dot). Lipid composition for FRET: protein-free, 45% PC, 13.5% PE, 1.5% Rho-PE, 10% PS, 25% Chol, 4% PI, and 1% PIP 2 . To vary PIP 2 concentrations, PI content was adjusted accordingly. ( a,b ) Increasing PIP 2 content up to 5% induced Ca 2+ -independent C2AB membrane binding, whereas 1% PIP 2 showed little effect. ( c,d ) The KAKA mutant of the C2AB domain reduced Ca 2+ -independent membrane binding. C2AB binding in b,d was normalized as F 0 /F, where F 0 is the initial value of the donor fluorescence intensity. All experiments were conducted under physiological ionic conditions (1 mM MgCl₂ and 3 mM ATP) in the absence of Ca 2+ . Data in b,d are means ± SD from three independent experiments. Unpaired two-tailed t -test was used ( d ). The C2AB domain exhibited Ca 2+ -independent binding to 1% PIP 2 liposomes, but this interaction was disrupted by 1 mM MgCl 2 /3 mM ATP due to charge shielding ( Supplementary Fig. 2 ), as reported before 20 . As in a vesicle fusion assay, MgCl 2 /ATP were used to removes contaminating Ca 2+ 30 and disrupt nonspecific interactions of the C2AB domain with membranes. Notably, increasing PIP 2 concentrations to 5% induced robust Ca 2+ -independent binding in a dose-dependent manner ( Fig. 3a,b ). Furthermore, the KAKA mutant of the C2AB domain, in which lysine residues were replaced with alanine (K326A, K327A) 32 , 33 , showed significantly reduced binding ( Fig. 3c,d ), highlighting the role of the polybasic region in mediating PIP 2 -masking. Download figure Open in new tab Supplementary Figure 2 ATP completely disrupts Ca 2+ -independent membrane binding of the C2AB domain. The membrane binding of the C2AB domain was analyzed using FRET measurement as described in Fig. 3 . The C2AB domain of Syt-1 (residues 97–421) was labeled with Alexa Fluor 488 at S342C (donor dye, green dot), while liposomes incorporated Rhodamine (Rho)-PE (acceptor dye, red dot). Lipid composition for FRET: protein-free, 45% PC, 13.5% PE, 1.5% Rho-PE, 10% PS, 25% Chol, 4% PI, and 1% PIP 2 . Treatment with 1 mM MgCl 2 and 3 mM ATP completely disrupted Ca 2+ -independent C2AB binding to liposomes containing 1% PIP 2 . As a control, 100 µM Ca 2+ induced C2AB membrane binding, which was reversed by 1 mM EGTA. Physiological ionic conditions (1 mM MgCl 2 and 3 mM ATP) abolished C2AB membrane binding to liposomes that contain 1% PIP 2 . Ca 2+ -independent C2AB membrane binding was observed when membranes contained 3∼5% PIP 2 , as shown in Fig. 3a ,b . Inhibition of Ca 2+ -independent vesicle fusion by masking PIP 2 Next, we investigated whether Syt-1 and CPLX-2 inhibit basal LDCV fusion using a lipid-mixing assay. Neither Syt-1 nor CPLX-2 inhibited vesicle fusion when PM-liposomes contained 1% PIP 2 ( Fig. 2c-f ), consistent with the absence of C2AB binding to membranes containing 1% PIP 2 in the presence of Mg 2+ /ATP ( Fig. 3a,b ). To evaluate the robust PIP 2 masking by the C2AB domain, we increased PIP 2 concentration in PM-liposomes to 5%. Under these conditions, the C2AB domain of Syt-1 inhibited basal LDCV fusion in a dose-dependent manner ( Fig. 4a ). Since CPLX-2 also has PIP 2 -binding affinity 19 , we tested CPLX-2 to reduce basal fusion by masking PIP 2 . As expected, CPLX-2 inhibited Ca 2+ -independent vesicle fusion in a dose-dependent manner, when PM-liposomes contained 5% PIP 2 ( Fig. 4b ). However, the C2AB domain demonstrated approximately threefold higher potency compared to CPLX-2, with IC 50 of 0.89 µM for the C2AB domain and 2.99 µM for CPLX-2 ( Fig. 4c ). At the maximum concentration, the C2AB domain and CPLX-2 inhibit basal fusion by ∼50%, whereas MARCKS ED completely blocks basal fusion 16 . Download figure Open in new tab Figure 4 Syt-1 and CPLX-2 inhibit Ca 2+ -independent vesicle fusion by masking PIP 2 . LDCV fusion with PM-liposomes using a lipid-mixing assay. PM-liposomes contained 5% PIP 2 and incorporated the stabilized Q-SNARE complex consisting of syntaxin-1A and SNAP-25A in a 1:1 molar ratio by the C-terminal VAMP-2 fragment (residues 49–96). The C2AB domain of Syt-1 ( a ) and CPLX-2 ( b ) inhibited Ca 2+ -independent vesicle fusion in a dose-dependent manner. ( c ) The inhibitory effects were quantified, with IC 50 as 0.89 µM for C2AB and 2.99 µM for CPLX-2. ( d,e ) The KAKA mutant of the C2AB domain showed reduced inhibitory effect on vesicle fusion. ( f ) Co-incubation of 1 µM C2AB domain and 1 µM CPLX-2 did not have additional inhibitory effects on basal fusion. Physiological ionic strength with 1 mM MgCl 2 /3 mM ATP was used. Data in c,e,f are means ± SD from three to four independent experiments. One-way ANOVA test with Bonferroni’s post-hoc correction was used. Further experiments with the KAKA mutant of the C2AB domain, which replaces lysine residues with alanine, revealed reduced inhibition of vesicle fusion compared to the wild-type C2AB domain ( Fig. 4d, e ), supporting the importance of the polybasic region of the C2AB domain in PIP 2 -masking to block basal fusion. Additionally, no significant synergistic effect on basal fusion inhibition was observed when 1 µM of the C2AB domain and 1 µM of CPLX-2 were combined ( Fig. 4f ). Taken together, Syt-1 and CPLX-2 reduce Ca 2+ -independent vesicle fusion by masking PIP 2 , not by clamping SNARE assembly. PIP 2 is a lipid catalyst to trigger vesicle fusion through electrostatic dehydration, and PIP 2 -masking by Syt-1 and CPLX-2 inhibits basal fusion. No effect of Syt-1 and CPLX-2 on SNARE disassembly Alpha- s oluble N SF a ttachment p rotein (α-SNAP) binds to the Q-SNARE complexes and interferes with full SNARE zippering, thus slowing down fusion and causing tight docking as a result of partially assembled SNARE proteins 24 . The SNARE complex is disassembled by the AAA+ ATPase N -ethylmaleimide-sensitive factor (NSF) and a cofactor, alpha-SNAP. Alpha-SNAP first binds to the SNARE complex with a high binding affinity, and then recruits NSF, thereby catalyzing SNARE disassembly in an ATP-dependent manner 34 . In the final set of experiments, we tested whether Syt-1 and CPLX-2 can compete with alpha-SNAP for SNARE disassembly. SNARE disassembly was monitored using a FRET-based assay, as described earlier 24 , 35 . Soluble VAMP-2 CD labeled with Oregon Green was assembled with the Q-SNARE complex containing Texas Red-labeled SNAP-25A to form the ternary SNARE complex on liposomes 25 , as in Fig. 2a . Alpha-SNAP (500 nM) and NSF (60 nM) were then added in the presence of MgCl 2 and ATP to activate SNARE disassembly, leading to dissociation of labeled proteins ( Fig. 5a,b ). Disassembly of the SNARE complex was detected by an increase in donor fluorescence intensity. Preincubation with 5 µM of either the C2AB domain of Syt-1 or CPLX-2 did not alter the disassembly of the SNARE complex mediated by alpha-SNAP and NSF ( Fig. 5a,b ). CPLX-2 initially slowed the disassembly kinetics of SNARE complexes, but this effect was rescued within 4 minutes ( Fig. 5a ), because alpha-SNAP binds with higher affinity to membrane-bound SNARE complexes 35 . Download figure Open in new tab Figure 5 Syt-1 and CPLX-2 do not affect SNARE disassembly. ( a,b ) SNARE disassembly was monitored using FRET measurement. VAMP-2 CD was labeled with Oregon Green and SNAP-25A in the Q-SNARE complex was labeled with Texas Red. The four-helix ternary SNARE complex was formed by SNARE assembly, as described in Fig. 2a . ( a ) SNARE disassembly was measured as an increase in donor fluorescence intensity, corresponding to the disassembly of the SNARE complex. Alpha-SNAP (500 nM) and NSF (60 nM) were added as indicated by the arrow. Preincubation of the SNARE complex with 5 µM C2AB domain or 5 µM CPLX-2 for 5 min showed no inhibition of SNARE disassembly. Liposome lipid composition: 45% PC, 15% PE, 10% PS, 25% Chol, 4% PI, and 1% PIP 2 . ( b ) SNARE disassembly is presented as a percentage of control. 3 mM MgCl 2 and 3 mM ATP were included in buffer to activate NSF-mediated disassembly. Data are means ± SD from three independent experiments. ( c,d ) Monitoring SNARE disassembly on LDCV membranes using fluorescence anisotropy. LDCVs were incubated with the soluble Q-SNARE complexes (50 nM) comprising syntaxin-1A (residues 183–262), SNAP-25A, and VAMP-2 fragment (residues 49–96), leading to the assembly of membrane-anchored ternary SNARE complexes. Syntaxin-1A (residues 183–262) in the soluble Q-SNARE complex was labeled with Alexa Fluor 488 at position Cys225. Endogenous VAMP-2 from LDCVs replaced VAMP-2 fragment (residues 49–96) to form the ternary SNARE complex. Sequential additions of LDCVs, alpha-SNAP (500 nM), and NSF (60 nM) resulted in stepwise increases in fluorescence anisotropy signals. SNARE disassembly triggered by MgCl 2 led to dissociation of syntaxin-1A, observed as a decrease in anisotropy signals. 3 mM ATP was included in buffer and as indicated by the arrow, 3 mM MgCl 2 was applied to activate NSF-mediated SNARE disassembly. ( d ) Co-incubation of 5 µM C2AB domain and 5 µM CPLX-2 did not alter SNARE disassembly. To validate these findings, we employed an alternative way of monitoring SNARE disassembly, i.e., fluorescence anisotropy. Endogenous VAMP-2 from LDCVs was assembled with the soluble Q-SNARE complexes (50 nM) to form membrane-anchored ternary SNARE complexes on LDCV membranes ( Fig. 5c,d ). Syntaxin-1A (residues 183–262) in the soluble Q-SNARE complex was labeled with Alexa Fluor 488 at position Cys225. SNARE assembly increased fluorescence anisotropy, confirming the formation of the ternary SNARE complex, as reported previously 24 . The SNARE complex formation on LDCV membranes was further validated using SDS-PAGE ( Supplementary Fig. 3 ). Download figure Open in new tab Supplementary Figure 3 SNARE disassembly mediated by alpha-SNAP and NSF. SNARE disassembly was assessed by analyzing the ternary SNARE complex on SDS-PAGE gels. LDCVs were incubated with the soluble Q-SNARE complexes (50 nM) comprising syntaxin-1A (residues 183–262), SNAP-25A, and VAMP-2 (residues 49–96), resulting in the formation of membrane-anchored ternary SNARE complexes on LDCVs. Endogenous VAMP-2 from LDCVs participated in the formation of ternary SNARE complexes. The light chain of tetanus neurotoxin (TeNT), a protease, selectively cleaves free VAMP-2. VAMP-2 incorporated into the SNARE complex was resistant to TeNT cleavage, migrating as a higher molecular weight band on SDS-PAGE gels, confirming SNARE assembly. The addition of alpha-SNAP (500 nM), NSF (60 nM), 3 mM ATP, and 3 mM MgCl 2 induced disassembly of the ternary SNARE complex. Degradation of endogenous VAMP-2 by TeNT confirmed SNARE disassembly. Samples were not boiled to preserve the SNARE complex. Sequential additions of alpha-SNAP (500 nM) and NSF (60 nM) induced additional stepwise increases in fluorescence anisotropy, reflecting protein binding to the ternary SNARE complex ( Fig. 5c ). Activation of NSF-driven disassembly was initiated by adding MgCl 2 , which stabilizes ATP hydrolysis 24 . Co-incubation of CPLX-2 and the C2AB domain had no effect on SNARE disassembly ( Fig. 5d ), indicating that the high affinity of alpha-SNAP for the SNARE complex effectively outcompete Syt-1 and CPLX-2, when the SNARE complex is incorporated in membranes 35 . Altogether, Syt-1 and CPLX-2 do not influence SNARE assembly or disassembly, even under conditions where they interact with SNARE proteins. Discussion We propose a paradigm shift for the molecular mechanisms of vesicle fusion, emphasizing the role of lipids as catalysts in facilitating this process. PIP 2 and cholesterol are critical plasma membrane lipids required for vesicle fusion ( Fig. 6a ). PIP 2 acts as a lipid catalyst for vesicle fusion through electrostatic dehydration 16 , 17 . Cholesterol catalyzes Ca 2+ -dependent vesicle fusion by strengthening Syt-1-induced membrane bending 25 , 31 . Vesicle docking is initiated by SNARE assembly, which brings vesicle and plasma membranes into close proximity ( Fig. 6b ). Syt-1 and CPLX-2 inhibit Ca 2+ -independent basal fusion by masking PIP 2 , thereby arresting fusion in a docking state. Notably, neither Syt-1 nor CPLX-2 clamps SNARE assembly. PIP 2 -masking arrests fusion due to the hydration energy barriers 16 . PIP 2 is a lipid catalyst, because PIP 2 i) attracts cations that neutralize electrostatic repulsion between membranes and ii) repel water molecules, reducing hydration energy and facilitating membrane fusion ( Fig. 6b ). Upon Ca 2+ influx, Syt-1 is inserted into the plasma membrane, inducing membrane deformation and bending. Cholesterol plays a critical role in lowering the energy barrier for fusion by stabilizing Syt-1-induced membrane deformation ( Fig. 6c ). The final step involves the formation of a fusion pore, enabling the release of neurotransmitters and hormones ( Fig. 6d ). This model highlights the catalytic roles of lipids in driving vesicle fusion and neurotransmitter release. Download figure Open in new tab Figure 6 Schematic overview of Syt-1 and CPLX to inhibit Ca 2+ -independent basal fusion. ( a ) PIP 2 and cholesterol are essential lipids of the plasma membrane for vesicle fusion. PS is present both in vesicle and plasma membrane. ( b ) Vesicle docking is initiated by SNARE assembly, which brings the vesicle and plasma membranes into close proximity. PIP 2 acts as a lipid catalyst for vesicle fusion by inducing electrostatic dehydration. PIP 2 attracts cations that i) neutralize the electrostatic repulsion between the membranes and ii) repel water molecules, reducing hydration energy and facilitating membrane merger. Syt-1 masks PIP 2 and arrest Ca 2+ -independent basal fusion in a state of vesicle docking. CPLX interacts with PIP 2 on the plasma membrane, similarly inhibiting basal fusion, but without clamping SNARE assembly. ( c ) Syt-1 is inserted to the plasma membrane in response to high Ca 2+ . PIP 2 induces the trans -interaction of synaptotagmin-1 with the plasma membrane. Syt-1 mediates evoked vesicle fusion through membrane deformation and bending, which are strengthened by cholesterol. ( d ) The formation of the fusion pore allows the release of neurotransmitters and hormones. The SNARE clamping hypothesis was proposed to explain the observed increase in spontaneous neurotransmitter release (mini-frequency) upon deletion of Syt-1 across various cell types 36 – 42 . Several models suggest that Syt-1 and CPLX act as clamping factors to arrest vesicle fusion, yet these models largely rely on data obtained under low ionic strength conditions, which may not accurately represent physiological environments 6 . Consequently, the mechanisms underlying SNARE clamping remain controversial 6 . For instance, while atomic-resolution X-ray crystallography suggests that Syt-1 interacts with the SNARE complex through a primary interface (reviewed in Ref. 7 ), solution NMR studies indicate that the SNARE–Syt-1 interaction is mediated by the polybasic region 43 . These contradictory findings raise further questions about the true nature of the SNARE–Syt-1 interaction. Corroborating earlier reports 20 , we demonstrate that no interaction occurs between Syt-1 and the SNARE complex under physiological ionic conditions, including Mg 2+ /ATP ( Fig. 1a,b ). Using fluorescence anisotropy, we rule out any unconventional SNARE–Syt-1 interactions, regardless of CPLX-2 ( Fig. 1 ), arguing against the SNARE clamping model. EPR spectroscopy provides strong evidence of no detectable interaction between the SNARE complex and Syt-1 in the presence of ATP 21 , consistent with our observations. Similarly, FRET experiments using nanodiscs reveal that ATP-induced charge shielding dramatically reduces SNARE–Syt-1 interaction 22 . The K d of the C2AB domain for PIP 2 -containing nanodiscs is 4.9 ± 1.1 µM, while nanodiscs containing the SNARE complex lower the K d to 0.37 µM, suggesting a robust interaction between the SNARE complex and Syt-1 22 . However, treatment with Mg 2+ /ATP reverses this interaction, increasing the K d to 3.3 µM. Additionally, Mg 2+ /ATP completely disrupts the SNARE–Syt-1 interaction in the presence of 1 mM Ca 2+22 . Our fluorescence anisotropy data further validate no SNARE–Syt-1 interaction under physiological ionic conditions ( Fig. 1 ), reinforcing the conclusion that the SNARE–Syt-1 interaction is not physiologically relevant. We highlighted several factors that lead to experimental artifacts in studies of Syt-1 6 . First, the use of chimera protein complexes in the studies of NMR and X-ray crystallography, where the C2AB domain is artificially linked to SNAP-25 via a short linker 14 , 15 , can introduce artifacts that mislead data interpretation. To avoid such issues, the soluble C2AB domain should be used to verify SNARE–Syt-1 interactions in structural studies. Second, PIP 2 should be excluded from membranes, such as nanodiscs, when confirming SNARE–Syt-1 interactions, as the C2AB domain strongly binds to PIP 2 -containing membranes, potentially confounding results. Third, protein structure studies and functional assay should be conducted under physiological ionic conditions, incorporating Mg 2+ /ATP. Syt-1 interactions with membranes or SNARE proteins are tightly regulated by electrostatic effects, making its function and activity highly dependent on ionic strength and salt concentration. Addressing these factors is essential to accurately understand Syt-1 function and interactions under physiological conditions. Electrophysiology data support our model that Syt-1 inhibits spontaneous vesicle fusion by masking PIP 2 . Substitution of three lysine residues in the polybasic region of the C2B domain with glutamines (K379,380,384Q) increases spontaneous release approximately twofold at the Drosophila neuromuscular junction (NMJ) 44 . Similarly, deletion of basic residues (R398,399Q or K326,327E) elevates miniature release frequency and spontaneous fusion in mouse neurons, further supporting the role of PIP 2 -masking in inhibiting basal vesicle fusion 45 , 46 . Under physiological ionic conditions, the polybasic region of the C2B domain plays a key role in PIP 2 interaction 18 . Our data support that the KAKA mutation reduces PIP 2 masking ( Fig. 3c, d ), which correlates with a diminished inhibitory effect on basal fusion ( Fig. 4d, e ). Additionally, R398 and R399 of the C2B domain bind to PIP 2 -containing membranes 43 , and the R398Q,R399Q mutation impairs membrane binding without interfering with the C2AB–SNARE interaction 47 . These findings emphasize the importance of the polybasic region in regulating spontaneous release by masking PIP 2 . SNARE assembly is essential for vesicle fusion by bringing vesicles into close proximity with the plasma membrane in a docking state, however, our data challenge the hypothesis that SNARE proteins serve as the primary fusion machinery by providing assembly-driven energy to overcome the barriers for vesicle fusion. First, the fact that SNARE complexes are already pre-assembled in a tightly docked state prior to fusion raises critical questions about how the energy barrier is effectively overcome to facilitate membrane merger. Second, Syt-1 and CPLX have no clamping effect on SNARE assembly, and SNARE zippering and vesicle fusion are rarely arrested by SNARE-interacting proteins 16 , 24 , criticizing the SNARE clamp hypothesis. Third, PIP 2 -masking arrests vesicle fusion in a tightly docked state despite partial SNARE assembly, challenging SNAREs as fusion machinery. Altogether, SNARE assembly is not sufficient to overcome the energy barriers required for fusion. We propose a paradigm shift, identifying PIP 2 as a lipid catalyst for fusion by lowering the hydration energy barrier. Competing Interests The authors have no competing interests. Data availability The datasets that support the findings of this study are openly available. Author Contributions Conceptualization, Y.P.; Methodology, Software, Validation, Formal analysis, Investigation, Visualization, Y.P., H.Y.A.M., K.C.S.; Supervision, Y.P.; Project administration and Funding acquisition, Y.P.; Original draft preparation, Y.P.; Review and editing, H.Y.A.M., K.C.S.; All authors have read and agreed to the published version of the manuscript. Online Methods Materials ATP disodium salt was from Sigma-Aldrich (Cat. A2383). Oregon Green 488 maleimide (Cat. O6034), Texas Red C2-maleimide (Cat. T6008), Alexa Fluor 594 C5 maleimide (Cat. A10256) and Alexa Fluor 488 C5 maleimide (Cat. A10254) were purchased from Thermo Fisher Scientific. All lipids were purchased from Avanti Polar lipids. Antibody against VAMP-2 (Cat. 104 318, clone number 69.1) was from Synaptic Systems (Göttingen, Germany). Purification of large dense-core vesicles (LDCVs) LDCVs, also known as chromaffin granules, were purified from bovine adrenal medullae by using continuous sucrose gradient and resuspended with fusion buffer containing 120 mM K-glutamate, 20 mM K-acetate, and 20 mM HEPES.KOH, pH 7.4, as described previously 28 . Briefly, fresh bovine adrenal glands were obtained from a local slaughterhouse. The cortex and fat were removed, then the medullae were minced with scissors in 300 mM sucrose buffer (300 mM sucrose, 20 mM HEPES, pH 7.4 adjusted with KOH), and homogenized using a homogenizer. After centrifugation at 1,000g for 15 min at 4 °C, the pellet containing nuclei and cell debris (P1) was discarded. The supernatant (S1) was further centrifuged (12,000g, 15 min, 4 °C), then subjected to an additional cycle of resuspension and centrifugation as a washing step. The resulting pellet (P2, crude LDCV fraction) was resuspended in 300 mM sucrose buffer and loaded on top of a continuous sucrose gradient (from 300 mM to 1.9 M) to remove other contaminants including mitochondria. LDCVs were collected from the pellet after centrifugation at 110,000g for 60 min in a Beckman SW 41 Ti rotor, then resuspended with the buffer (120 mM K-glutamate, 20 mM K-acetate, 20 mM HEPES.KOH, pH 7.4). Protein purification All SNARE, Syt-1, and CPLX-2 constructs based on Rattus norvegicus sequences were expressed in E. coli strain BL21 (DE3) and purified by Ni 2+ -NTA affinity chromatography followed by ion-exchange chromatography as described elsewhere 20 , 24 , 26 , 31 , 32 , 48 , 49 . The stabilized Q-SNARE complex was composed of syntaxin-1A (residues 183–288) and SNAP-25A (no cysteine, cysteines replaced by alanines) in a 1:1 molar ratio by the C-terminal VAMP-2 fragment (residues 49–96) and the soluble Q-SNARE complex with syntaxin-1A (residues 183–262) were purified as described earlier 31 . The soluble cytoplasmic domain (CD) of stabilized Q-SNARE complex with syntaxin-1A (residues 183–262) was purified as described earlier 31 . The binary Q-SNARE complex containing the full-length syntaxin-1A (residues 1–288) and SNAP-25A (no cysteine, cysteines replaced by alanines) was expressed using co-transformation 26 . Soluble cytoplasmic region of VAMP-2 (residues 1–96) and the C2AB fragment of synaptotagmin-1 (residues 97–421), and the KAKA mutant (K326A, K327A) were purified by Mono S column (GE Healthcare, Piscataway, NJ) as described previously 32 , 49 . The stabilized Q-SNARE complexes were purified by Ni 2+ -NTA affinity chromatography followed by ion-exchange chromatography on a Mono Q column (GE Healthcare, Piscataway, NJ) in the presence of 50 mM n-octyl-β-D-glucoside (OG) 26 . Soluble VAMP-2 lacking the transmembrane domain called VAMP-2 CD (residues 1–96), and C2AB domain of synaptotagmin-1 (residues 97–421) were purified by Mono S column 24 . Chinese hamster NSF and bovine α-SNAP (wild-type) were expressed and purified as described in detail elsewhere 24 , 35 . CPLX-2 (residues 1–134) was purified as explained previously 48 . Protein structures were visualized with PyMOL; PDB 1BYN for the C2A domain, 1K5W for the C2B domain, 3IPD for the SNARE complex, 3C98 for syntaxin-1A. Protein labeling The point-mutated C2AB domain (S342C)(C2AB-Alexa 488) 32 , KAKA mutant of the C2AB domain 32 , and VAMP-2 (residues 49–96, T79C) in the stabilized Q-SNARE complex 26 , 31 were labeled with Alexa Fluor 488 C5 maleimide. Full-length syntaxin-1A (1–288) monomer was labeled with Alexa Fluor 594 C5 maleimide at T197C 20 . The ternary SNARE complex consisting of syntaxin-1A (183– 288), labeled SNAP-25A (Cys130), and VAMP-2 CD (49-96) was purified on Mono Q column (GE Healthcare, Piscataway, NJ). Syntaxin-1A (residues 183–262) in the soluble Q-SNARE complex was labeled with Alexa Fluor 488 C5 maleimide at position Cys225 24 . A single cysteine SNAP-25A mutant (Cys130) in the stabilized Q-SNARE complex was labeled with Texas Red C2-maleimide 20 , 24 . VAMP-2 CD (Cys28) was labeled with Oregon Green 488 maleimide 24 . For fluorescent labeling, proteins were incubated with 10-fold molar excess of the fluorophores for 2 hr and unbound dye was removed by gel filtration on a PD-10 column. Protein labeling efficiencies were >70∼80% as assessed by UV-visible spectroscopy. Lipid composition of liposomes All lipids were from Avanti Polar lipids, unless stated otherwise. Lipid composition (molar percentages) of PM-liposomes containing the Q-SNARE complex: 45% PC (L-α-phosphatidylcholine, Cat. 840055), 15% PE (L-α-phosphatidylethanolamine, Cat. 840026), 10% PS (L-α-phosphatidylserine, Cat. 840032), 25% Chol (cholesterol, Cat. 700000), 4% PI (L-α-phosphatidylinositol, Cat. 840042), and 1% PIP 2 (Cat. 840046). In case of removing PS/PIP 2 , PC contents were accordingly adjusted. For vesicle fusion lipid-mixing assays, 1.5% 1,2-dioleoyl- sn -glycero-3-phosphoethanolamine-N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) (NBD-PE, Cat. 810145) as a donor and 1.5% 1,2-dioleoyl- sn -glycero-3-phosphoethanolamine-N-lissamine rhodamine B sulfonyl ammonium salt (Rhodamine-PE, Cat. 810150) as an acceptor dye were incorporated in PM-liposomes (accordingly 12% unlabeled PE). For FRET measurement using the C2AB domain labeled with Alexa 488, 1.5% Rhodamine-DOPE was included in liposomes as an acceptor dye. Lipid composition of liposomes: protein-free, 45% PC, 13.5% PE, 1.5% Rho-PE, 10% PS, 25% Chol, 4% PI, and 1% PIP 2 . For FRET measurement using labeled SNARE proteins, liposome lipid composition is as follows: 45% PC, 15% PE, 10% PS, 25% Chol, 4% PI, and 1% PIP 2 . Preparation of proteoliposomes Incorporation of the stabilized Q-SNARE complex into large unilamellar vesicles (LUVs) was achieved by OG-mediated reconstitution, called the direct method, i.e. incorporation of proteins into preformed liposomes 16 , 20 , 26 , 29 . LUVs prepared by the direct method were used, unless stated otherwise. Briefly, lipids dissolved in chloroform were mixed according to lipid composition. The solvent was removed using a dry nitrogen stream in a fume hood to form a lipid film, and then lipids were resuspended in 0.5 mL buffer containing 150 mM KCl and 20 mM HEPES/KOH pH 7.4. After sonication on ice, multilamellar vesicles were extruded using polycarbonate membranes of pore size 100 nm (Avanti Polar lipids) to give uniformly-distributed LUVs with average diameter of 110 nm 29 . After the preformed LUVs had been prepared, SNARE proteins (for PM-liposomes) were incorporated into liposomes by using OG, a mild non-ionic detergent, then OG was removed by dialysis overnight in 1 L buffer containing 150 mM KCl and 20 mM HEPES/KOH pH 7.4 together with 2 g SM-2 adsorbent beads. Protein to lipid ratio in proteoliposomes was 1:500 (n/n). Vesicle fusion assay A FRET-based lipid-mixing assay was performed to monitor native vesicle fusion in vitro 20 , 26 – 29 . LDCV fusion assays were performed at 37°C in 1 mL fusion buffer containing 120 mM K-glutamate, 20 mM K-acetate, 20 mM HEPES-KOH (pH 7.4), 1 mM MgCl 2 , and 3 mM ATP. ATP should be made freshly before experiments because it is easily destroyed by freezing and thawing. Free Ca 2+ concentration in the presence of ATP and Mg 2+ was calibrated using the MaxChelator simulation program. The fluorescence dequenching signal was measured using Fluoromax (Horiba Jobin Yvon) with wavelengths of 460 nm for excitation (Ex) and 538 nm for emission (Em). Fluorescence values were normalized as a percentage of maximum donor fluorescence (total fluorescence) after addition of 0.1% Triton X-100 at the end of experiments. Fluorescence anisotropy measurements Anisotropy measurement 20 , 24 was carried out at 37°C in 1 mL buffer containing 120 mM K-glutamate, 20 mM K-acetate, and 20 mM HEPES-KOH (pH 7.4), 1 mM MgCl 2 , and 3 mM ATP, unless stated otherwise. Anisotropy ( r ) was calculated as r = (I VV − G × I VH )/(I VV + 2 ×G × I VH ), where I VV denotes the fluorescence intensity with vertically polarized excitation and vertical polarization on the detected emission, and I VH denotes the fluorescence intensity when using a vertical polarizer on the excitation and horizontal polarizer on the emission. G is a grating factor used as a correction for the instrument’s differential transmission of the two orthogonal vector orientations. Liposomes contained no PS/PIP2: 60% PC, 15% PE, and 25% Chol. Anisotropy (a.u.) was presented as A/A 0 , where A 0 is the initial value. The stabilized Q-SNARE complex (VAMP-2, residues 49–96) and soluble Q-SNARE complex (Syntaxin-1A, residues 183–262) were labeled with Alexa 488; Ex/Em = 488/516 nm. Protein to lipid ratio in proteoliposomes was 1:500 (n/n). Fluorescence resonance energy transfer (FRET) The C2AB domain (40 nM) labeled with Alexa 488 (a donor dye) was incubated with liposomes that include 1.5% Rhodamine-DOPE (an acceptor dye); donor fluorescence signal was measured with wavelengths of 488 nm for excitation and 516 nm for emission 29 . SNAP-25A in the stabilized Q-SNARE complex was labeled with Texas Red at Cys130 (an acceptor dye) and VAMP-2 CD (40 nM) was labeled with Oregon Green 488 at Cys28 (a donor dye). SNARE assembly led to quenching of donor fluorescence with excitation and emission wavelengths for 488 nm and 520 nm, respectively. Donor fluorescence signal was measured at 37°C using Fluoromax (Horiba Jobin Yvon) in 1 mL buffer containing 120 mM K-glutamate, 20 mM K-acetate, 20 mM HEPES-KOH (pH 7.4), 1 mM MgCl 2 , and 3 mM ATP. FRET was normalized as F/F 0 , where F 0 represents the initial value of the donor fluorescence intensity. Determination of tetanus toxin-resistant SNARE complexes The light chain of TeNT is a protease that selectively degrades free VAMP-2, whereas VAMP-2, partially or fully assembled in the ternary SNARE complex, is resistant to cleavage 24 , 50 . Incubation of LDCVs with liposomes containing the stabilized Q-SNARE complex for 10 min at 37°C led to the formation of ternary SNARE complex 24 . For the experiments of SNARE disassembly ( Supplementary Fig. 3 ), LDCVs were preincubated with the soluble Q-SNARE complex (50 nM) for 30 min at 37 °C to induce the ternary SNARE complex formation with VAMP-2 on the LDCV membrane. The ternary SNARE complex was analyzed by SDS-PAGE and immunoblotting with antibody against VAMP-2. 500 nM α-SNAP and 50 nM NSF were added to induce disassembly of the SNARE complex for 30 min at 37 °C. At this time, 200 nM TeNT should be included together to cleave disassembled free VAMP-2 in order to block SNARE re-assembly. The ternary SNARE complex formation on LDCV membrane caused the shift of VAMP-2 to an SDS-resistant band of higher molecular weight noted the ternary complex, which was resistant to TeNT, whereas TeNT completely cleaved free VAMP-2 51 . The samples were not boiled to observe the ternary SNARE complex. Statistical analysis Data analysis was performed using OriginPro 2019 software (OriginLab Corporation, Northampton, MA, USA) and GraphPad Prism 9 (GraphPad Software, San Diego, CA, USA). Data are means ± standard deviation (SD). One-way ANOVA test with Bonferroni correction was used to determine any statistically significant differences among three or more independent groups. Probabilities p < 0.05 were considered significant. Acknowledgements We thank Dr. Reinhard Jahn for the plasmids and samples. We thank Dr. Ahmed Elalawy and Sarra Karrar from Widam Food Company for the arrangement of adrenal glands. This work was supported by the grant from Qatar Biomedical Research Institute (Project Number SF 2019 004 and IGP5-2022-001 to Y.P.), the HBKU Thematic Research Grant (Project Number VPR-TG02-06 to Y.P.) and the Academic Research Grant (Project Number ARG01-0508-230099 to Y.P.). References ↵ Jahn , R. , Cafiso , D. C. & Tamm , L. K . Mechanisms of SNARE proteins in membrane fusion . Nature reviews. Molecular cell biology 25 , 101 – 118 ( 2024 ). https://doi.org/10.1038/s41580-023-00668-x OpenUrl CrossRef PubMed ↵ De Camilli , P. & Jahn , R . Pathways to regulated exocytosis in neurons . Annu Rev Physiol 52 , 625 – 645 ( 1990 ). doi: 10.1146/annurev.ph.52.030190.003205 OpenUrl CrossRef PubMed Web of Science ↵ Park , Y. & Kim , K. T . Short-term plasticity of small synaptic vesicle (SSV) and large dense-core vesicle (LDCV) exocytosis . Cellular signalling 21 , 1465 – 1470 ( 2009 ). doi: 10.1016/j.cellsig.2009.02.015 OpenUrl CrossRef PubMed ↵ Rizo , J. , David , G. , Fealey , M. E. & Jaczynska , K . On the difficulties of characterizing weak protein interactions that are critical for neurotransmitter release . FEBS Open Bio 12 , 1912 – 1938 ( 2022 ). doi: 10.1002/2211-5463.13473 OpenUrl CrossRef PubMed ↵ Brunger , A. T. & Leitz , J . The Core Complex of the Ca(2+)-Triggered Presynaptic Fusion Machinery . J Mol Biol , 167853 ( 2022 ). doi: 10.1016/j.jmb.2022.167853 OpenUrl CrossRef ↵ Park , Y. & Ryu , J. K . Models of synaptotagmin-1 to trigger Ca(2+) -dependent vesicle fusion . FEBS Lett 592 , 3480 – 3492 ( 2018 ). doi: 10.1002/1873-3468.13193 OpenUrl CrossRef PubMed ↵ Brunger , A. T. & Leitz , J . The Core Complex of the Ca(2+)-Triggered Presynaptic Fusion Machinery . J Mol Biol 435 , 167853 ( 2023 ). doi: 10.1016/j.jmb.2022.167853 OpenUrl CrossRef ↵ Llinas , R. , Steinberg , I. Z. & Walton , K . Relationship between presynaptic calcium current and postsynaptic potential in squid giant synapse . Biophys J 33 , 323 – 351 ( 1981 ). S0006-3495(81)84899-0 [pii] doi: 10.1016/S0006-3495(81)84899-0 OpenUrl CrossRef PubMed Web of Science ↵ Sabatini , B. L. & Regehr , W. G . Timing of neurotransmission at fast synapses in the mammalian brain . Nature 384 , 170 – 172 ( 1996 ). doi: 10.1038/384170a0 OpenUrl CrossRef PubMed Web of Science ↵ Radecke , J. et al. Morphofunctional changes at the active zone during synaptic vesicle exocytosis . EMBO Rep 24 , e55719 ( 2023 ). doi: 10.15252/embr.202255719 OpenUrl CrossRef PubMed ↵ Papantoniou , C. et al. Munc13- and SNAP25-dependent molecular bridges play a key role in synaptic vesicle priming . Sci Adv 9 , eadf6222 ( 2023 ). doi: 10.1126/sciadv.adf6222 OpenUrl CrossRef PubMed ↵ Hua , Y. & Scheller , R. H . Three SNARE complexes cooperate to mediate membrane fusion . Proceedings of the National Academy of Sciences of the United States of America 98 , 8065 – 8070 ( 2001 ). doi: 10.1073/pnas.131214798 OpenUrl Abstract / FREE Full Text ↵ Mohrmann , R. , de Wit , H. , Verhage , M. , Neher , E. & Sorensen , J. B . Fast vesicle fusion in living cells requires at least three SNARE complexes . Science 330 , 502 – 505 ( 2010 ). doi: 10.1126/science.1193134 OpenUrl Abstract / FREE Full Text ↵ Zhou , Q. et al. Architecture of the synaptotagmin-SNARE machinery for neuronal exocytosis . Nature 525 , 62 – 67 ( 2015 ). doi: 10.1038/nature14975 OpenUrl CrossRef PubMed ↵ Jaczynska , K. et al. A lever hypothesis for Synaptotagmin-1 action in neurotransmitter release . Proceedings of the National Academy of Sciences of the United States of America 122 , e2417941121 ( 2025 ). doi: 10.1073/pnas.2417941121 OpenUrl CrossRef PubMed ↵ Ali Moussa , H. Y., et al. PIP(2) Is An Electrostatic Catalyst for Vesicle Fusion by Lowering the Hydration Energy: Arresting Vesicle Fusion by Masking PIP(2) . ACS Nano 18 , 12737 – 12748 ( 2024 ). doi: 10.1021/acsnano.3c09614 OpenUrl CrossRef PubMed ↵ Moussa , H. Y. A. , Shin , K. C. & Park , Y . Ca(2+)/calmodulin and protein kinase C (PKC) reverse the vesicle fusion arrest by unmasking PIP(2) . Sci Adv 11 , eadr9859 ( 2025 ). doi: 10.1126/sciadv.adr9859 OpenUrl CrossRef ↵ Perez-Lara , A. et al. PtdInsP(2) and PtdSer cooperate to trap synaptotagmin-1 to the plasma membrane in the presence of calcium . Elife 5 ( 2016 ). doi: 10.7554/eLife.15886 OpenUrl CrossRef PubMed ↵ Liang , Q. et al. Complexin-1 and synaptotagmin-1 compete for binding sites on membranes containing PtdInsP(2) . Biophysical journal 121 , 3370 – 3380 ( 2022 ). doi: 10.1016/j.bpj.2022.08.023 OpenUrl CrossRef PubMed ↵ Park , Y. et al. Synaptotagmin-1 binds to PIP(2)-containing membrane but not to SNAREs at physiological ionic strength . Nature structural & molecular biology 22 , 815 – 823 ( 2015 ). doi: 10.1038/nsmb.3097 OpenUrl CrossRef PubMed ↵ Nyenhuis , S. B. et al. Conserved arginine residues in synaptotagmin 1 regulate fusion pore expansion through membrane contact . Nat Commun 12 , 761 ( 2021 ). doi: 10.1038/s41467-021-21090-x OpenUrl CrossRef PubMed ↵ Voleti , R. , Jaczynska , K. & Rizo , J . Ca(2+)-dependent release of synaptotagmin-1 from the SNARE complex on phosphatidylinositol 4,5-bisphosphate-containing membranes . Elife 9 ( 2020 ). doi: 10.7554/eLife.57154 OpenUrl CrossRef PubMed ↵ Park , Y . Fluorescence Anisotropy for Monitoring cis- and trans-Membrane Interactions of Synaptotagmin-1 . Methods Mol Biol 2887 , 175 – 182 ( 2025 ). https://doi.org/10.1007/978-1-0716-4314-3_12 OpenUrl CrossRef PubMed ↵ Park , Y. et al. alpha-SNAP interferes with the zippering of the SNARE protein membrane fusion machinery . The Journal of biological chemistry 289 , 16326 – 16335 ( 2014 ). doi: 10.1074/jbc.M114.556803 OpenUrl Abstract / FREE Full Text ↵ Siddiqui , T. J. et al. Determinants of synaptobrevin regulation in membranes . Mol Biol Cell 18 , 2037 – 2046 ( 2007 ). doi: 10.1091/mbc.E07-01-0049 OpenUrl Abstract / FREE Full Text ↵ Park , Y. et al. Controlling synaptotagmin activity by electrostatic screening . Nature structural & molecular biology 19 , 991 – 997 ( 2012 ). doi: 10.1038/nsmb.2375 OpenUrl CrossRef PubMed Gumurdu , A. et al. MicroRNA exocytosis by large dense-core vesicle fusion . Sci Rep 7 , 45661 ( 2017 ). doi: 10.1038/srep45661 OpenUrl CrossRef PubMed ↵ Birinci , Y. , Preobraschenski , J. , Ganzella , M. , Jahn , R. & Park , Y . Isolation of large dense-core vesicles from bovine adrenal medulla for functional studies . Sci Rep 10 , 7540 ( 2020 ). doi: 10.1038/s41598-020-64486-3 OpenUrl CrossRef PubMed ↵ Ali Moussa , H. Y., et al. Requirement of Cholesterol for Calcium-Dependent Vesicle Fusion by Strengthening Synaptotagmin-1-Induced Membrane Bending . Adv Sci (Weinh ) , e2206823 ( 2023 ). doi: 10.1002/advs.202206823 OpenUrl CrossRef ↵ Ali Moussa , H. Y. & Park , Y. Electrostatic regulation of the cis- and trans-membrane interactions of synaptotagmin-1 . Sci Rep 12 , 22407 ( 2022 ). doi: 10.1038/s41598-022-26723-9 OpenUrl CrossRef PubMed ↵ Pobbati , A. V. , Stein , A. & Fasshauer , D . N- to C-terminal SNARE complex assembly promotes rapid membrane fusion . Science 313 , 673 – 676 ( 2006 ). doi: 10.1126/science.1129486 OpenUrl Abstract / FREE Full Text ↵ Radhakrishnan , A. , Stein , A. , Jahn , R. & Fasshauer , D . The Ca2+ affinity of synaptotagmin 1 is markedly increased by a specific interaction of its C2B domain with phosphatidylinositol 4,5-bisphosphate . The Journal of biological chemistry 284 , 25749 – 25760 ( 2009 ). M109.042499 [pii] doi: 10.1074/jbc.M109.042499 OpenUrl Abstract / FREE Full Text ↵ Li , L. et al. Phosphatidylinositol phosphates as co-activators of Ca2+ binding to C2 domains of synaptotagmin 1 . The Journal of biological chemistry 281 , 15845 – 15852 ( 2006 ). doi: 10.1074/jbc.M600888200 OpenUrl Abstract / FREE Full Text ↵ Clary , D. O. , Griff , I. C. & Rothman , J. E . SNAPs, a family of NSF attachment proteins involved in intracellular membrane fusion in animals and yeast . Cell 61 , 709 – 721 ( 1990 ). doi: 10.1016/0092-8674(90)90482-t OpenUrl CrossRef PubMed Web of Science ↵ Winter , U. , Chen , X. & Fasshauer , D . A conserved membrane attachment site in alpha-SNAP facilitates N-ethylmaleimide-sensitive factor (NSF)-driven SNARE complex disassembly . The Journal of biological chemistry 284 , 31817 – 31826 ( 2009 ). doi: 10.1074/jbc.M109.045286 OpenUrl Abstract / FREE Full Text ↵ Littleton , J. T. , Stern , M. , Perin , M. & Bellen , H. J . Calcium dependence of neurotransmitter release and rate of spontaneous vesicle fusions are altered in Drosophila synaptotagmin mutants . Proc Natl Acad Sci U S A 91 , 10888 – 10892 ( 1994 ). OpenUrl Abstract / FREE Full Text DiAntonio , A. & Schwarz , T. L . The effect on synaptic physiology of synaptotagmin mutations in Drosophila . Neuron 12 , 909 – 920 ( 1994 ). 0896-6273(94)90342-5 [pii] OpenUrl CrossRef PubMed Web of Science Pang , Z. P. , Sun , J. , Rizo , J. , Maximov , A. & Sudhof , T. C . Genetic analysis of synaptotagmin 2 in spontaneous and Ca2+-triggered neurotransmitter release . EMBO J 25 , 2039 – 2050 ( 2006 ). 7601103 [pii] doi: 10.1038/sj.emboj.7601103 OpenUrl Abstract / FREE Full Text Xu , J. , Pang , Z. P. , Shin , O. H. & Sudhof , T. C . Synaptotagmin-1 functions as a Ca2+ sensor for spontaneous release . Nat Neurosci 12 , 759 – 766 ( 2009 ). nn2320 [pii] doi: 10.1038/nn.2320 OpenUrl CrossRef PubMed Web of Science Chicka , M. C. , Hui , E. , Liu , H. & Chapman , E. R . Synaptotagmin arrests the SNARE complex before triggering fast, efficient membrane fusion in response to Ca2+ . Nat Struct Mol Biol 15 , 827 – 835 ( 2008 ). nsmb.1463 [pii] doi: 10.1038/nsmb.1463 OpenUrl CrossRef PubMed Web of Science Burgalossi , A. et al. SNARE protein recycling by alphaSNAP and betaSNAP supports synaptic vesicle priming . Neuron 68 , 473 – 487 S0896-6273(10)00761-0 [pii] doi: 10.1016/j.neuron.2010.09.019 OpenUrl CrossRef PubMed Web of Science ↵ Littleton , J. T. , Stern , M. , Schulze , K. , Perin , M. & Bellen , H. J . Mutational analysis of Drosophila synaptotagmin demonstrates its essential role in Ca(2+)-activated neurotransmitter release . Cell 74 , 1125 – 1134 ( 1993 ). OpenUrl CrossRef PubMed Web of Science ↵ Brewer , K. D. et al. Dynamic binding mode of a Synaptotagmin-1-SNARE complex in solution . Nature structural & molecular biology 22 , 555 – 564 ( 2015 ). doi: 10.1038/nsmb.3035 OpenUrl CrossRef PubMed ↵ Mackler , J. M. & Reist , N. E . Mutations in the second C2 domain of synaptotagmin disrupt synaptic transmission at Drosophila neuromuscular junctions . J Comp Neurol 436 , 4 – 16 ( 2001 ). OpenUrl CrossRef PubMed Web of Science ↵ Courtney , N. A. , Bao , H. , Briguglio , J. S. & Chapman , E. R . Synaptotagmin 1 clamps synaptic vesicle fusion in mammalian neurons independent of complexin . Nat Commun 10 , 4076 ( 2019 ). doi: 10.1038/s41467-019-12015-w OpenUrl CrossRef PubMed ↵ Toulme , E. , Salazar Lazaro , A. , Trimbuch , T. , Rizo , J. & Rosenmund , C . Neurotransmitter release is triggered by a calcium-induced rearrangement in the Synaptotagmin-1/SNARE complex primary interface . Proceedings of the National Academy of Sciences of the United States of America 121 , e2409636121 ( 2024 ). doi: 10.1073/pnas.2409636121 OpenUrl CrossRef PubMed ↵ Xue , M. , Ma , C. , Craig , T. K. , Rosenmund , C. & Rizo , J . The Janus-faced nature of the C(2)B domain is fundamental for synaptotagmin-1 function . Nature structural & molecular biology 15 , 1160 – 1168 ( 2008 ). doi: 10.1038/nsmb.1508 OpenUrl CrossRef PubMed Web of Science ↵ Pabst , S. et al. Rapid and selective binding to the synaptic SNARE complex suggests a modulatory role of complexins in neuroexocytosis . The Journal of biological chemistry 277 , 7838 – 7848 ( 2002 ). doi: 10.1074/jbc.M109507200 OpenUrl Abstract / FREE Full Text ↵ Stein , A. , Radhakrishnan , A. , Riedel , D. , Fasshauer , D. & Jahn , R . Synaptotagmin activates membrane fusion through a Ca2+-dependent trans interaction with phospholipids . Nature structural & molecular biology 14 , 904 – 911 ( 2007 ). nsmb1305 [pii] doi: 10.1038/nsmb1305 OpenUrl CrossRef PubMed Web of Science ↵ Hayashi , T. et al. Synaptic vesicle membrane fusion complex: action of clostridial neurotoxins on assembly . EMBO J 13 , 5051 – 5061 ( 1994 ). OpenUrl CrossRef PubMed Web of Science Park , Y. et al. Controlling synaptotagmin activity by electrostatic screening . Nat Struct Mol Biol 19 , 991 – 997 nsmb.2375 [pii] doi: 10.1038/nsmb.2375 OpenUrl CrossRef PubMed View the discussion thread. Back to top Previous Next Posted March 24, 2025. Download PDF Email Thank you for your interest in spreading the word about bioRxiv. NOTE: Your email address is requested solely to identify you as the sender of this article. Your Email * Your Name * Send To * Enter multiple addresses on separate lines or separate them with commas. You are going to email the following Synaptotagmin-1 and complexin inhibit spontaneous vesicle fusion by masking PIP2, not by clamping SNARE assembly Message Subject (Your Name) has forwarded a page to you from bioRxiv Message Body (Your Name) thought you would like to see this page from the bioRxiv website. Your Personal Message CAPTCHA This question is for testing whether or not you are a human visitor and to prevent automated spam submissions. Share Synaptotagmin-1 and complexin inhibit spontaneous vesicle fusion by masking PIP2, not by clamping SNARE assembly Houda Yasmine Ali Moussa , Kyung Chul Shin , Yongsoo Park bioRxiv 2025.03.24.644869; doi: https://doi.org/10.1101/2025.03.24.644869 Share This Article: Copy Citation Tools Synaptotagmin-1 and complexin inhibit spontaneous vesicle fusion by masking PIP2, not by clamping SNARE assembly Houda Yasmine Ali Moussa , Kyung Chul Shin , Yongsoo Park bioRxiv 2025.03.24.644869; doi: https://doi.org/10.1101/2025.03.24.644869 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Neuroscience Subject Areas All Articles Animal Behavior and Cognition (7629) Biochemistry (17660) Bioengineering (13881) Bioinformatics (41913) Biophysics (21436) Cancer Biology (18578) Cell Biology (25482) Clinical Trials (138) Developmental Biology (13372) Ecology (19889) Epidemiology (2067) Evolutionary Biology (24302) Genetics (15599) Genomics (22483) Immunology (17728) Microbiology (40365) Molecular Biology (17163) Neuroscience (88540) Paleontology (666) Pathology (2830) Pharmacology and Toxicology (4821) Physiology (7637) Plant Biology (15130) Scientific Communication and Education (2045) Synthetic Biology (4290) Systems Biology (9818) Zoology (2269)