Inside the brain, cells called neurons relay messages from one place to another in the form of electrical signals. When an electrical signal reaches a junction between two neurons (known as a synapse) it triggers small particles called calcium ions to enter one of the cells. This influx of calcium causes vesicles to fuse with the membrane surrounding the neuron and release molecules called neurotransmitters into the small gap between the two neurons. These molecules travel across the gap to activate an electrical signal in the second neuron which then carries the message onwards.

A protein known as synaptotagmin-1 senses calcium ions at synapses and works together with a group of proteins known as the SNARE complex to help vesicles fuse with the cell membrane. Previous studies have reported three different structures of synaptotagmin-1 bound to the SNARE complex in a different way. But it was unclear which of these binding states actually result in the release of neurotransmitters. To address this question, Voleti, Jaczynska and Rizo studied how and when synptotagmin-1 and the SNARE complex bind together using two approaches known as NMR spectroscopy and fluorescence spectroscopy.

The experiments suggest that before calcium enters the synapse, synaptotagmin-1 is already bound to a surface on the SNARE complex. This binding inhibits the release of neurotransmitters and has been reported in previous studies. Adding calcium ions causes synaptotagmin-1 to be released from the SNARE complex. This allows synaptotagmin-1 to interact with the membrane and cooperate with the SNARE complex to trigger vesicle fusion.

Finding out how neurons release neurotransmitters at synapses may help us to understand how the brain works. This could provide new insights into how defects in the synapse lead to neurological disorders, such as schizophrenia, and potentially aid the development of new treatments for such conditions.

The release of neurotransmitters by Ca²⁺-evoked synaptic vesicle exocytosis is an exquisitely regulated process that is critical for communication between neurons. Release involves tethering of synaptic vesicles to presynaptic active zones, priming of the vesicles to a release-ready state, and Ca²⁺-triggered fusion of the vesicle and plasma membranes (Südhof, 2013). Extensive research has allowed reconstitution of basic features of synaptic vesicle fusion with the central components of the neurotransmitter release machinery (Liu et al., 2016; Ma et al., 2013) and led to defined models for their functions (Brunger et al., 2018; Rizo, 2018). The SNAP receptors (SNAREs) syntaxin-1, SNAP-25 and synaptobrevin form a tight four-helix bundle called the SNARE complex that brings the vesicle and plasma membranes together and is key for membrane fusion (Hanson et al., 1997; Poirier et al., 1998; Söllner et al., 1993; Sutton et al., 1998). This complex is disassembled by N-ethylmaleimide sensitive factor (NSF) and soluble NSF adaptors proteins (SNAPs) (Söllner et al., 1993), whereas SNARE complex assembly is organized in an NSF-SNAP-resistant manner by Munc18-1 and Munc13s (Ma et al., 2013; Prinslow et al., 2019). Release is triggered very fast (<1 ms) after Ca²⁺ influx by the Ca²⁺ sensor Synaptotagmin-1 (Syt1) (Fernández-Chacón et al., 2001).

Syt1 is a synaptic vesicle protein with two C₂ domains (named C₂A and C₂B) that form most of its cytoplasmic region and adopt β-sandwich structures that bind multiple Ca²⁺ ions through loops at the top of the β-sandwich (Fernandez et al., 2001; Sutton et al., 1995; Ubach et al., 1998). These loops also mediate Ca²⁺-dependent binding to phospholipids (Chapman and Davis, 1998; Frazier et al., 2003; Zhang et al., 1998), which is crucial for neurotransmitter release (Fernández-Chacón et al., 2001; Rhee et al., 2005). A polybasic region on the side of the C₂B domain β-sandwich also contributes to membrane binding, in part via interactions with phosphatidylinositol 4,5-bisphosphate (PIP₂) (Bai et al., 2004; Li et al., 2006). In addition, the Syt1 C₂B domain can bind simultaneously to two membranes in a Ca²⁺ dependent manner through its Ca²⁺-binding loops and two arginines at the bottom of the β-sandwich (R398,R399) that are crucial for Syt1 function (Xue et al., 2008), suggesting that Syt1 cooperates with the SNAREs in bringing the vesicle and plasma membranes together to mediate membrane fusion (Araç et al., 2006; Seven et al., 2013). Induction of membrane curvature by insertion of the C₂ domain Ca²⁺-binding loops into the bilayer was also proposed to stimulate membrane fusion (Martens et al., 2007). While these ideas are attractive, the mechanism of action of Syt1 remains enigmatic, in part because dozens of papers have described Syt1-SNARE interactions but it is unclear which of these interactions is physiologically relevant and which ones arise merely from the high promiscuity of these proteins (Jahn and Scheller, 2006; Rizo et al., 2006). Moreover, Syt1 is believed to function in a tight interplay with complexins whereby complexins play both inhibitory and stimulatory functions and Ca²⁺ binding to Syt1 releases the inhibition (Giraudo et al., 2006; Schaub et al., 2006; Tang et al., 2006), but the underlying mechanism is unknown.

Potentially critical insights were provided by three structures of Syt1-SNARE complexes (Brewer et al., 2015; Zhou et al., 2015; Zhou et al., 2017), but the observed binding modes were drastically different (Figure 1A–G). Analysis in solution by NMR spectroscopy revealed a dynamic interaction involving the polybasic region of the Syt1 C₂B domain, while R398,R399 at the bottom of C₂B remain available for membrane binding (Brewer et al., 2015; Figure 1A). The functional relevance of this binding mode was supported by the finding that release was disrupted by mutations of basic residues that contact the SNAREs (including R322E,K325E), but not by nearby basic residues that do not point toward the SNAREs (K324E,K326E) (Figure 1D), while both mutations disrupted PIP₂ binding similarly (Brewer et al., 2015). The second structure, determined by X-ray crystallography, revealed a so-called primary interface involving two regions of the C₂B domain, one formed largely by E295 and Y338 (region I) and another including R398,R399 (region II) (Figure 1B,E,F; Zhou et al., 2015). In this structure, the polybasic region is available for PIP₂ binding, an arrangement that was supported by a low-resolution cryo-electron microscopy (cryo-EM) structure of Syt1-SNARE complexes coating lipid tubes (Grushin et al., 2019). The physiological relevance of the primary interface was supported by the strong impairments of release caused by mutations in both region I and II (E295A/Y338W and R398Q/R399Q). However, Syt1-SNARE co-immunoprecipitation (co-IP) was not significantly altered by the R398Q/R399Q mutation and only moderately disrupted by the E295A/Y338W mutation (Zhou et al., 2015).

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X-ray crystallography of a tripartite Syt1-complexin-1-SNARE complex yielded a third structure (Zhou et al., 2017) where complexin-1 forms an α-helix that binds to a groove between synaptobrevin and syntaxin-1, as observed for a binary complexin-1-SNARE complex (Chen et al., 2002), and an α-helix of the C₂B domain binds to the same groove, continuing the complexin-1 helix (Figure 1C,G). Isothermal titration calorimetry (ITC) supported the existence of this interaction in solution and suggested that Syt1 cooperates with complexin-1 for binding to the SNARE complex, providing an explanation for the finding that the dominant negative effect on release caused by mutation of the Syt1 C₂B domain Ca²⁺ binding sites (Mackler et al., 2002; Wu et al., 2017) requires complexin-1 (Zhou et al., 2017). The functional importance of the tripartite interface was supported by the observation that a L387Q/L394Q mutation disrupted binding and impaired neurotransmitter release. However, as pointed out in Zhou et al., 2017, the primary interface was also observed in these crystals, which had a 1:1:1 Syt1-SNARE complex-complexin-1 stoichiometry. Hence, one of the two interfaces might constitute a crystal contact. Moreover, a screen for mutations that abrogate the dominant negative effect of mutations in the Syt1 C₂B domain Ca²⁺-binding sites in Drosophila yielded a large number of mutations in the primary interface but none in the tripartite interface (Guan et al., 2017), indicating that the dominant negative effect may require Syt1-SNARE complex binding through the primary rather than the tripartite interface.

The role of Ca²⁺ in Syt1-SNARE interactions has also been enigmatic. Many studies reported that Ca²⁺ strongly stimulates binding of Syt1 to SNAREs and SNARE complexes both in solution and on membranes [e.g. (Chapman et al., 1995; Dai et al., 2007; Davis et al., 1999; Gerona et al., 2000; Huang and Cafiso, 2008; Li et al., 1995; Lynch et al., 2007; Zhou et al., 2013)]. Paradoxically, the Syt1 Ca²⁺-binding loops are not involved in SNARE complex binding in the structures described above, although Ca²⁺ is believed to stimulate binding by increasing the positive electrostatic potential of the C₂B domain, as the SNARE complex is negatively charged. Note also that Ca²⁺ disrupted Syt1-SNARE complexes assembled on lipid tubes (Grushin et al., 2019), but another study reported binding of Syt1 to membrane-anchored SNARE complex in the absence and presence of Ca²⁺ (Wang et al., 2016). Conversely, ATP was reported to abolish Syt1-SNARE complex at physiological ionic strength, with or without Ca²⁺ (Park et al., 2015).

To shed light into this strikingly confusing picture and decipher how the functions of Syt1 and the SNAREs are coupled, we performed a systematic analysis of Syt1-SNARE complex interactions in solution and on membranes. NMR data show that a soluble complexin-1-SNARE complex binds to the polybasic region and the primary interface of the Syt1 C₂B domain with similar affinities, but binding to the tripartite interface could not be detected. Analyses of binding of a Syt1 fragment spanning the two C₂ domains (C₂AB) to nanodiscs or nanodiscs containing anchored SNARE complexes using FRET show that C₂AB binds to membrane-anchored SNARE complexes under a variety of conditions, and that complexin-1 does not affect the affinity. Binding of the SNARE complex to the primary interface is impaired by the R398Q/R399Q mutation but is enhanced by the E295A/Y338W mutation, both in solution and on nanodiscs, showing that the primary interface is involved in binding to the nanodisc-anchored SNARE complex. Importantly, Ca²⁺-dependent binding of C₂AB to SNARE complexes anchored on PIP₂-nanodiscs is almost abolished in the presence of ATP, but Ca²⁺-independent binding remains. We also observed that the R322E/K325E mutation disrupts Ca²⁺-dependent binding of C₂AB to PIP₂-containing nanodiscs much more strongly than the K324E/K326E mutation, in correlation with the effects of these mutations on release (Brewer et al., 2015). Together with previous data, these results suggest a model whereby Syt1 binds to the SNARE complex through the primary interface and to PIP₂ through the polybasic region before Ca²⁺ influx; Ca²⁺ binding to the C₂B domain induces a specific, PIP₂-dependent interaction with the plasma membrane, disrupting the interaction with the SNARE complex and allowing cooperation between the SNAREs and Syt1 in inducing membrane fusion.

Because of the well-established functions of the SNAREs as the engines of membrane fusion and of Syt1 as the Ca²⁺ sensor that triggers synchronous neurotransmitter release, elucidating the role(s) of Syt1-SNARE interactions is crucial to understand how Ca²⁺ sensing is coupled to membrane fusion during release. The recent determination of three structures of Syt1-SNARE complexes led to intriguing models of Ca²⁺-triggered release (Brewer et al., 2015; Zhou et al., 2015; Zhou et al., 2017), but yielded a confusing picture because of the striking differences among the structures. Paradoxically, Ca²⁺ was generally believed to enhance Syt1-SNARE binding, but the C₂ domain Ca²⁺-binding loops were not involved in SNARE binding in any of the structures. Together with available data, the study presented here suggests that Syt1 binds to the SNARE complex before Ca²⁺ influx, most likely through the primary interface, and that Ca²⁺ actually releases this interaction, inducing tight membrane binding that involves specific interactions of the C₂B polybasic region with PIP₂. We propose that the Syt1-SNARE complex keeps the release machinery in a state that hinders membrane fusion but at the same time is ready for fast release when Ca²⁺ induces binding of the Syt1 Ca²⁺-binding loops to the membrane, releasing the Syt1-SNARE interaction and enabling cooperation between Syt1 and the SNAREs in membrane fusion (Figure 10).

Figure 10

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To rationalize the immense amount of data available on Syt1-SNARE interactions, it is crucial to decipher which data reflect physiologically relevant interactions and which arise from the promiscuity of these proteins, the lack of key components in a reduced system and/or the choice of experimental conditions. This task is hindered by the fact that relevant interactions may be necessarily weak because of the very nature of this dynamic system, which is expected to undergo quick, drastic rearrangements during the events that lead to fast membrane fusion upon Ca²⁺ influx. Weak interactions can be dramatically enhanced by co-localization within the confines of a primed synaptic vesicle, but they must be distinguished from other, perhaps stronger but irrelevant interactions that are detectable in vitro (Magdziarek et al., 2020). Another complicating aspect is that basic sequences such as the polybasic region and R398,R399 can bind to SNAREs and to membranes, both of which are acidic, and binding to a wrong molecule can occur in the absence of the bona fide target. Perhaps the most confusing factor was the observation by multiple labs that Ca²⁺ strongly enhanced binding of Syt1 to SNAREs or SNARE complexes in solution (see introduction), leading to the widespread belief that elucidating how Ca²⁺-bound Syt1 binds to the SNARE complex was a ‘Holy Grail’ to understand neurotransmitter release. However, it now seems clear that Ca²⁺ releases Syt1 from SNARE complexes anchored on PIP₂-containing membranes such as the plasma membrane and that the Ca²⁺-induced increase in Syt1-SNARE binding in solution arose merely because Ca²⁺ increases the positive electrostatic potential of the Syt1 C₂ domains (Fernandez et al., 2001; Shao et al., 1997). This enhanced Syt1-SNARE affinity is offset by the specific, Ca²⁺ -dependent interaction of Syt1 with PIP₂-containing membranes, which is incompatible with SNARE complex binding and is much stronger. This key conclusion arises from several lines of evidence.

First, our titrations with NDs and cisSC-NDs show that physiological conditions including Mg-ATP almost abolish Ca²⁺-dependent binding of C₂AB* to the SNARE complex anchored on PIP₂-containing membranes (Figure 9B), consistent with previous results (Park et al., 2015). A previous study reported that Syt1-SNARE complex interactions persist in the presence of ATP (Wang et al., 2016), but this conclusion was tested only in the absence of Ca²⁺ and is thus consistent with our results (Figure 9A).

Second, and particularly important is the observation of the high specificity of Ca²⁺-dependent binding of C₂AB* to PIP₂-containing NDs, which is disrupted much more strongly by the R322E/K325E mutation than by the K324E/K326E mutation (Figures 6G and 9F), in correlation with the effects of these mutations on neurotransmitter release (Brewer et al., 2015). Note that, remarkably, the striking difference in the effects of the two mutations on Ca²⁺-dependent binding of C₂AB* to PIP₂-containing NDs was not observed in Ca²⁺-dependent binding to NDs lacking PIP₂ (Figure 6E) or Ca²⁺-independent binding to NDs with or without PIP₂, and with or without SNARE complex (Figure 6A–D). As explained above, modeling readily explains these observations, as R322 and K325 (but not K324 and K326) are expected to be well positioned to interact with PIP₂ upon Ca²⁺-induced binding of the C₂B domain to the membrane (Figure 10B). In the absence of Ca²⁺, insertion of the Ca²⁺-binding loops into the membrane is hindered by the negative charge of the loops (Fernandez et al., 2001) and, consequently, more parallel orientations of the C₂B domain with respect to the membrane that allow simultaneous binding of the primary interface to the SNARE complex (Figure 10A) are favored. These parallel orientations bring K324 and K326 near the membrane and hence PIP₂ can readily interact with these residues as well as with R322 and K325.

Third, binding of the C₂B domain to the membrane in the perpendicular orientation induced by Ca²⁺ and PIP₂ is incompatible with the binding modes observed in the three structures of Syt1-SNARE complexes that have been determined. The elongated SNARE complex could not remain membrane-anchored or would have strong steric clashes if it remained bound to the C₂B domain through the primary or tripartite interfaces, and binding of the SNARE complex to the polybasic region is hindered because this region interacts with PIP₂. These conclusions are supported by the observation that highly efficient FRET between C₂AB* and the SNARE complex labeled at residue 76 of SNAP-25 was observed under a variety of conditions except for those that allowed Ca²⁺-dependent binding of C₂AB* to PIP₂-containing SNARE complex nanodiscs (Figure 8). Note also that the physiological relevance of our NMR structure was supported by the differential disruption of C₂B-SNARE complex binding and neurotransmitter release caused by the R322E/K325E and K324E/K326E mutations (Brewer et al., 2015). However, the physiological data can now be explained by the differential effects of these mutations on Ca²⁺-dependent binding of C₂AB* to PIP₂-containing membranes (Figures 6G and 9F), which occurs with high affinity (in the low nM range) and is much tighter than SNARE complex binding (Figure 8D).

Syt1 does bind to membrane-anchored SNARE complexes in the absence of Ca²⁺, even in the presence of ATP (Figure 9A; Wang et al., 2016). Our NMR studies show that there are two main Ca²⁺-independent binding modes between the C₂B domain and CpxSC in solution (Figure 1) that are mediated by the primary and polybasic interfaces. The strong disruption of CpxSC binding to the primary interface caused by the R398Q/R399Q mutation (Figure 2B,C) shows that R398,R399 contribute substantially to the energy of binding at this interface. E295 and Y338 likely contribute also to the binding energy, but additional mutations will be necessary to assess this contribution, as the E295A/Y338W mutation actually enhances binding (Figure 4). These observations contrast with the finding that co-IP of Syt1 with the SNAREs was not substantially affected by the R398Q/R399Q mutation and was moderately disrupted by the E295A/Y338W mutation (Zhou et al., 2015). This discrepancy might arise because co-IP likely detects SNARE binding to the polybasic region upon membrane solubilization, and depends not only on affinities but also on off rates. The effects of the R398Q/R399Q and E295A/Y338W mutations observed by NMR correlate with those observed on Ca²⁺-independent binding of C₂AB* to cisSC-NDs (Figures 6B,D, 7A, 8 and 9D), showing that binding to the SNARE complex on the nanodiscs is mediated in part by the primary interface. However, the R322E/K325E and K324E/K326E mutations also decreased the differences in the affinities of C₂AB* for NDs and cisSC-NDs (Figure 6—figure supplements 1–3; Figure 9—figure supplement 3), indicating that there are additional binding modes where the C₂B polybasic region interacts with the SNAREs. These other binding modes are unlikely to be biologically relevant, as the K324E/K326E mutation does not impair neurotransmitter release substantially (Brewer et al., 2015).

The functional importance of binding of Syt1 to the SNARE complex through the primary interface is overwhelmingly supported by physiological data (Guan et al., 2017; Zhou et al., 2015). Hence, it is most likely that binding of C₂AB* to the SNARE complex in cisSC-NDs through the polybasic region arises from limitations of our in vitro experiments and that Syt1 is bound to the SNARE complex through the primary interface before Ca²⁺ influx. Although this interaction is rather weak, it is likely to be dramatically stabilized by co-localization at the site of fusion and may be favored over other binding modes by other factors. For instance, our experiments used 1% PIP₂, which corresponds to the average PIP₂ content of the plasma membrane, but clusters containing 6% PIP₂ have been detected (James et al., 2008; van den Bogaart et al., 2011). High PIP₂ concentration should favor simultaneous binding of the C₂B domain polybasic region to this lipid and of the primary interface to the SNARE complex (Figure 10A,C). Such a state has been observed by cryo-EM on lipid nanotubes, albeit at limited resolution (Grushin et al., 2019). The structure revealed a slanted orientation of the SNARE complex that should hinder full zippering at the C-terminus and was disrupted by Ca²⁺. It is unclear whether the orientation was dictated by helical packing on the nanotubes and whether Ca²⁺ disrupted such packing, but these results are consistent with the notion that Ca²⁺ releases the interaction between Syt1 and membrane-anchored SNARE complexes. Even if the SNARE complex is less slanted in the primed state, a more parallel orientation of the SNARE complex with respect to the membrane (e.g. Figure 10A) would still hinder C-terminal zippering, which could explain the increase in spontaneous release observed in Syt1 KO mice (Xu et al., 2009). The proposed state is also consistent with the observation that complexin-1 does not alter the affinity of C₂AB* for cisSC-NDs (Figure 9E), as complexin-1 and Syt1 bind to opposite sides of the SNARE complex (Figure 10A). In this state, the helix formed by complexin-1 would point toward the vesicle membrane (Figure 10A,C), also hindering SNARE complex zippering (Trimbuch et al., 2014). This model provides a basis for the findings that complexin-1 is required for the dominant negative effect of Syt1 bearing mutations in the C₂B domain Ca²⁺-binding sites (Zhou et al., 2017) and that Syt1 is necessary for the inhibition of spontaneous release by complexin in Drosophila (Jorquera et al., 2012).

Our finding that the R398Q/R399Q mutation strongly disrupts binding of the C₂B domain to the SNARE complex through the primary interface (Figure 2B,C) suggests that such disruption underlies the dramatic impairment of Ca²⁺-evoked neurotransmitter release caused by this mutation, as well as the abrogation of clamping of spontaneous release by Syt1 (Xue et al., 2008; Zhou et al., 2015). The E295A/Y338W mutation in the primary interface also disrupted neurotransmitter release but still allowed clamping of spontaneous release, which resembles the phenotype observed for the R322E/K325E mutation in the polybasic region (Zhou et al., 2015). These findings are readily explained by our model and our biochemical data, as the E295A/Y338W enhances (rather than weakens) Ca²⁺-independent binding of Syt1 to the SNARE complex (Figures 4, 7A, 8 and 9D) and hence hinders release of the inhibitory Syt1-SNARE interaction. Similarly, the R322E/K325E mutation impairs Ca²⁺- and PIP₂-dependent membrane binding, which is critical to release this inhibitory interaction.

Altogether, the available data support the notion that, before Ca²⁺ influx, the trans-SNARE complex, complexin-1 and Syt1 form a macromolecular assembly between the vesicle and plasma membranes that inhibits release but is ready to trigger fast membrane fusion upon Ca²⁺ influx because it brings Syt1 close to the SNAREs (Figure 10C left panel). In this model, Ca²⁺ triggers tight, specific binding of the C₂B domain to PIP₂ and other lipids in the plasma membrane while binding of C₂B to the SNARE complex is released. The mechanisms underlying the last events leading to membrane fusion are still unclear, but we speculate that the Syt1 C₂ domains bridge the vesicle and plasma membranes (Araç et al., 2006; Figure 10C, middle panel). This action could cooperate with the SNAREs in bringing the membranes together, and insertion of the C₂ domain Ca²⁺-binding loops into the membrane can induce membrane curvature (Martens et al., 2007) to facilitate membrane fusion (Figure 10C, right panel). Note that R398,R399 are critical for the membrane-membrane bridging activity of Syt1 and therefore the strong disruption of neurotransmitter release caused by mutating these arginines might arise from impairment of this activity (Araç et al., 2006; Xue et al., 2008) in addition to abrogating SNARE binding.

Clearly, the proposed model needs further testing and multiple aspects need to be unraveled to understand the mechanism of neurotransmitter release. Thus, syntaxin-1 is known to form clusters on the plasma membrane (Khuong et al., 2013), and formation of Syt1 oligomers has been proposed to underlie the primed state and to exert an inhibitory activity that is released by Ca²⁺ (Wang et al., 2017). Such clusters and oligomers could not be present in our nanodisc experiments. However, the Syt1 oligomers are compatible with the state proposed in Figure 10A (Tagliatti et al., 2020) and hence can be readily incorporated into the model of Figure 10C. Note also that, while our NMR studies could not detect binding of the Syt1 C₂B domain to CpxSC through the tripartite interface (Figures 2E and 3), it is premature to completely rule out the relevance of the tripartite structure given the potential functional importance of very weak interactions (Magdziarek et al., 2020). Nevertheless, it is also premature to accept this structure as physiologically relevant, and further evidence for such relevance needs to be obtained, ideally using mutations that exclusively replace residues in the interface rather than interior residues that are important for protein stability. The model of Figure 10 provides a strong foundation to address these issues and incorporate additional proteins that may play key roles in the last steps of neurotransmitter release.

The NMR data corresponding to Figs. 1-4 and corresponding figure supplements are publicly available at https://doi.org/10.5061/dryad.0zpc866vt. Source data are provided for all the FRET experiments shown in Figs. 5-9 and corresponding figure supplements.

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Author details

1. Rashmi Voleti

   1. Department of Biophysics, University of Texas Southwestern Medical Center, Dallas, United States
   2. Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, United States
   3. Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Conceptualization, Formal analysis, Investigation, Methodology, Writing - review and editing

   Contributed equally with

   Klaudia Jaczynska

   Competing interests

   No competing interests declared

2. Klaudia Jaczynska

   1. Department of Biophysics, University of Texas Southwestern Medical Center, Dallas, United States
   2. Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, United States
   3. Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Conceptualization, Formal analysis, Investigation, Methodology, Writing - review and editing

   Contributed equally with

   Rashmi Voleti

   Competing interests

   No competing interests declared

3. Josep Rizo

   1. Department of Biophysics, University of Texas Southwestern Medical Center, Dallas, United States
   2. Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, United States
   3. Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Writing - original draft

   For correspondence

   Jose.Rizo-Rey@UTSouthwestern.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-1773-8311

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