The release of neurotransmitters by Ca²⁺-triggered synaptic vesicle exocytosis is key for communication between neurons. The high speed of this process arises in part because synaptic vesicles are first tethered at the plasma membrane and undergo priming processes that leave them ready for fast fusion when an action potential induces Ca²⁺ influx (Südhof, 2013). Research for over three decades has led to extensive characterization of the neurotransmitter release machinery (Brunger et al., 2018; Rizo, 2022), allowing reconstitution of basic features of synaptic vesicle fusion with the central components of this machinery (Lai et al., 2017; Liu et al., 2016; Ma et al., 2013) and uncovering key aspects of the underlying mechanisms. The SNARE proteins syntaxin-1, SNAP-25 and synaptobrevin form a tight four-helix bundle called the SNARE complex that assembles (zippers) from the N- to the C-terminus to bring the vesicle and plasma membranes together, and is key for membrane fusion (Hanson et al., 1997b; Poirier et al., 1998; Söllner et al., 1993; Sutton et al., 1998). This complex is disassembled by NSF and SNAPs (no relation to SNAP-25) to recycle the SNAREs for another round of fusion (Mayer et al., 1996; Söllner et al., 1993). Munc18-1 and Munc13-1 play central roles in synaptic vesicle priming by organizing assembly of the SNARE complex via an NSF-SNAP-resistant pathway (Ma et al., 2013; Prinslow et al., 2019) whereby Munc18-1 first binds to a self-inhibited ‘closed’ conformation of syntaxin-1 (Dulubova et al., 1999; Misura et al., 2000). Munc13-1 later opens syntaxin-1 (Ma et al., 2011; Yang et al., 2015) while bridging the vesicle and plasma membranes (Quade et al., 2019; Xu et al., 2017), and Munc18-1 forms a template for SNARE complex assembly (Baker et al., 2015; Jiao et al., 2018; Parisotto et al., 2014; Sitarska et al., 2017).

Synaptotagmin-1 (Syt1) acts as the Ca²⁺ sensor that triggers fast release (Fernández-Chacón et al., 2001) in a tight interplay with complexins whereby both Syt1 and complexins play inhibitory and active roles (Giraudo et al., 2006; Reim et al., 2001; Schaub et al., 2006; Tang et al., 2006). The Ca²⁺ sensor function is performed by the two C₂ domains of Syt1 (C₂A and C₂B), which form most of its cytoplasmic region and bind multiple Ca²⁺ ions via loops at the tip of β-sandwich structures (Fernandez et al., 2001; Sutton et al., 1995; Ubach et al., 1998; Figure 1—figure supplement 1A). These loops also mediate Ca²⁺-dependent binding of both C₂ domains to phospholipids, which is critical for release (Fernández-Chacón et al., 2001; Rhee et al., 2005). The C₂B domain also binds to PIP₂ through a polybasic region on the side of the β-sandwich (Bai et al., 2003), which induces binding to the plasma membrane. Moreover, the C₂B domain can bind to the SNARE complex through three different surfaces (Brewer et al., 2015; Zhou et al., 2015; Zhou et al., 2017), although only binding through a so-called primary interface (Zhou et al., 2015) is firmly established as physiologically relevant (Guan et al., 2017; Voleti et al., 2020). Such binding is disrupted upon Ca²⁺-dependent binding of Syt1 to PIP₂-containing membranes (Voleti et al., 2020).

Complexin-1 binds tightly to the SNARE complex through a central α-helix that is preceded by an accessory helix (Chen et al., 2002) and may play a stimulatory role in release by promoting formation of a primed state with enhanced release probability (Chen et al., 2002), by protecting trans-SNARE complexes from disassembly by NSF and αSNAP (Prinslow et al., 2019) and/or by synchronizing Ca²⁺-triggered fusion mediated by the SNAREs and Syt1 (Diao et al., 2012). The complexin-1 accessory helix inhibits release (Xue et al., 2007), possibly because it causes steric clashes with the vesicle, hindering C-terminal assembly of the SNARE complex (Radoff et al., 2014; Trimbuch et al., 2014). These and other findings suggested that complexin-1 and Syt1 bind simultaneously to the SNARE complex in the primed state of synaptic vesicles, stabilizing this state and preventing premature fusion; in this model, Ca²⁺ influx relieves the inhibition by inducing dissociation of Syt1 from the SNAREs and enabling cooperation between Syt1 and the SNAREs in promoting fusion (Voleti et al., 2020).

Despite this wealth of knowledge, fundamental questions remain about the mechanism of neurotransmitter release, particularly regarding how the SNAREs and Syt1 trigger fast, Ca²⁺-dependent membrane fusion. Major hurdles to address this question are the dynamic nature of this process and the fact that the protein complexes that trigger fusion are assembled between two membranes. Although important clues on the nature of the primed macromolecular assembly have been obtained with structural studies of soluble proteins or complexes anchored on one membrane (Chen et al., 2002; Grushin et al., 2019; Voleti et al., 2020; Zhou et al., 2015), this assembly is most likely affected by its location between two membranes. This feature strongly hinders the possibility of crystallization, while application of NMR spectroscopy for structure elucidation is hampered by the large size of any reconstituted two-membrane system (Voleti et al., 2021). Conversely, the small size of the SNAREs, Syt1 and complexin-1 hinders visualization by cryo-EM. Moreover, it is extremely challenging to capture transient states formed during the pathway to Ca²⁺-triggered membrane fusion experimentally.

Molecular dynamics (MD) simulations offer a powerful tool to analyze dynamic biomolecular processes and model cellular membranes (Marrink et al., 2019). Simulations using continuum and/or coarse-grained representations have provided important insights into SNARE-mediated membrane fusion (Fortoul et al., 2015; Kasson et al., 2006; Manca et al., 2019; McDargh et al., 2018; Mostafavi et al., 2017; Risselada et al., 2011; Sharma and Lindau, 2018). Continuum models can access the longest timescales, but require experimental data or atomistic simulations to parameterize the material properties, and often need to constrain geometries or material properties due to lack of context-dependent parameters (Fortoul et al., 2015). Coarse-grained molecular simulation approaches are freed from some of these constraints but at the expense of reduced simulation speed, and are limited in their ability to capture certain entropic effects and protein conformational changes (see below). To date, coarse-grained models of SNARE-mediated fusion have accessed the low microsecond timescale (Kasson et al., 2006; Risselada et al., 2011; Sharma and Lindau, 2018). All-atom simulations are better suited to reproduce the finely-balanced network of interactions between proteins, Ca²⁺, and lipids that are expected to lead to membrane fusion but, because of the large size of the systems involved (millions of atoms), the low microsecond time scale has only recently become accessible through the most powerful available high-performance computing resources. In this context, it is worth noting that the delay from Ca²⁺ influx into the presynaptic terminal to observation of postsynaptic currents in rat cerebellar synapses at 38 °C is 60 μs (Sabatini and Regehr, 1996), and that multiple events occur within this time frame, including Ca²⁺ binding to the sensor, release of inhibitory interactions that hinder premature fusion, Ca²⁺-evoked synaptic vesicle fusion, opening of the fusion pore, diffusion of neurotransmitters through the synaptic cleft, binding of the neurotransmitters to their postsynaptic receptors and opening of the channels that underlie the postsynaptic currents. These observations suggest that the fusion step may occur in just a few microseconds and hence that it may be possible to recapitulate the initiation of Ca²⁺-dependent synaptic vesicle fusions in all-atom MD simulations starting with a properly designed initial configuration.

Here, we present all-atom MD simulations with explicit water molecules of systems containing four trans-SNARE complexes bridging two flat bilayers or a vesicle and a flat bilayer, without or with fragments of Syt1 and/or complexin-1. Because of the limited simulation times and the absence of key components, our results cannot lead to definitive conclusions but they help visualize potential trajectories and intermediates along the pathway to fusion and reveal intriguing features, leading to predictions or hypotheses that can be tested experimentally and with additional simulations. Our data indicate that trans-SNARE complexes strongly pull two membranes together, as expected, but have a tendency to induce extended membrane-membrane adhesion interfaces that have been observed experimentally but fuse slowly (Hernandez et al., 2012; Witkowska et al., 2021). Our results also suggest that, in the primed state of synaptic vesicles, Syt1 and complexin-1 form a spring-loaded macromolecular assembly with trans-SNARE complexes that hinders formation of such extended contact interfactes and premature bilayer merger, but is ready for fast membrane fusion upon Ca²⁺ influx.

Enormous advances have been made to elucidate the molecular mechanisms underlying neurotransmitter release and have suggested that Syt1 and complexin are bound to trans-SNARE complexes in the primed state that renders synaptic vesicles ready for fast fusion upon Ca²⁺ influx. However, the configuration of the resulting macromolecular assembly is still unclear. Our MD simulations, together with previously available data, suggest that trans-SNARE complexes alone induce extended vesicle-plasma membrane contact interfaces that fuse slowly. Our results also indicate that binding of Syt1 and complexin-1 to trans-SNARE complexes in primed vesicles leads to a spring-loaded arrangement that hinders formation of such extended contact interfaces, keeping the system ready for fast fusion but at the same time hindering premature fusion before Ca²⁺ influx.

Our results need to be interpreted with caution because of the limited simulation times, the dependence of the results on the initial configurations and the absence of key elements of the release machinery. Nevertheless, multiple features observed in the simulations make sense from structural and energetic points of view, and are consistent with abundant experimental data available on this system. The SNARE four-helix bundle strongly drew the membranes together in our simulations with SNAREs alone (Figure 1A–D), in agreement with the high stability of the four-helix bundle [e.g. Gao et al., 2012]. However, assembly of the four-helix bundle brought the flat bilayers within a few nm from each other, and it is unclear how the SNAREs exert additional force on the membranes to induce fusion. A major problem with the widespread notion that the SNARE motif, juxtamembrane linker and TM region of synaptobrevin and syntaxin-1 form continuous helices that force fusion as they zipper from the N- to the C-terminus (Hanson et al., 1997a; Sutton et al., 1998; Weber et al., 1998) is that the bent conformations of the linkers envisioned in these models are unrealistic from an energetic point of view. Although optical tweezer data suggested that interactions between the juxtamembrane linkers contribute to exerting force on the membranes to induce fusion (Gao et al., 2012) helix continuity in the linkers is not required for neurotransmitter release (Kesavan et al., 2007; Zhou et al., 2013). Substantial release was observed even upon insertion of a five-residue sequence into the synaptobrevin linker (Kesavan et al., 2007) despite the fact that this sequence should break the register of linker-linker interactions and contained two (helix disrupting) glycine residues. Note also that the optical tweezer data were obtained in the absence of membranes and that, in vivo, the linkers are likely to interact with the lipids given the proximity of each linker to the adjacent membrane and the abundance of basic residues in the linker sequence (and aromatic residues in the case of synaptobrevin). The extensive interactions of the linkers with the membranes observed in all our simulations support this prediction.

Based on conformational grounds, it is not surprising that the juxtamembrane linkers became unstructured during the simulation that we performed to generate a trans-SNARE complex starting from the crystal structure of the cis-SNARE complex. Note however that we did not perform a systematic analysis to examine the range of linker structures that are compatible with the geometry of a trans-SNARE complex. In any case, the linkers are expected to be unstructured before SNARE complex assembly, which is supported by EPR data (Kim et al., 2002). Therefore, configurations with unstructured linkers were natural, unbiased starting points for the simulations, and helical conformation could be adopted by the linkers during the simulations if they were preferred. However, the linkers remained unstructured during all our simulations, which facilitated the extensive linker-membrane interactions observed. These interactions may have contributed to pull the two flat bilayers together in our first simulation (Figure 1A–D) and to induce the formation of an extended contact interface in the simulation with a vesicle and a flat bilayer (Figure 2J). Coarse-grained simulations have suggested that formation of such interfaces can also arise from entropic forces that favor outward movement of the SNARE complexes, away from the center of the interface (Mostafavi et al., 2017). These extended interfaces have been observed by cryo-EM and by fluorescence microscopy, and evolve to fusion in long time scales (seconds-minutes) (Diao et al., 2012; Hernandez et al., 2012; Witkowska et al., 2021), in agreement with data showing that liposome fusion occurs minutes after liposome docking (Cypionka et al., 2009). SNARE-mediated fusion was also observed at faster time scales (Domanska et al., 2009; Heo et al., 2021). Thus, it is plausible that fusion is slower under conditions that favor formation of extended interfaces. Interestingly, cryo-EM studies indicated that formation of such extended interfaces is hindered by other proteins involved in Ca²⁺-evoked release, which favor point-of-contact interfaces that fuse faster (Diao et al., 2012; Gipson et al., 2017).

Our two simulations including Syt1 C₂AB and the complexin-1 (27–72) fragment are consistent with this proposal, as these proteins appeared to hinder formation of extended interfaces (Figure 4—figure supplement 4), but longer simulations will be required to further test this notion. The initial configurations used in these simulations (Figure 3A, Figure 4—figure supplement 2A) were built to mimic and investigate potential states of the Syt1-SNARE-complexin-1 macromolecular assemblies that are likely central components of the primed state of synaptic vesicles. Overwhelming evidence supports the physiological relevance of the binding modes of the SNARE complex to complexin-1 and to the primary interface of the Syt1 C₂B domain used to build these initial configurations (Chen et al., 2002; Guan et al., 2017; Xue et al., 2007; Zhou et al., 2015). These binding modes were largely preserved in the eight primed complexes during the two simulations, although there were slight rearrangements in the primary interface that allow closer interactions of the critical R398 side chain of the C₂B domain (Xue et al., 2008) with acidic residues from the SNARE complex (Figure 4F).

In the initial configurations, we placed the C₂B domain between the SNARE complex and the flat bilayer to facilitate interactions of a polybasic β-strand (residues 321–327) with PIP₂ that are believed to mediate binding of the C₂B domain to the plasma membrane (Bai et al., 2003). Interestingly, each of the eight primed complexes was re-oriented during the two simulations to enable extensive interactions of the C₂B domain with the flat bilayer that involve not only the polybasic β-strand but also other basic residues from the same side of the β-sandwich that are also known to play a key role in release [e.g. K313; Brewer et al., 2015; Figure 4G, Figure 4—figure supplement 1F–H, Figure 4A-D]. Thus, the resulting C₂B domain-flat bilayer binding mode is consistent with the physiological importance of multiple residues of the polybasic face (Brewer et al., 2015). This binding mode resembles that observed in previous all-atom simulations of a C₂AB-SNARE-complexin-1 assembly on a flat bilayer (Bykhovskaia, 2021) and makes sense from a physicochemical point of view given the numerous electrostatic interactions involved. It is also worth noting that the extent of C-terminal assembly of the SNARE four-helix bundle was distinct in each of the eight complexes. Thus, the orientation of the C₂B domain-SNARE-Cpx1(27-72) assembly with respect to the membranes consistently observed in the eight complexes can be accommodated by different extents of C-terminal assembly. All these observations suggest that the C₂B domain binds to the plasma membrane in a similar mode in the primed state of synaptic vesicles and that some of the configurations of the complexes visited in the simulations resemble those present in the primed state.

Due to the C₂B-flat bilayer interactions, the SNARE four-helix bundle came near the flat bilayer, with the C₂B domain on one side rather than between the bundle and the flat bilayer, and with the helix of Cpx1(27-72) oriented toward the vesicle (Figure 4A–D, Figure 4—figure supplement Figure 4—figure supplements 2 and 3A–D). This orientation dictates that formation of a continuous, straight helix by Cpx1(27-72) leads to steric clashes with the vesicle if the SNARE four-helix bundle is fully or close to fully zippered, consistent with the proposal that such steric clashes underlie the inhibitory role of the accessory helix and with physiological data supporting this notion (Trimbuch et al., 2014). These steric clashes most likely cause, at least to some extent, the distinct ways by which the Cpx1(27-72) helix was bent in the different complexes (Figures 3D–K–4A–D, Figure 4—figure supplement 3A-D). These variations may also arise in part from random motions and are consistent with data suggesting that the accessory helix can adopt additional orientations that depend on the extent of C-terminal zippering (Choi et al., 2016; Kümmel et al., 2011). However, NMR data showed that the accessory helix sequence has a high propensity to form α-helical conformation that nucleates the central helix and that structures with a continuous, straight helix as observed by X-ray crystallography are substantially populated when there is no steric hindrance (Chen et al., 2002; Pabst et al., 2000; Radoff et al., 2014). Overall, these observations show that, even if there are natural motions in the complexin-1 helix, transient formation of a continuous straight helix results in steric clashes with the vesicle if the four-helix bundle zippers fully.

It is difficult to estimate the energy barrier imposed by this behavior on SNARE zippering, but it is known that the energy cost of breaking just one hydrogen bond to distort the complexin-1 helix can be about 1.8 k_BT (Nick Pace et al., 2014), which could translate to 7.2 k_BT by cooperative action of four primed complexes. Such an energy barrier would slow down SNARE zippering by a factor of 1340 (=e^7.2). For comparison, the enhancements of spontaneous neurotransmitter release observed upon deletion of complexin range from 3 to >20 (Huntwork and Littleton, 2007; Martin et al., 2011). Thus, even a small energy barrier caused by steric hindrance can account for these enhancements. Note also that we observed occasional interactions of the Cpx1(27-72) accessory helix with C-terminal residues of the synaptobrevin and SNAP-25 SNARE motifs that are favored by proximity (e.g. Figure 3G, Figure 3—figure supplement 2G), and recent cross-linking experiments suggested that very weak interactions of the complexin accessory helix with synaptobrevin and SNAP-25 hinder C-terminal zippering and release (Malsam et al., 2020). It is unlikely that specific interactions underlie the complexin inhibitory function, as this function was retained when the accessory helix was replaced with an unrelated helical sequence, and helix propensity appears to be the key determinant for the inhibitory function (Radoff et al., 2014). However, very weak complexin-SNARE interactions that do not need to be specific may slow down C-terminal SNARE zippering and thus contribute also to inhibition of release by the complex accessory helix.

While all these observations support the proposal that the consistent overall configuration of the Syt1 C₂B domain-SNARE-Cpx1(27-72)-flat bilayer assembly observed for the eight primed complexes in the two simulations resembles that present in primed vesicles, there are clear uncertainties with regard to the extent of C-terminal SNARE zippering and the vesicle-plasma membrane distance, which are closely related. In our two simulations, the vesicle was drawn into contact with the flat bilayer. However, other factors in addition to complexin are likely to hinder zippering in vivo, most notably Munc13-1 because in the absence of Ca²⁺ it bridges membranes in approximately perpendicular orientations that keep the membranes apart (Camacho et al., 2021; Grushin et al., 2022; Quade et al., 2019). Analyses by high-pressure freezing electron tomography (ET) (Imig et al., 2014) and cryo-ET (Radhakrishnan et al., 2021) showed that docked vesicles exhibit a distribution of distances from the plasma membrane that range from 0 to several nm, and a density map built from a subset of subtomograms revealed a distance of 3.5 nm. However, a recent model that can explain a large amount of available presynaptic plasticity data invoked two primed states, one that involves partially assembled SNARE complexes and has low release probability (loose state), and another with more fully assembled SNARE complexes that has a much higher release probability (tight state; Neher and Brose, 2018). Hence, it is plausible that the vesicles that are closest to the plasma membrane account for much of the neurotransmitter release observed.

Based on these observations and our simulations, we envision two scenarios. In one scenario, vesicles in the tight primed state are very close to or in contact with the plasma membrane and include primed complexes that resemble PC1 at the end of each of our two simulations (Figure 3E, Figure 4—figure supplement 2E). In both of these poses, the four-helix bundle was almost fully zippered, but the vesicle was not as close to the flat bilayer around the C-terminus of the four-helix bundle as in the simulation with only SNAREs (Figure 6A–C), which led to an extended vesicle-flat bilayer interface (Figure 2J). Since complexin-1 binding stabilizes the C-terminus of the SNARE complex (Chen et al., 2002), it is tempting to speculate that, to maximize the speed of release, complexin-1 favors formation of an almost fully zippered, tight primed state that is ready for fast Ca²⁺-triggered fusion, but hinders the ‘final pull’ of the SNARE four-helix bundle to bring the membranes together, preventing premature fusion as well as formation of an extended interface. In this spring-loaded model, dissociation of the C₂B domain from the SNARE complex upon Ca²⁺ influx (Voleti et al., 2020) allows rotations of the four-helix bundle that eliminate the steric clashes and facilitate cooperation between the Syt1 C₂ domains and the SNAREs in triggering fusion by a mechanism that remains unclear. Hence, this model can explain the finding that complexins are required for the dominant negative effect of mutations in the Ca²⁺-binding site of the Syt1 C₂B domain (Zhou et al., 2017). In the second scenario, the distance from the vesicle to the plasma membrane in the tight primed state is a few nm. Manually moving the vesicle in the pose of Figure 6B to a distance of 3 nm from the flat bilayer (Figure 6D) shows that the same spring-loaded configuration of the Syt1 C₂B domain-SNARE-Cpx1(27-72)-flat bilayer assembly could be kept by stretching the synaptobrevin juxtamembrane linker, and the Cpx1(27-72) helix would still hinder progress toward final zippering and fusion. This configuration is also largely compatible with longer vesicle-flat bilayer distances if the linker is stretched further and/or there is partial SNARE unzippering. In this case, steric clashes of the Cpx1(27-72) helix with the vesicle might be alleviated or eliminated, but they would occur as soon there is full zippering and hence would still hinder vesicle fusion.

Figure 6

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Clearly, these models will need to be tested with further MD simulations and experimentation, and there are multiple factors of the primed state of synaptic vesicles that were not included in the simulations. As mentioned above, particularly important for this state is Munc13-1, but other missing elements include Munc18-1, the N-terminal region of syntaxin-1, the SNAP-25 linker joining its SNARE motifs, the N- and C-terminal regions of complexin-1 and the TM region plus linker sequence of Syt1 [reviewed in Rizo, 2022]. Moreover, the Syt1 C₂B domain can also bind to the SNARE complex through a so-called tripartite interface whereby an α-helix of the C₂B domain is adjacent to the complexin-1 helix (Zhou et al., 2017), which could stabilize the complexin-1-SNARE interface to hinder rotations that might lead to dissociation (see Figure 3K, Figure 3—figure supplement 2F). There is also evidence that Syt1 forms oligomeric rings that hinder spontaneous neurotransmitter release (Tagliatti et al., 2020), and it is plausible that the SNAREs alone can induce fast fusion in different configurations that we did not study. Furthermore, some evidence suggested many years ago that direct binding of Ca²⁺ to phospholipids could trigger synaptic vesicle fusion (Papahadjopoulos et al., 1976) and, although it is now generally believed that Ca²⁺ triggers neurotransmitter release by binding to Syt1, it is plausible that Ca²⁺-phospholipid interactions might also contribute to trigger membrane fusion. Addressing all these issues with all-atom MD simulations will be challenging because of the limited simulation times that are currently reachable. Continuum and coarse-grained simulations, which have already provided important insights into SNARE-mediated membrane fusion (Fortoul et al., 2015; Manca et al., 2019; McDargh et al., 2018; Mostafavi et al., 2017; Risselada et al., 2011; Sharma and Lindau, 2018), offer the opportunity to explore much longer time scales with similar systems. Hence, a marriage of approaches whereby the most interesting results obtained by such simulations are investigated in further detail by all-atom simulations will likely provide a powerful strategy to unravel the intricate mechanisms that govern fast Ca²⁺-triggered membrane fusion. The systems that we present here provide a framework to pursue these studies and gradually incorporate additional elements of the neurotransmitter release machinery.

Most files corresponding to our molecular dynamics simulations are available in the dryad database (doi:10.5061/dryad.ns1rn8pw6). Because of the very large size of trajectory files, it was not practical to deposit them in this database, but these files are available from the corresponding author upon reasonable request.

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Author details

1. Josep Rizo

   1. Department of Biophysics, The 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, Resources, Validation, Visualization, 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

2. Levent Sari

   1. Department of Biophysics, The University of Texas Southwestern Medical Center, Dallas, United States
   2. Green Center for Systems Biology, The University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Conceptualization, Formal analysis, Methodology, Validation, Visualization, Writing – review and editing

   Competing interests

   No competing interests declared

3. Yife Qi

   Department of Medicinal Chemistry, School of Pharmacy, Fudan University, Shanghai, China

   Contribution

   Funding acquisition, Investigation, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

4. Wonpil Im

   1. Department of Biological Sciences, Lehigh University, Bethlehem, United States
   2. Department of Chemistry, Lehigh University, Bethlehem, United States
   3. Department of Bioengineering, Lehigh University, Bethlehem, United States
   4. Department of Computer Science and Engineering, Lehigh University, Bethlehem, United States

   Contribution

   Funding acquisition, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5642-6041

5. Milo M Lin

   1. Department of Biophysics, The University of Texas Southwestern Medical Center, Dallas, United States
   2. Green Center for Systems Biology, The University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   Conceptualization, Methodology, Validation, Visualization, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-8680-2685

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