The instructions sent to, from and within the brain are rapidly transmitted along neurons in the form of electrical signals. These signals cannot pass across the small gaps—called synapses—that separate neighboring neurons. Instead, neurons release chemicals called neurotransmitters into the synapses, and these relay the signal to the next neuron.

The neurotransmitters are stored inside neurons in small bubbles called vesicles. To release these neurotransmitters into the synapse, the membrane that encloses the vesicle fuses with the membrane that surrounds the neuron. To fuse the membranes, proteins embedded in the vesicle membrane interact with similar proteins in the neuron membrane to form a structure called a SNARE complex. Additional proteins control membrane fusion to ensure that the signal is passed to the other neuron at the right time and with the appropriate efficiency.

Among these proteins are the complexins, which are often found attached to SNARE complexes. Although different parts of complexins can both help and hinder membrane fusion, a part known as an accessory helix is thought to have only one role—to stop the membranes from fusing together. Several models have been suggested for how the accessory helix interferes with fusion. However, after performing a range of analyses by diverse biophysical techniques, Trimbuch, Xu et al. suggest these models are unlikely to describe the process accurately.

Instead, Trimbuch, Xu et al. propose a new model based on the electrostatic properties of two molecules that are both negatively charged. An accessory helix taken from a fruit fly complexin was more negatively charged than a mammalian version, and experiments showed it was also better at preventing the release of neurotransmitters. It is thought that the negative charges on the helix hold the membranes apart because the helix is located between the membranes, which are also negatively charged. Consistent with this model, Trimbuch, Xu et al. showed that the membranes fused more easily when some of the negative charges on the accessory helix were replaced with positive charges. The next challenges are to test the model further with additional studies, and to explain how other proteins work with complexins to control neurotransmitter release.

Neurotransmitter release is crucial for interneuronal communication and is exquisitely regulated by a sophisticated protein machinery (Sudhof, 2013). Great advances have been made in elucidating the mechanism of release (Brunger et al., 2009; Sorensen, 2009; Jahn and Fasshauer, 2012; Rizo and Sudhof, 2012) and basic aspects of this process have been reconstituted with eight central components of the release machinery (Ma et al., 2013), leading to a model with defined roles for each component. In this model, the neuronal soluble N-ethylmaleimide-sensitive factor attachment protein receptors (SNAREs) synaptobrevin, syntaxin-1 and SNAP-25 form a tight four-helix bundle called the SNARE complex (Sollner et al., 1993; Poirier et al., 1998; Sutton et al., 1998) that brings the synaptic vesicle and plasma membranes together (Hanson et al., 1997) and is critical for membrane fusion; N-ethylmaleimide-sensitive factor (NSF) and soluble NSF attachment proteins (SNAPs) disassemble the SNARE complex (Sollner et al., 1993) to recycle the SNAREs for another round of fusion (Mayer et al., 1996), and may favor physiological membrane fusion by disassembling syntaxin-1-SNAP-25 complexes (Ma et al., 2013); Munc18-1 and Munc13s orchestrate SNARE complex assembly in an NSF/SNAP resistant manner (Ma et al., 2013), and may play a direct role in fusion (Dulubova et al., 2007; Li et al., 2011b); and synaptotagmin-1 acts as a Ca²⁺ sensor (Fernandez-Chacon et al., 2001), likely by bridging the two membranes (Arac et al., 2006; Xue et al., 2008a).

Tight regulation of neurotransmitter release also depends critically on complexins, small soluble proteins that bind to the SNARE complex (McMahon et al., 1995) and play activating and inhibitory functions. Absence of complexins leads to a severe impairment of Ca²⁺-evoked exocytosis and to varied effects on spontaneous release ranging from small decreases to dramatic increases, depending on the system (Reim et al., 2001; Huntwork and Littleton, 2007; Xue et al., 2008b; Maximov et al., 2009; Hobson et al., 2011; Martin et al., 2011; Yang et al., 2013). These results likely arise from an interplay between stimulatory and inhibitory activities of different regions of complexins (Xue et al., 2007, 2009; Cho et al., 2010; Kaeser-Woo et al., 2012). Complexin I (CpxI) is largely unstructured in solution (Pabst et al., 2000) but forms a central α-helix that binds to the SNARE complex and is preceded by an accessory helix (Bracher et al., 2002; Chen et al., 2002) (Figure 1A). The central helix is crucial for both the activating and inhibitory functions of complexins, while the accessory helix inhibits release (Xue et al., 2007; Maximov et al., 2009); the complexin N-terminus plays an activating function, releasing the inhibition by the accessory helix (Xue et al., 2010; Yang et al., 2010), and the C-terminal sequence has activating and inhibitory roles (Kaeser-Woo et al., 2012).

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Cell–cell fusion assays and reconstitution studies also indicated dual roles for complexins that likely recapitulate to some extent their physiological functions (Giraudo et al., 2006; Schaub et al., 2006; Yoon et al., 2008; Malsam et al., 2009, 2012; Diao et al., 2012), but in most cases these studies revealed only stimulatory or inhibitory roles. The activating function has been proposed to arise from stabilization of the SNARE complex by complexin binding (Chen et al., 2002), from interactions of the complexin N-terminus with the C-terminus of the SNARE complex (Xue et al., 2010), and from binding of the complexin C-terminal region to phospholipids (Seiler et al., 2009), but these models remain to be validated. The inhibitory activity of complexins attracted much attention because several studies suggested that complexins prevent exocytosis before Ca²⁺ influx and synaptotagmin-1 releases the inhibition upon Ca²⁺ binding by displacing complexins from the SNARE complex (Giraudo et al., 2006; Schaub et al., 2006; Tang et al., 2006; Roggero et al., 2007). Later analyses showed that CpxI is not fully displaced but there is competition between synaptotagmin-1 and part of CpxI for binding to the SNARE complex on membranes (Dai et al., 2007; Xu et al., 2013). Genetic interaction studies in hippocampal neurons showed that complexins regulate release similarly in the absence or presence of synaptotagmin-1, indicating that complexins function independently of synaptotagmin-1 (Xue et al., 2007, 2010), but this finding does not exclude the notion that complexins and synaptotagmin-1 may cooperate in regulating release. For example, the dramatic increase of spontaneous release observed in complexin nulls in Drosophila requires synaptotagmin-1 (Jorquera et al., 2012), and absence of complexins in hippocampal neurons sensitizes release to loss of 50% of synaptotagmin-1 expression (Xue et al., 2010).

While it is clear that the complexin accessory helix inhibits release (Xue et al., 2007; Maximov et al., 2009), a satisfactory model for this activity has not emerged yet. We initially proposed that part of this helix might replace part of the synaptobrevin SNARE motif in trans partially assembled SNARE complexes, hindering C-terminal assembly of the complex (Xue et al., 2007; Figure 1C). A similar insertion model, but envisioning that the entire accessory helix replaces part of synaptobrevin, was proposed later and was supported by the enhanced inhibition in cell–cell fusion assays caused by replacing charged with hydrophobic residues in the accessory helix of CpxI ('superclamp mutants') and by the design of a ‘poor-clamp’ mutation (K26A) that impairs the inhibitory activity (Giraudo et al., 2009). The crystal structure of a fragment of a CpxI superclamp mutant (D27L, E34F, R37A) bound to a SNARE complex with C-terminally truncated synaptobrevin suggested an alternative model whereby the central helix of one CpxI molecule binds to a SNARE complex and the accessory helix inserts into another SNARE complex, resulting in a zigzag array (Kummel et al., 2011; Figure 1B). However, formation of such a complex with wild type (WT) CpxI would be highly unfavorable thermodynamically because three charged residues would be placed into hydrophobic environments.

The study described here was designed to investigate how the complexin accessory helix inhibits neurotransmitter release, testing the insertion and zigzag models as well as additional models that emerged subsequently (Figure 1D,E). Using NMR spectroscopy and isothermal titration calorimetry (ITC), we show that the accessory helix of CpxI does not insert into synaptobrevin-truncated SNARE complexes in solution. Furthermore, in stark contrast with the cell–cell fusion data, rescue experiments in complexin I-III triple knockout (KO) neurons reveal that superclamp mutations in CpxI lead to slightly stimulatory or no effects on neurotransmitter release, while the poor-clamp K26A mutation impairs release. We also find that the accessory helix of complexin from drosophila melanogaster inhibits spontaneous release more strongly than the accessory helix of mammalian CpxI, which may arise from the more negatively charged nature of the former. Indeed, a mutation that increases the negative charge of the CpxI accessory helix inhibits release and a mutation that decreases the negative charge enhances release. These results suggest a model whereby the location of the negatively charged accessory helix between the synaptic vesicle and plasma membranes causes electrostatic and perhaps steric repulsion with the membranes, thus hindering membrane fusion and neurotransmitter release (Figure 1E).

Complexins are small proteins that play stimulating and inhibitory roles in neurotransmitter release. The inhibitory function was attributed to insertion of the complexin accessory helix into the C-terminus of partially assembled SNARE complexes (Xue et al., 2007; Giraudo et al., 2009; Kummel et al., 2011), but the validity of this model was unclear. In the study presented here, we were unable to detect any interaction between C-terminally-truncated SNARE complexes and the accessory helix of WT CpxI or the CpxI superclamp mutant using highly sensitive biophysical methods in solution. Moreover, we find that the effects of superclamp and poor-clamp CpxI mutations on neurotransmitter release do not correlate with their effects on cell–cell fusion assays, actually pointing in opposite directions. We also show that mutations that increase the negative charge of the accessory helix inhibit neurotransmitter release while mutations that increase its positive charge enhance release. These results strongly argue against the insertion and zigzag models for the inhibitory activity of the complexin accessory helix and suggest a simple, alternative model whereby the negative charges of the accessory helix and perhaps steric hindrance repel both membranes, thus hindering membrane fusion and neurotransmitter release.

A major question that arises from our study is: Can the insertion and zigzag models be now completely ruled out? In addressing this question, it is critical to consider the available data and the arguments that have been used to support these models:

1. We originally proposed that part of the accessory helix might insert into partially assembled SNARE complexes (Figure 1C) to explain the inhibitory function of this helix (Xue et al., 2007), but this proposal was not based on any biochemical data and the model could be questioned based on the paucity of hydrophobic residues in the accessory helix.

2. The related model whereby the entire accessory helix inserts into a partially assembled SNARE complex was supported by the effects of superclamp and poor-clamp mutations (e.g., D27L, E34F, R37A, and K26A) in cell–cell fusion assays (Giraudo et al., 2009). However, the model envisioned that the three charged side chains replaced in the superclamp mutant (D27, E34, and R37) insert into the hydrophobic groove left in the SNARE complex, which is very unlikely from a thermodynamic point of view. Moreover, our data show that the effects of these mutations on cell–cell fusion do not correlate with their effects on neurotransmitter release: the superclamp D27L, E34F, R37A mutation that enhanced inhibition of cell–cell fusion had little effect on release, with a slight tendency to stimulate release, while the K26A mutation that diminishes the inhibitory acitivity of CpxI in cell–cell fusion actually impairs release (Figure 7).

3. It has been argued (e.g., Kummel et al., 2011) that the D27L, E34F, R37A superclamp mutation inhibits neurotransmitter release in vivo based on rescue experiments on complexin knockdown neurons (Yang et al., 2010). However, these rescue experiments revealed no effect of the D27L, E34F, R37A mutation on evoked release, and it was unclear whether a modest inhibitory effect on spontaneous release was significant. Our rescue data with this mutant using complexin I-III triple KO neurons (Figure 7) are consistent with the results of the knockdown rescues considering the small nature of the effects observed, and the experimental differences between the two approaches. Thus, neither of the two studies supports the insertion or zigzag models.

4. The crystal structure of the CpxI(26-83) D27L, E34F, R37A superclamp mutant with the Δ60 synaptobrevin-truncated SNARE complex was purported to demonstrate an alternative insertion model whereby the accessory and central helices bind to different SNARE complexes (zigzag model) (Kummel et al., 2011) (Figure 1B). However, the binding mode observed for the mutant accessory helix is highly unlikely thermodynamically for WT CpxI for the above mentioned reason that three mutated side chains are charged and hence are unlikely to insert into a hydrophobic groove. Correspondingly, our extensive NMR analyses did not detect any interaction between the WT CpxI accessory helix and synaptobrevin-truncated SNARE complexes in solution (Figures 2, 3, Figure 3—figure supplement 1C). We were also unable to detect any interaction in solution between truncated SNARE complexes and the accessory helix even for CpxI superclamp mutant fragments (Figure 2—figure supplement 2, Figure 3—figure supplement 1A,D, Figure 4E). A plausible explanation for these findings is that the interaction between the superclamp accessory helix and the truncated SNARE complex observed in the crystals is very weak in solution, and hence could not be observed in our assays, but is stabilized by crystallization. Note that, although the interface area with the SNAREs is larger for the accessory helix (ca. 900 Ă² calculated with PISA; Krissinel and Henrick, 2007) than for the central helix (ca. 540 Ă²), the atomic B-factors of the residues of the accessory helix in the interface with the SNAREs are much larger than those in the central helix interface, with little electronic density for the side chains of the accessory helix interface (Figure 1—figure supplement 2). Interestingly, it has been suggested that motion at a crystal packing interface is intermediate between that of a solvent accessible surface and that of a protein core, even for large interfaces (Carugo and Argos, 1997). It is also worth noting that Kummel et al. (2011) described another structure obtained with a similar CpxI mutant (D27L, E34M, R37A) where four of the eight crystallographically distinct complexes exhibited the same type of interaction with SCΔ60 observed for the D27L, E34F, R37A CpxI mutant, but the other four complexes had an alternative interaction of the accessory helix with the groove of the truncated SNARE complex where the register was shifted by two helical turns (Kummel et al., 2011).

5. A competition assay monitored by ITC was used to support the conclusion that the accessory helix of WT CpxI inserts into SCΔ60 (Kummel et al., 2011). We can reproduce these data (Figure 4D) but it is clear that the underlying assumption that 1.5 equivalentes of CpxI(47-134) saturate SCΔ60 is incorrect (Figure 4B,C) and that these ITC assays do not detect an interaction of the accessory helix with SCΔ60.

6. Experiments with a surface force apparatus (SFA) were proposed to support the zigzag model (Li et al., 2011a). However, the effects caused by WT complexin II in these experiments could be subject to more than one interpretation; for instance, they are compatible with both the zigzag model and the electrostatic hindrance model that we propose here.

7. The zigzag model was also proposed to be supported by FRET measurements showing that the distances between probes placed on CpxI and SNARE complexes increase on truncation of the synaptobrevin C-terminus (Krishnakumar et al., 2011; Kummel et al., 2011), consistent with the fact that the accessory helix remains close to the SNAREs in the CpxI(26-83)-SNARE complex structure (Figure 1A) but points away from the SNAREs in the structure of the CpxI superclamp mutant bound to SCΔ60 (Figure 1B). We believe that the interpretation of the FRET measurements in terms of static structures constitutes an oversimplification because multiple evidence suggests that there is some flexibility in the accessory helix even in the non-truncated complex, including the poor dispersion of the NH cross-peaks of the accessory helix, their sharper line widths compared to cross-peaks of the central helix, the fact that the NH and Cα chemical shifts of the accessory helix change much less than those of the central helix upon SNARE complex binding and the high B factors observed in the crystal structure of the complex (Chen et al., 2002). Indeed, such flexibility is also consistent with the distance of 20 Å measured by FRET with probes placed at residue 38 of CpxI and residue 193 of SNAP-25 of the SNARE complex (Kummel et al., 2011), since the probes would be expected to be much closer according to the structure of the complex (Chen et al., 2002) (the closest distance between the side chains of these residues is 10 Å). Moreover, our NMR data indicate that the truncations in the SNARE complex further increase the flexibility of the accessory helix (Figure 2—figure supplement 1C,D) and produce flexibility in the N-terminus of the CpxI central helix (Figure 2C) as well as in the C-terminus of SNAP-25 where the FRET donor probe was placed (Figure 3B, Figure 3—figure supplement 1C). The loss in FRET efficiency observed by Kummel et al. upon truncation of the SNARE complex can be readily explained by all these increases in flexibility and the fact that SCΔ60 lacks key residues of synaptobrevin that contact CpxI without the truncation (Figure 1—figure supplement 1).

8. Kummel et al. (2011) concluded that flexibility could not explain the FRET efficiency observed between probes placed at residue 38 of CpxI and residue 193 of SNAP-25 in SCΔ60 because there was no detectable FRET in experiments performed with a CpxI mutant containing a flexible GPGP sequence between the accessory and central helices. However, a small decrease in donor fluorescence was actually observed for this mutant (Figure 4D of Kummel et al., 2011) and, based on other measurements shown in Figure 4C and Supplementary Table 3 in Kummel et al., 2011, such decrease would correspond to a distance of ca. 42 Å, just 8 Å longer than the distance measured without the GPGP insertion. Considering that a flexible sequence of four residues can readily span 8 Å, that the insertion of these residues is expected to push away the fluorescence probes, and that the error in such long distances is expected to be rather large because of the low associated FRET efficiencies, the results obtained with the GPGP mutant are not inconsistent with the conclusion that increased flexbility underlies the decreased FRET caused by the synaptobrevin truncation.

We believe that, although the arguments presented above and the overall available data argue strongly against the validity of the insertion and zigzag models, it might be premature to completely rule out these models given the complexity of this system. Note for instance that our NMR and ITC data were obtained with truncated SNARE complexes in solution and hence do not rule out the possibility that the CpxI accessory helix interacts with trans-SNARE complexes partially assembled between membranes. Moreover, while cell–cell fusion assays do not correlate with our electrophysiological data or with the stimulatory function of complexins in release, these assays were crucial to establish the functional interplay between complexins and synaptotagmin-1 (Giraudo et al., 2006) and did provide support for the insertion model (Giraudo et al., 2009). Hence, we believe that it is advisable to keep an open mind about the insertion or zigzag models, but views considering either of these models proven need to be revised, and alternative models need to be considered.

The model proposed here is attractive because of its simplicity and because it emerges naturally from the realization that the accessory helix of complexins is negatively charged (see Supplementary figure 1 of Huntwork and Littleton (2007)), together with examination of the crystal structure of the WT CpxI(26-83)/SNARE complex. Thus, binding of the central helix to the SNARE complex places the accessory helix right between the membranes at the space where they need to be brought together for fusion (Figure 1E), and such action is very likely to be hindered at least to some extent by steric and electrostatic repulsion between the negatively charged membranes and the accessory helix. Note also that the stronger inhibitory activity of the accessory helix of dmCpx, compared to the mammalian CpxI accessory helix (Figure 8), cannot arise from increased hydrophobicity, but can be explained by our electrostatic hindrance model. The model is also supported by the inhibition of release caused by the 5E mutation, as well as by the increase in release caused by the 3R mutation (Figure 9). Moreover, the impairment in release that we observed for the K26A mutant (Figure 7; charge change = −1), and the increase in spontaneous release caused in rescue experiments on complexin knockdown neurons by a K26E, L41K, E47K mutation (charge change = +1) (Yang et al., 2010) also correlate in general terms with this model. However, these results need to be interpreted with caution, since it is plausible that the 5E and 3R mutations may alter the helical character of the accessary helix, and the considerable magnitude of the effects caused by the K26A and K26E, L41K, E47K mutations suggest that they do not arise only from changes in overall electrostatic potential. Thus, K26 might be involved in some additional, as yet unidentified interaction. Note that K26 does not really form part of the accessory helix in the CpxI(26-83)/SNARE complex and may be involved in releasing the inhibition caused by the accessory helix (Xue et al., 2010).

It is also important to note that the accessory helix does not act on its own and its function depends on the neighboring N-terminus. Our previous studies demonstrate an overall facilitatory effect of complexin for evoked and spontaneous release that requires binding of the complexin N-terminus back onto the SNARE complex (Xue et al., 2010). This proposed loop may serve two functions, namely to further stabilize the trans-SNARE complex and to properly position the accessory helix near the putative fusion area to reduce spontaneous release through its electrostatic repulsion effect. Alternatively, it is plausible that the function of the accessory helix may involve interactions with other components of the release machinery. Clearly, more research will be required to further test the validity of our model, ideally using diverse approaches. Such approaches should include challenging biophysical studies involving trans-SNARE complexes formed between two membranes, reconstitution experiments that have already yielded a wealth of information on complexins (e.g., Schaub et al., 2006; Yoon et al., 2008; Malsam et al., 2009, 2012; Diao et al., 2012), and correlations with additional studies of neurotransmitter release in neurons.

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Author details

1. Thorsten Trimbuch

   NeuroCure Cluster of Excellence, Neuroscience Research Center, Charité-Universitätsmedizin Berlin, Berlin, Germany

   Contribution

   TT, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Contributed equally with

   Junjie Xu

   Competing interests

   No competing interests declared.

2. Junjie Xu

   Department of Biophysics, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   JX, Conception and design, Acquisition of data, Analysis and interpretation of data

   Contributed equally with

   Thorsten Trimbuch

   Competing interests

   No competing interests declared.

3. David Flaherty

   Department of Biophysics, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   DF, Acquisition of data, Analysis and interpretation of data

   Competing interests

   No competing interests declared.

4. Diana R Tomchick

   Department of Biophysics, University of Texas Southwestern Medical Center, Dallas, United States

   Contribution

   DRT, Conception and design, Acquisition of data, Analysis and interpretation of data

   Competing interests

   No competing interests declared.

5. 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

   JR, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   jose@arnie.swmed.edu

   Competing interests

   No competing interests declared.

6. Christian Rosenmund

   NeuroCure Cluster of Excellence, Neuroscience Research Center, Charité-Universitätsmedizin Berlin, Berlin, Germany

   Contribution

   CR, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   christian.rosenmund@charite.de

   Competing interests

   CR: Reviewing editor, eLife.

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