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Munc18-1 prevents the formation of ectopic SNARE complexes in living cells
Claire N. Medine, Colin Rickman, Luke H. Chamberlain, Rory R. Duncan


Membrane trafficking in eukaryotic cells must be strictly regulated both temporally and spatially. The assembly at the plasma membrane of the ternary SNARE complex, formed between syntaxin1a, SNAP-25 and VAMP, is essential for efficient exocytotic membrane fusion. These exocytotic SNAREs are known to be highly promiscuous in their interactions with other non-cognate SNAREs. It is therefore an important cellular requirement to traffic exocytotic SNARE proteins through the endoplasmic reticulum and Golgi complex while avoiding ectopic interactions between SNARE proteins. Here, we show that syntaxin1a traffics in an inactive form to the plasma membrane, requiring a closed-form interaction, but not N-terminal binding, with munc18-1. If syntaxin is permitted to interact with SNAP-25, both proteins fail to traffic to the plasma membrane, becoming trapped in intracellular compartments. The munc18-1–syntaxin interactions must form before syntaxin encounters SNAP-25 in the Golgi complex, preventing the formation of intracellular exocytotic SNARE complexes there. Upon delivery to the plasma membrane, most SNARE clusters in resting cells do not produce detectable FRET between t-SNARE proteins. These observations highlight the crucial role that munc18-1 plays in trafficking syntaxin through the secretory pathway.


Every eukaryotic cell relies on the SNARE proteins to mediate the vesicular targeting, docking and fusion processes that underlie membrane trafficking and exocytosis. Essential to the exocytotic process are the interactions of the plasma membrane proteins (t-SNAREs) syntaxin and SNAP-25 (synaptosome-associated protein 25 kDa) and the vesicular protein synaptobrevin (v-SNARE) (Burgoyne and Morgan, 2003; Sudhof, 1995). Membrane fusion requires the assembly of all three proteins into a trimeric, four-helical complex to promote fusion of the two opposing bilayers in living cells (Sutton et al., 1998; Burgoyne and Morgan, 2003; Sudhof, 1995). This process is tightly regulated by a conserved set of accessory proteins that operate throughout the trafficking pathway. Members of the Sec1p/munc18 (SM) protein family represent one such set of modulators that might add to the specificity of SNARE protein interactions. Indeed, mutations of SM proteins are characterised by a severe disruption of general secretion or neurotransmitter release (Brenner, 1974; Harrison et al., 1994; Novick and Schekman, 1979; Verhage et al., 2000). The possible functions of SM proteins were obtained originally from the observations that munc18-1 (STXB1) binds directly to the monomeric form of syntaxin 1, rendering the t-SNARE unable to form the SDS-resistant ternary SNARE complex (Hata et al., 1993; Pevsner et al., 1994). However, owing to the diversity of SM proteins and SNARE interactions, these functions have remained enigmatic.

Syntaxin can adopt two structurally distinct forms (Dulubova et al., 1999): a closed form in which the three-helical Habc domain is folded back on to the syntaxin SNARE helix, and an open, elongated structure with an accessible SNARE helix. Munc18-1 binds to the closed form of syntaxin with high affinity, rendering the syntaxin unable to enter into SNARE complexes (Hata et al., 1993; Pevsner et al., 1994; Rickman et al., 2007). In addition, munc18-1 can interact with an extreme N-terminal motif of syntaxin (Dulubova et al., 2007; Rickman et al., 2007; Shen et al., 2007). This interaction can occur with syntaxin in the open conformation and most likely is also present in the closed-form mode of interaction. The association of munc18-1 with the syntaxin N-terminus allows this interaction to be maintained throughout the formation of both the binary (syntaxin–SNAP-25 dimer) and ternary SNARE complex, both in vitro and in cells (Dulubova et al., 2007; Rickman et al., 2007). The interaction of munc18-1 with the N-terminal peptide of syntaxin increases the rate of membrane fusion in an in vitro assay (Shen et al., 2007), but the role of the closed-form interaction in the process of exocytosis remains undefined (Burgoyne and Morgan, 2007).

The t-SNARE proteins localise in clusters or spots on the plasma membrane, of between 60-750 nm in diameter, depending on the imaging approach used in visualisation (Lang et al., 2001; Lang et al., 2002; Ohara-Imaizumi et al., 2002; Rickman et al., 2004; Sieber et al., 2007). These clusters have been shown to define the site of vesicle docking and exocytosis in a variety of neuroendocrine cells (Lang et al., 2001; Lang et al., 2002; Ohara-Imaizumi et al., 2002). It is not clear, however, whether every cluster is fusion competent, and indeed it has never been demonstrated convincingly that the t-SNARE proteins contained within each cluster interact before the fusion event occurs. Experiments performed using inside-out lysed sheets of plasma membrane showed that exogenously added VAMP or syntaxin (but interestingly not SNAP-25) interacted with endogenous SNAREs in clusters in `aged' preparations but not in freshly prepared membrane fragments (Lang et al., 2002); this work led to the current hypothesis that the SNAREs are constitutively active.

Several studies have suggested that binding of syntaxin to munc18-1 is required for trafficking of syntaxin to the cell surface (Rowe et al., 2001; Rowe et al., 1999). This requirement has also been observed for the homologous pair of proteins in the yeast Saccharomyces cerevisiae, Vps45p and Tlg2p (SM protein and cognate syntaxin, respectively), whereby ablation of Vps45p resulted in a marked reduction in Tlg2p expression. However, others have reported that syntaxin can traffic to the plasma membrane in the absence of munc18-1 in a variety of different cell types (Schutz et al., 2005; Toonen et al., 2005). Studies using a munc18-1-null mouse demonstrated that syntaxin can be detected at the cell surface (Toonen et al., 2005; Verhage et al., 2000), but, in this system, syntaxin levels are reduced by 70% and other munc18 isoforms still exist (Toonen et al., 2005). It has been suggested that munc18-1 is a `docking factor', based on observations using the same null mouse (Voets et al., 2001). More recently, however, it was demonstrated that docking is actually syntaxin dependent (de Wit et al., 2006). Under certain culture conditions, it has been reported that PC12 cells possess an excess of syntaxin over munc18-1 (Schutz et al., 2005), and this could be used as an argument to suggest that munc18-1 is not required for SNARE trafficking. Other studies (Liu et al., 2004; Rickman et al., 2007) have shown that syntaxin and munc18-1 interact in intracellular membranes in intact cells. However, neither the functional significance, nor the mode(s) of interaction, of such intracellular interactions have been defined.

Here, we show that syntaxin forms stable intracellular SNARE complexes with SNAP-25 if the proteins encounter one another before munc18-1 binding to syntaxin. The formation of such ectopic exocytotic SNARE complexes occurs predominantly in the Golgi complex, preventing trafficking of both t-SNAREs. Using quantitative imaging approaches including colocalisation and fluorescence lifetime imaging (FLIM), we demonstrate that this intracellular trapping of the t-SNAREs depends on their direct interaction and that munc18-1 binding to the closed form of syntaxin, but not the N-terminal peptide motif, inhibits the formation of this complex. Syntaxin thus traffics efficiently through the Golgi complex while bound to munc18-1, and, once at the cell surface, remains inactive even in the presence of SNAP-25 colocalised in clusters. Plasma membrane clusters containing both syntaxin1a and SNAP-25 are heterogeneous in their interaction status. However, the influx of Ca2+ significantly increased the number of SNARE clusters containing interacting t-SNARE proteins. These findings provide important information on the role of munc18-1 in the SNARE trafficking life cycle.


Munc18-1 facilitates syntaxin, but not SNAP-25, trafficking to the cell surface in neuroendocrine cells

It has previously been shown that coexpression of munc18-1 is required for efficient trafficking of syntaxin in nonspecialised cells (Martinez-Arca et al., 2003; Rowe et al., 2001). It has not been demonstrated convincingly, however, whether a SNARE-trafficking function for munc18-1 exists in specialised secretory cells such as neuroendocrine cell lines. To address this, we expressed fluorescent fusions of syntaxin or syntaxin mutants in Neuro2a (N2a) cells, both in isolation or in conjunction with fluorescent munc18-1 (Fig. 1A). Full-length syntaxin (Syx1-288) was never observed to traffic to the cell surface in either HEK293 cells or N2a cells in the absence of coexpressed munc18-1. For all cell types, the expression levels were similar and only cells with the lowest detectable expression levels were selected. Fluorescence correlation spectroscopy (FCS) was performed to estimate the fluorescent protein concentration from multiple points on the plasma membrane of N2a cells expressing fluorescent syntaxin and munc18-1. These experiments used cells expressing heterologous proteins at the same levels as those used for FLIM and colocalisation analyses. FCS revealed that the absolute number of molecules per excitation volume, on the plasma membrane of these cells, was approximately 10 (data not shown). As the excitation volume of the diffraction-limited focal point is sub-femtolitre, the concentration of the proteins under study is in the nanomolar range in the plasma membrane. It should be noted that there are potential caveats to using heterologous proteins expressed on a background of endogenous proteins. We have taken steps, therefore, to minimise overexpression and avoid potential targeting or mislocalisation problems. Other work has utilised primary embryonic cells from animal null models; these approaches are powerful but also subject to caveats: chronic ablation of protein targets might have undesired nonspecific effects on both development and cellular function (de Wit et al., 2006; Gulyas-Kovacs et al., 2007; Toonen et al., 2005). In addition, the proteins studied are present in multiple isoforms in these models and these might functionally substitute for the ablated target. In the absence of a `clean' cellular model, devoid of all munc18 isoforms, and through careful experimental design, low-level acute expression of heterologous proteins in a cellular environment provides a tractable system to investigate the function of syntaxin 1a and munc18-1.

One argument against the role of munc18-1 as a chaperone for syntaxin would be the ability of a mutant syntaxin, spending more time in the `open' conformation (SyxL165E, E166E; Syx1-288[open]) (Dulubova et al., 1999), to traffic to the cell surface. Although munc18-1 has been shown recently to interact with this mutant through an alternative, N-terminal, binding site (Rickman et al., 2007; Shen et al., 2007), the recognised ability of the open mutant to form SNARE complexes while bound to munc18-1 (Rickman et al., 2007) would argue against a `protective' role for munc18-1. To address this, we performed similar experiments, where we expressed Syx1-288[open] either alone or together with munc18-1. In the absence of munc18-1, Syx1-288[open] remained trapped in intracellular membranes (Fig. 1A). When coexpressed with munc18-1, however, Syx1-288[open] trafficked efficiently to the cell surface (Fig. 1A). Both Syx1-288 and Syx1-288[open] interact with munc18-1 in intracellular membranes and on the cell surface (Fig. 1A) (Rickman et al., 2007) (see text below and Figs 3, 4 for details). While these findings could argue against munc18-1 being a chaperone for syntaxin, Syx1-288[open] is not constitutively `open' – it is known to spend time in the closed state (Fasshauer and Margittai, 2004). In addition, this mutant has an affinity for munc18-1 identical to that of wild-type Syx1-288 (5 nM) (Rickman et al., 2007); abolition of the N-terminal motif still allows `open' syntaxin to bind to munc18-1 with high affinity (44 nM) (Rickman et al., 2007). Finally, it was shown recently that Syx1-288[open] can interact with munc18-1 in living cells, but that the interaction in intracellular membranes was significantly decreased compared with wild-type Syx1-288 (Rickman et al., 2007).

Fig. 1.

SNAP-25 but not syntaxin traffics readily to the plasma membrane in live cells, but ectopic complexes can form. (A) Live N2a cells expressing Syx1-288, Syx1-288 coexpressed with munc18-1, Syx1-288[open] or Syx1-288[open] coexpressed with munc18-1. The FLIM data confirmed that Syx1-288 and Syx1-288[open] interact with munc18-1. (B) Fields of PC12 cells expressing the same constructs were scored for syntaxin trafficking to the cell surface. The percentage of cells with surface syntaxin was plotted. (C) N2a cells expressing SNAP-251-206 or truncated SNAP-251-121 were imaged by CLSM. Shown are representative equatorial sections; n>20 images. Bar, 10 μm. (D) Wild-type or mutant SNAP-251-206 (green) and Syx1-288 (red) were expressed in live N2a cells and imaged by CLSM, as before. The merged image shows areas of coincidence in yellow hues. The two-dimensional histogram represents the intensity of each channel in each voxel, with a colour scale representing frequency. The residual map corresponds to weighted residuals from the linear regression of the histogram, thus indicating fluorescent channel covariance. The hue is from –1 to 1, with cyan corresponding to a zero residual. Shown are representative equatorial sections; n>20 images. Bar, 5 μm.

Syntaxin has been shown to traffic to the plasma membrane in PC12 cells without coexpressed munc18-1. In addition, it is thought that syntaxin exists in a large excess over munc18-1 in these cells (Schutz et al., 2005). Both these observations argue against a role for munc18-1 in trafficking syntaxin. To eliminate the possibility that the requirement for munc18-1 in trafficking syntaxin in N2a cells was specific to this cell line, we repeated the same experiments in PC12 cells (Fig. 1B). In this case, syntaxin was seen to traffic efficiently to the cell surface, but in a minority of cells. Whereas 19±2% (mean±s.e.m., n=153 cells from at least four experiments) of cells had syntaxin on the cell surface in the absence of coexpressed munc18-1, 93±5% (mean±s.e.m., n=122) of co-transfected cells had both proteins on the cell surface. Also, 11±4% (mean±s.e.m., n=135) of cells expressing Syx1-288[open] had syntaxin on the cell surface, whereas 37±6% (mean±s.e.m., n=100; significantly lower than for Syx1-288; t test, P<0.001) had Syx1-288[open] and munc18-1. We conclude, therefore, that munc18-1 has an essential role in the trafficking of syntaxin to the cell surface in intact neuroendocrine cells. SNAP-25 trafficking, however, did not rely on munc18-1, or on the presence of any specialised protein (Loranger and Linder, 2002), as it trafficked efficiently to the cell surface in both HEK293 and N2a cells.

Syntaxin and SNAP-25 colocalise in intracellular membranes in the absence of munc18-1

We hypothesised that the role of munc18-1 in trafficking syntaxin might be dependent on spatio-temporal factors. To address this issue, we performed similar experiments in both N2a and HEK293 cells, where we coexpressed Syx1-288 and SNAP-25 or SNAP-25 mutants in the absence of additional munc18-1. These experiments were designed to test the ability of syntaxin to form complexes with SNAP-25; are the SNAREs reactive or non-reactive in intact intracellular membranes? Quantitative colocalisation analyses demonstrated that neither SNARE trafficked to the cell surface in this situation, even in the presence of the endogenous munc18-1 in N2a cells (Fig. 1D). A two-dimensional histogram was generated by analysing the fluorescence intensity, for each channel present in a single voxel, and plotting this on the histogram. This approach was applied to every voxel in the three-dimensional image stack, with the colour scale of the histogram corresponding to the frequency of voxels for each pair of intensity values. This histogram is fit by linear least-squares regression, yielding a Pearson's correlation coefficient. If the concentrations of both proteins are colinear, there will be a high degree of covariance between them – as would be expected for two proteins that interact with a defined stoichiometry. From this fit, residuals for each voxel were calculated and displayed as a `residual map' to highlight areas of covariance (Fig. 1D). Syntaxin and SNAP-25 exhibited a high degree of coincidence and of covariance in perinuclear membranes, reminiscent of the Golgi complex in these cells. To improve our understanding of this effect, we coexpressed Syx1-288 alongside SNAP-251-121. It was demonstrated previously that the single N-terminal SNARE motif of SNAP-25 is sufficient for interaction with syntaxin (Fasshauer et al., 1997). The construct used in our study includes this region necessary for SNARE interaction, in addition to the flexible linker required for localisation to the plasma membrane (Gonzalo et al., 1999). In this case, SNAP-251-121 was seen to traffic to the cell surface, but most remained colocalised with Syx1-288 in intracellular membranes (Fig. 1D). These experiments provided evidence to support the hypothesis that the t-SNAREs are in fact reactive in living cells and that another, spatially and temporally localised, factor is required to prevent them from forming SNARE complexes in intracellular membranes.

Syntaxin and SNAP-25 complexes form in the Golgi complex

Next, we determined the intracellular site of colocalisation and interaction between Syx1-288 and SNAP-25. We performed similar experiments as before, where we coexpressed fluorescent fusions to the t-SNAREs, but this time also coexpressed a marker of the Golgi complex, GRASP-65. A red-fluorescent (mCherry) protein fusion to this marker localises to the cis-Golgi (J. Lane, personal communication) and is spectrally distinct from both cerulean-Syx1-288 and EYFP-SNAP-25. Quantitative colocalisation analysis, as before, confirmed that the site of greatest t-SNARE covariance was the Golgi complex (Fig. 2A). Frequency distribution histograms were generated from these data, emphasising that the weighted residuals centered around zero (i.e. highest covariance) were in the Golgi complex (Fig. 2A, right).

This quantitative analysis of colocalisation between mutant proteins in living cells provided evidence suggesting that the t-SNAREs are able to interact in intracellular membranes. However, colocalisation data are limited by the resolution of the microscope (maximally 200 nm) and are not a direct indication of protein interactions. To increase our understanding of the mechanism whereby SNAP-25 appears trapped by coexpressed Syx1-288, we employed FLIM to detect and quantify directly Förster resonance energy transfer (FRET). FLIM quantifies the excited state fluorescence lifetime of a fluorophore, the duration of which is exquisitely sensitive to the microenvironment it inhabits (Medine et al., 2007). Thus FRET, to an interacting acceptor molecule, dramatically shortens the donor fluorescence lifetime (Lakowicz, 1999); this effect can be quantified directly in each pixel of an image. Importantly, the amount of lifetime quenching is dependent on the distance between the proteins and the fluorophores used; thus, the actual numerical values obtained for each protein pair will vary. Furthermore, the low light levels required for FLIM allowed us to use the same cells as before, selected for the lowest detectable levels of expression. In contrast to other, intensity based, approaches for detecting FRET, this approach has the advantage of being directly quantitative and requires only one measurement. No complex arithmetical adjustments to the pixel data are required, and two-photon excitation is inherently in focus, providing high-resolution data.

Fig. 2.

Syntaxin and SNAP-25 colocalise and can interact in the Golgi complex in live cells. Syx1-288 and SNAP-251-206 coexpressed in live N2a cells and imaged by CLSM. (A) Residual map corresponds to weighted residuals indicating fluorescent channel covariance. The hue is from –1 to 1, with cyan corresponding to a zero residual. Bar, 10 μm. (B) The Golgi complex marker GRASP65-mCherry was coexpressed in live N2a cells and imaged by CLSM (left panel). Residual map of Syx1-288 and SNAP-251-206 covariance in the Golgi complex, as defined by a GRASP65-mCherry mask (middle panel). The weighted residuals of Syx1-288 and SNAP-251-206 contained within the Golgi complex, as defined by GRASP65-mCherry (black circles), alongside residual values from elsewhere in the cell (grey circles), were plotted as a frequency distribution histogram (right panel). (C) mCerulean-Syx1-288 (donor) fluorescence in the absence of EYFP-SNAP-251-206 (acceptor) exhibited an intracellular distribution. The colour scale in the FLIM map represents the fluorescence lifetime [1900 pseconds (red) – 2400 pseconds (blue)]. The fluorescence lifetime values were plotted as a frequency distribution histogram, showing the 99% confidence interval of the Syx1-288 fluorescence lifetime distribution (red line). Syx1-288 alone has a single fluorescence lifetime of 2388±47 pseconds. The excited-state fluorescence decay of Syx1-288 in the absence of an energy acceptor followed a mono-exponential decay (light grey circles) (right panel). (D) mCerulean-Syx1-288 (donor) fluorescence in the presence of SNAP-251-206 (acceptor) exhibited an intracellular distribution. The colour scale in the FLIM map represents the fluorescence lifetime. The fluorescence lifetime values were plotted as a frequency distribution histogram, showing the 99% confidence interval of the Syx1-288 fluorescence lifetime distribution (non-FRET; red line). The fluorescence lifetime of mCerulean-Syx1-288 was shortened significantly to 2176 ±134 pseconds (Mann-Whitney, n=5, P<0.01) when coexpressed with EYFP-SNAP-251-206. In the presence of an energy acceptor, the fluorescence decay of Syx1-288 was fit by a bio-exponential decay function (dark grey circles) (C, right panel). Syx1-288 (donor) fluorescence in the presence of SNAP-251-206 (acceptor) showing in red the pixels containing donor fluorescence lifetimes below than the 99% confidence interval (right panel). Data are expressed as the mean±s.e.m.

We used FLIM analysis to detect FRET between cerulean-Syx1-288 (the donor) and EYFP-SNAP-25 (the acceptor; Fig. 2B). FLIM analysis of N2a cells expressing cerulean-Syx1-288 alone revealed a mono-exponential fluorescence lifetime decay of 2388±47 pseconds (mean±s.d., n=5), in agreement with previous studies of mCerulean (Rizzo et al., 2004) and of cerulean-Syx1-288 (Rickman et al., 2007). In the presence of coexpressed EYFP-SNAP-25, however, this donor lifetime was shortened significantly to 2176±134 pseconds (Mann-Whitney U non-parametric test; P<0.01; mean±s.d., n=5). These data are presented as intensity images, frequency distribution histograms and as FLIM maps (Fig. 2B). FLIM maps represent the measured fluorescence lifetime in each pixel of an image as a false-colour to reveal the intracellular locations where FRET occurs. In this case, statistically significant donor fluorescence lifetime quenching was restricted to the perinuclear Golgi region, where the t-SNAREs colocalise, consistent with the conclusion that the t-SNAREs are highly reactive in the absence of the correct levels of munc18-1 in the appropriate intracellular location.

Munc18-1 interacts with syntaxin in intracellular membranes, preventing formation of ectopic SNARE complexes and permitting efficient trafficking

If our hypothesis is accurate, that munc18-1 at different intracellular locations can prevent Syx1-288 from entering into ectopic SNARE complexes, then coexpressing munc18-1 with the t-SNAREs should allow both Syx1-288 and SNAP-25 to traffic efficiently to the cell surface. We have demonstrated previously that munc18-1 interacts with syntaxin1a in intracellular membranes (Rickman et al., 2007). To explore this further, we coexpressed both t-SNAREs, as before with munc18-1, in HEK293 cells. We used fibroblasts in this experiment as they do not express any endogenous munc18 isoform (Rowe et al., 2001). Coexpression of Syx1-288, or a mutant of Syx lacking the ability to bind to munc18-1 through its N-terminal peptide motif (Syx1-288 Δ6) (Rickman et al., 2007) with SNAP-25 and munc18-1, resulted in the efficient trafficking of both t-SNAREs to the cell surface (Fig. 3). However, when Syx1-288[open] (Rickman et al., 2007) or Syx1-288[open] Δ6 was coexpressed with SNAP-25 and munc18-1, the t-SNAREs remained colocalised in the Golgi complex, as before. Trafficking to the cell surface was thus significantly reduced; we hypothesise that Syx1-288[open] can preferentially form complexes with SNAP-25 in cells even in the presence of munc18-1, as shown previously in vitro (Rickman et al., 2007). These data indicate that munc18-1 can prevent the formation of intracellular t-SNARE complexes and, importantly, show that binding to the closed form of Syx1-288 is required for this function.

SNARE clusters are heterogeneous in their interaction status

If munc18-1 binding to Syx1-288 is required to prevent formation of ectopic t-SNARE complexes, but this in turn inhibits Syx1-288 from entering the ternary SNARE complex, how does exocytosis proceed? The t-SNAREs are known to colocalise in clusters at the cell surface (Lang et al., 2001). Our model is that SNAP-25 can traffic to the cell surface without the involvement of any other specialised factors, as long as closed Syx1-288 is first stabilised by munc18-1 binding. We again used FLIM to determine whether the newly delivered t-SNAREs could interact on the plasma membrane in the presence of munc18-1 (Fig. 4A). Colocalisation analyses as before confirmed that 82±2% of Syx1-288 colocalised with SNAP-25 in these clusters, and that the clusters observed were indistinguishable from endogenous clusters in terms of size, number and density (R.R.D., unpublished). In resting cells, we were unable to detect significant donor (cerulean-Syx1-288) lifetime quenching, even in the presence of colocalised EYFP-SNAP-25. These data can be interpreted in two ways; either the newly arrived t-SNAREs do not interact at the surface before fusion, or the t-SNARE interaction adopts a conformation where the fluorophores are sufficiently distant, or of an orientation, to preclude FRET. We attempted to dissect these possibilities by using a SNAP-25 construct with the fluorescent fusion to the C-terminus, and still no FRET could be detected (R.R.D., unpublished). In either case, these data demonstrate heterogeneity in resting SNARE clusters not previously recognised, owing to alternative SNARE complex conformational states or a sequestration of closed syntaxin by munc18-1.

Depolarisation of the cells with KCl induced FRET in puncta on the cell surface (Fig. 4A, lower panels). To further our understanding of this effect, we performed FLIM imaging at the base of the cells, attempting to acquire FLIM-FRET data from individual SNARE clusters there. As a stimulation paradigm, ionomycin was used to elevate intracellular Ca2+ levels directly, circumventing any potential indirect effect of Ca2+ channels on SNARE interactions. In this case, in both resting cells or in stimulated cells in a Ca2+-free, chelating medium, we saw heterogeneity in the SNARE cluster population (Fig. 4B). After ionomycin-induced Ca2+ influx, this situation was reversed; almost every t-SNARE cluster (95±4%, mean±s.e.m., n=15) now contained interacting SNAREs (Fig. 4C). We investigated this effect further, comparing the donor fluorescence lifetimes of cerulean-Syx colocalized with EYFP-SNAP-25 before and after treatment with KCl (Fig. 4D). Again, the proportion of FRET-positive SNARE clusters increased. The frequency distribution of all the donor fluorescence lifetimes in each sample was fit separately by nonlinear regression (Fig. 4E). These nonlinear fits were significantly different (extra sum-of-squares F test, P<0.0001; Fig. 4E). These data suggest that the formation of suitable syntaxin–SNAP-25 dimeric binary complexes, which can act as acceptor sites for VAMP-containing vesicles, might be a regulatable step of SNARE function.

Fig. 3.

Munc18-1 prevents ectopic t-SNARE interactions. (A) Wild-type or mutants of Syx1-288 (red), SNAP-25 (green) and munc18-1 were coexpressed in live HEK293 cells and imaged by CLSM. The merged image shows areas of coincidence in yellow hues. The two-dimensional histogram represents the intensity of each channel in each voxel, with a colour scale representing frequency. The residual map corresponds to weighted residuals from the line fit to the histogram, thus indicating fluorescent channel covariance. The hue is from –1 to 1, with cyan corresponding to a zero residual. (B) Similar results were obtained using N2a cells. Bars, 5 μm. (C) The covariance data from HEK293 cells coexpressing wild-type or mutants of Syx and SNAP251-206, in the absence (filled bars) or presence of munc18-1 (open bars) were quantified and expressed as Pearson's coefficient values. These data showed that the covariance between the t-SNAREs is unaffected by the presence of munc18-1, but that the location of the colocalisation is altered (see panel A; n⩾4). Data are expressed as the mean±s.e.m.


We have demonstrated that munc18-1 does indeed have an essential role in preventing formation of ectopic t-SNARE complexes during trafficking of syntaxin in neuroendocrine cells. The closed-form mode of binding to syntaxin is necessary and sufficient for this function. While munc18-1 clearly plays a crucial role in trafficking syntaxin to the plasma membrane in neuroendocrine cells, as shown here (Figs 1, 3 and 4), it is possible that a small percentage of syntaxin 1 is able to reach the plasma membrane independently of munc18-1. This possibility would reconcile our findings with previous observations that syntaxin is severely depleted, but still present at low levels, in organisms where a specific SM protein has been ablated (Toonen et al., 2005). Importantly, direct knockdown of expression of syntaxin 1 (de Wit et al., 2006) has been shown to result in a phenocopy of that observed in the munc18-1-knockout mouse (de Wit et al., 2006; Voets et al., 2001). This highlights the close interrelationship between SM proteins and their cognate syntaxin throughout their life cycle. Once at the membrane, the newly delivered t-SNAREs form interaction-heterogeneous clusters, with interactions modulated by elevation of intracellular Ca2+ levels. This enhancement of interaction between the t-SNAREs, illustrated in Fig. 4, could be a shift in the equilibrium from monomeric t-SNAREs to the binary intermediate. However, we cannot exclude that these interacting clusters might also be reporting a conformational change induced by VAMP during the exocytotic events.

Fig. 4.

SNARE clusters are heterogeneous. Intensity images and FLIM maps showing mCerulean-Syx1-288, in the presence of EYFP-SNAP-251-206 and munc18-1 in live N2a cells, imaged by FLIM and two-photon microscopy. (A) mCerulean-Syx1-288 (donor) fluorescence exhibited a plasma membrane distribution. The colour scale [1900 (red) – 2400 pseconds (blue)] in the FLIM map represents the donor fluorescence lifetime in resting cells (top) and after depolarisation with 55 mM KCl (lower panels). These data were plotted as a frequency distribution histogram, showing the 99% confidence interval of the donor lifetime distribution (red dashed line; bars are mean±s.e.m., n=5 experiments). Bar, 5 μm. (B) Similar experiments performed in the presence of ionomycin, but a Ca2+-free environment also revealed clusters at the base of the cell. The colour scale in the FLIM map represents the donor fluorescence lifetime. Bar, 5 μm. [Colour scale: 1500 (red) – 2000 pseconds (blue).] The boxed region of interest is shown in the lower panels and illustrates clusters at the plasma membrane. Pixels containing fluorescence lifetimes below the 99% confidence interval of the non-FRET distribution in panel A are shown in red. Bar, 1 μm. SNARE clusters where no FRET could be detected are highlighted with a dashed circle. (C) In the presence of ionomycin and Ca2+, mCerulean-Syx1-288 (donor) fluorescence in the presence of EYFP-SNAP-251-206 (acceptor) and munc18-1 showed clusters at the base of the cell. The colour scale [1500 (red) – 2000 pseconds (blue)] in the FLIM map represents the donor fluorescence lifetime in the presence of ionomycin and Ca2+. The donor fluorescence lifetime was significantly shortened, indicative of FRET between the t-SNAREs. Bar, 5 μm. The boxed region of interest is shown in the lower panels and illustrates clusters at the plasma membrane. Pixels containing fluorescence lifetimes below the 99% confidence interval of the non-FRET distribution in panel A are shown in red. SNARE clusters where no FRET could be detected are highlighted with a dashed circle. Bar, 1 μm. (D) Similar results were obtained using KCl depolarization. The boxed region of interest is shown in the right-hand panels as a zoomed image, showing that the proportion of FRET-positive t-SNARE clusters increased after KCl-induced depolarization. (E) The donor fluorescence lifetime data for each sample were plotted as a frequency distribution histogram. The fluorescence lifetimes in the KCl-treated samples (open circles, grey fit line) were significantly reduced compared with those from resting cells (filled circles, black fit line).

The role of munc18-1 has been difficult to define and controversial. Conflicting data regarding its role in trafficking syntaxin and its ability to interact with the assembled binary- and ternary-SNARE complexes have hampered progress. We present a model here based on the simplest interpretation of munc18-1 function. The role of munc18-1 in trafficking syntaxin through the secretory pathway relies on it being present in appropriate concentrations at the correct time and in the correct place. If the t-SNAREs encounter one another early in their biogenesis, before munc18-1 inactivation of syntaxin, they can form a misplaced SNARE complex. This complex cannot traffic out of the Golgi complex, perhaps because a fourth SNARE helix is provided by an intracellular SNARE, so trapping the ectopic complex. This hypothesis is supported by including the observations that syntaxin can interact promiscuously with intracellular SNAREs (Fasshauer et al., 1999). SNAP-25 is palmitoylated by Golgi-resident enzymes and requires a functional Golgi complex for membrane association (Gonzalo et al., 1999). Once through intracellular membrane systems, syntaxin remains inactive, while SNAP-25 can proceed efficiently and independently to the plasma membrane and associate with the inner leaflet (Loranger and Linder, 2002). Newly delivered SNAREs form clusters but might remain nonreactive until required. As munc18-1 can remain associated with syntaxin throughout the formation of the binary and ternary SNARE complex, it is possible that munc18-1 can be bound to syntaxin for its entire life cycle, in different conformational forms.

Data from inside-out membrane preparations showed that exogenously added SNARE proteins could interact spontaneously with pre-existing SNARE clusters (Lang et al., 2002). Interestingly, this observation relied on the `ageing' of the membrane preparations: freshly prepared sheets did not allow the exogenous SNAREs to interact. In addition, exogenously added recombinant SNAP-25 could not enter into a complex with the endogenous SNAREs in clusters. The authors concluded that the SNAREs are constitutively reactive and that no additional factor was required to prevent SNARE protein interactions (Lang et al., 2002). Our data are in complete agreement with these findings, but support a different conclusion. We argue that syntaxin is held in an inactive closed form by munc18-1 and that it is feasible therefore that the run-down of the membrane sheet preparation seen during `ageing' is due to loss of soluble munc18-1, along with other soluble cytoplasmic material. In addition, the observed failure of exogenous SNAP-25 to enter into complexes (Lang et al., 2002) also supports our model, where syntaxin is held in an inactive state until required. It has previously been observed in vitro that syntaxin and SNAP-25 can form a stable dimeric complex (Fasshauer and Margittai, 2004; Rickman et al., 2004) and that the formation of this binary intermediate is the rate-limiting step to formation of ternary SNARE complexes (Fasshauer and Margittai, 2004). It is common in biological pathways for the rate-limiting step to form the natural point of regulation (Fersht, 2003). It is highly likely, therefore, that formation of the t-SNARE binary intermediate is a crucial point of regulation in living cells. Indeed, the slow fusion kinetics observed for in vitro reconstituted fusion assays are dramatically accelerated by the presence of a stabilised binary t-SNARE intermediate (Pobbati et al., 2006).

The approaches used in this study permit the determination of protein interactions in situ, meaning that individual SNARE clusters can be examined without prior cell lysis and protein interaction disruption and re-formation in vitro. Thus, these data provide important insights into the spatial mechanisms regulating the biogenesis and function of SNARE proteins. Further work and development of the technologies will be required to quantify protein interactions in living cells to determine protein dynamics immediately before, during and after exocytosis.

Materials and Methods

Vectors and cell culture

The vector pEGFP-C2 was obtained from Clontech (Basingstoke, UK). An EYFP-SNAP-25 fusion was generated by ligation of SNAP-251-206 into BamHI/EcoRI sites of pEGFP-C, followed by the replacement of EGFP with EYFP. SNAP-25–EGFP in pEGFP-N1 was a gift from M. Linder. This construct was used in control FRET experiments. A SNAP-25 mutation, SNAP-251-121, was generated using site-directed mutagenesis with a Quickchange site-directed mutagenesis kit (Stratagene Europe). The plasmids pmCerulean-Syntaxin1a1-288, open (L165A, E166A) mutation and N-terminal truncations of syntaxin1a were described previously (Rickman et al., 2007). A native munc18-1 expression vector was constructed by amplifying the open reading frame of rat munc18-1 and ligating the PCR product to pTarget (Promega). The Golgi complex marker GRASP65-mCherry was obtained from Jon Lane (University of Bristol, UK). Neuroblastoma 2a (N2a) cells and human embryonic kidney (HEK293) cells were cultured in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum, 50 units of penicillin, 50 μg/ml streptomycin and maintained at 37°C in 5% (v/v) CO2, 95% (v/v) air. All cells were cultured on glass coverslips and transfections were performed using ExGen500 (Fermentas). In stimulation experiments, the medium was replaced with Krebs-Ringer bicarbonate buffer (115 mM NaCl, 5 mM KCl, 24 mM NaHCO3, 2.5 mM CaCl2, 1 mM MgCl2, 10 mM HEPES pH 7.4 and 0.1% BSA) for imaging. Cells were stimulated with Krebs-Ringer bicarbonate buffer adjusted to 55 mM KCl or by addition of 100 nM ionomycin and incubated at 37°C for 20 minutes before imaging. At the concentration and time scales used, we saw no morphological evidence of apoptosis.

Confocal laser scanning microscopy and image analysis

All imaging experiments were performed using a Zeiss LSM510 Axiovert confocal laser scanning microscope, equipped with a pulsed excitation source [MIRA 900 Ti:Sapphire femto-second pulsed laser, coupled with a VERDI 10 W pump laser (Coherent)]. Data acquisition was performed using a 1024×1024 pixel image size, using a Zeiss Plan NeoFLUAR 1.4 NA 63× oil-immersion objective lens. Separate fluorescence emission channels were collected simultaneously using multi-track or dual-laser excitation. All imaging was performed using live cells maintained at 37°C in 5% (v/v) CO2, 95% (v/v) air. Image data acquired at Nyquist sampling rates were deconvolved using Huygens software (Scientific Volume Imaging), and the resulting images analysed using NIH ImageJ software ( Residual maps were generated by calculating the residual of each voxel from the linear regression fit to the intensities of each channel within each voxel. The resulting residuals are displayed on a colour scale from –1 to 1 (with zero residual coloured cyan), with brightness corresponding to the combined intensity of the two channels. Cell peripheries were determined using transmitted light imaging combined with CLSM data.

TCSPC-FLIM acquisition and analysis

Time-correlated single-photon counting (TCSPC) measurements were made under 800-820 nm two-photon excitation (TPE), which efficiently excited cerulean, without any detectable direct excitation or emission from EYFP, using a fast photomultiplier tube (H7422; Hamamatsu Photonics UK) coupled directly to the rear port of the Axiovert microscope. TCSPC recordings were acquired for between 10 seconds and 60 seconds, mean photon counts were between 105-106 counts per second. Images were recorded at 256×256 pixels from a 1024×1024 image scan with 256 time bins over a 12 nsecond period. Off-line FLIM data analysis used pixel-based fitting software (SPCImage, Becker & Hickl). The fluorescence was assumed to follow a multi-exponential decay for the purposes of data fitting. In addition, an adaptive offset-correction was performed. This constant offset takes into consideration the time-independent baseline due to dark noise of the detector and the background caused by room light, calculated from the average number of photons per channel preceding the rising part of the fluorescence trace. To fit the parameters of the multi-exponential decay to the fluorescence decay trace measured by the system, a convolution with the instrumental response function was carried out. The optimisation of the fit parameters was performed by using the Levenberg-Marquardt algorithm, minimising the weighted chi-square quantity. This approach can be used to separate the interacting from the non-interacting donor fraction in our FRET systems. The long lifetime component τ2 was determined by control assays with cerulean alone, or expressed with (non-interacting) EYFP, as described above. This value was subsequently used as a fixed τ2 lifetime for all other experiments. As controls for non-specific FRET, or FRET between GFPs that might form dimers spontaneously when overexpressed in cells, we determined the fluorescence lifetimes of cerulean-Syx1-288 alone, cerulean alone, or cerulean-Syx1-288 co-transfected with EYFP (data not shown). No FRET was detected in any of these experiments.


We thank Jon Lane, University of Bristol, UK, for providing a vector encoding mCherry-GRASP65. SNAP-25 EGFP in pEGFP-N1 was a gift from Maurine Linder, Washington University in St Louis, USA. This work was supported by a Wellcome Trust Research Career Development Fellowship to R.R.D. and a Project Grant to R.R.D. and L.H.C.

  • Accepted September 25, 2007.


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