Munc18-1 is believed to prime or stimulate SNARE-mediated membrane fusion/exocytosis through binding to the SNARE complex, in addition to chaperoning its cognate syntaxins. Nevertheless, a Munc18-1 mutant that selectively loses the priming function while retaining the syntaxin chaperoning activity has not been identified. As a consequence, the mechanism that mediates Munc18-1-dependent priming remains unclear. In the course of analyzing the functional outcomes of a variety of point mutations in domain 3a of Munc18-1, we discovered insertion mutants (K332E/K333E with insertions of 5 or 39 residues). These mutants completely lose their ability to rescue secretion whereas they effectively restore syntaxin-1 expression at the plasma membrane as well as dense-core vesicle docking in Munc18-1 and Munc18-2 double-knockdown PC12 cells. The mutants can bind syntaxin-1A in a stoichiometric manner. However, binding to the SNARE complex is impaired compared with the wild type or the hydrophobic pocket mutant (F115E). Our results suggest that the domain 3a of Munc18-1 plays a crucial role in priming of exocytosis, which is independent of its syntaxin-1 chaperoning activity and is downstream of dense-core vesicle docking. We also suggest that the priming mechanism of Munc18-1 involves its domain-3a-dependent interaction with the SNARE complex.
Synaptic exocytosis/membrane fusion is highly governed by neuronal Sec1/Munc18 (SM) protein, Munc18-1. The essence of Munc18-1 has been clearly evidenced by strikingly impaired exocytosis of the null mutants of Munc18-1 and its orthologues in mice (Verhage et al., 2000; Voets et al., 2001), Drosophila (Rop) (Harrison et al., 1994) and Caenorhabditis elegans (Unc-18) (Hosono et al., 1992). Nevertheless, the precise modality of Munc18-1 contribution to exocytosis is still poorly understood. At least three important functions of Munc18-1 have been proposed (Han et al., 2010): (i) molecular chaperone of syntaxin-1, allowing proper localization and expression of syntaxin-1 (Arunachalam et al., 2008; Han et al., 2011; Han et al., 2009; Malintan et al., 2009; McEwen and Kaplan, 2008; Medine et al., 2007; Rowe et al., 2001; Rowe et al., 1999); (ii) priming via promotion of SNARE complex-mediated membrane fusion (Rodkey et al., 2008; Shen et al., 2007; Südhof and Rothman, 2009; Tareste et al., 2008); (iii) docking of large dense-core vesicles to the plasma membrane (Toonen et al., 2006; Voets et al., 2001).
We recently demonstrated that the domain 1 surface (residues 34–71) of Munc18-1 serves an important role in the chaperoning activity of Munc18-1, which is crucial for dense-core vesicle docking and exocytosis (Han et al., 2011; Han et al., 2009). The K46E/E59K mutant was discovered as the key ‘chaperoning mutant’ that essentially loses its abilities to bind to the ‘closed’ conformation of syntaxin-1 and consequently to restore syntaxin-1 expression, localization, dense-core vesicle docking and secretion in Munc18-1/2 double-knockdown (DKD) PC12 cells. However, the chaperoning function of Munc18-1 itself is insufficient to explain the near complete loss of exocytosis in Munc18-1-deficient neurons. For instance, Munc18-1 knockout in neurons causes a strong (∼50–75%), but not complete reduction in syntaxin-1A level (Verhage et al., 2000). Also, in contrast to severely perturbed plasmalemmal localization of syntaxin-1 in Munc18-1/2 DKD PC12 cells (Han et al., 2011; Han et al., 2009), synaptic localization of syntaxin-1 is unaffected in Munc18-1 deficient neurons despite its severe defect in neurotransmitter release (Verhage et al., 2000). This implies possible role of Munc18-1 downstream of syntaxin-1 chaperoning. Moreover, overexpression of SNAP-25 has been shown to rescue the docking but not the secretion phenotype in Munc18 deficient chromaffin cells which strongly indicates a post-docking role of Munc18-1 (de Wit et al., 2009). Although different studies have suggested the potential role of Munc18-1 in the post docking stage of exocytosis, the direct evidence for such function has been lacking. Our analysis of the previously alleged ‘priming mutants’ (e.g. E59K, D34V/M38V) (Deák et al., 2009; Gulyás-Kovács et al., 2007) in the surface of domain 1 cleft suggested that their phenotypes are better explained by their reduced syntaxin-1 chaperoning activity. Therefore, the Munc18-1 mutant that selectively loses priming function is yet to be identified (Han et al., 2011). Currently, the priming stage of exocytosis is not well defined largely due to the abstruse mechanism underlying this process. In this paper, we refer to the stage of exocytosis that occurs after the docking of vesicles but prior to the actual fusion as the ‘priming stage’.
Several groups have suggested the direct binding of Munc18-1 to the SNARE complex as an underlying mechanism for Munc18-1 dependent SNARE-mediated fusion (Dulubova et al., 2007; Rickman et al., 2007; Rodkey et al., 2008; Shen et al., 2007). At least two different binding modes have been proposed for this direct interaction. The first binding mode represents the interaction between Munc18-1 hydrophobic pocket and syntaxin-1 N-peptide (Dulubova et al., 2007; Hu et al., 2007; Shen et al., 2007). However, syntaxin N-peptide binding to Munc18-1 is not highly selective, suggesting that other parts of the SNARE complex are involved in binding to Munc18-1 (Hu et al., 2010). The second binding mode is the binding between domain 3a of Munc18-1 and synaptobrevin-2 (Xu et al., 2010). The latter binding mode supports the potential role of domain 3a in the binding of Munc18-1 to the SNARE complex which may be crucial for directly stimulating SNARE-dependent exocytosis (Boyd et al., 2008). In the present study, we have thoroughly examined the functional significance of the domain 3a of Munc18-1 in syntaxin-1 chaperoning, dense-core vesicle docking and exocytosis. During this process, we discovered the mutants that selectively abolish the exocytotic function of Munc18-1 while preserving its syntaxin-1 chaperoning activity and the ability to support dense-core vesicle docking. These mutants represent the long-sought-after ‘priming mutant’ of Munc18-1.
Discovery of domain 3a insertion mutants that selectively lose the ability to restore secretion in Munc18-1/2 DKD PC12 cells while retaining the abilities to restore syntaxin level and localization
The syntaxin-1 chaperoning function of Munc18-1 is primarily mediated by the tight binary interaction between the ‘closed’ syntaxin-1 and Munc18-1 cleft that is formed by domain 1 and domain 3a (Misura et al., 2000). Previous studies have focused on elucidating the role of domain 1 through analyses of domain 1 mutants (Deák et al., 2009; Gulyás-Kovács et al., 2007; Han et al., 2011; Han et al., 2009). In an attempt to investigate the functional significance of domain 3a which is relatively understudied, we initially mutated highly conserved domain 3a residues that were suggested to make a contact with syntaxin-1. These mutants include: E278K, K314L/R315L (Kauppi et al., 2002), K332E, K333E and K332E/K333E (Fig. 1). During the course of our study, a paper was published showing that transient overexpression of a dominant negative Y337L mutation in domain 3a can inhibit secretion from PC12 cells (Boyd et al., 2008). Accordingly, we generated additional mutants which include Q336A/Y337L, Y337L/Q338A and Q336A/Y337L/Q338A (Fig. 1). These Munc18-1 mutants were expressed stably as Emerald-GFP (EmGFP) fusion proteins in our Munc18-1/2 DKD PC12 cells (clones D7 and D16) (Han et al., 2011; Han et al., 2009) for phenotypic analyses. Unexpectedly, all of these point mutants retained intact abilities to interact with syntaxin-1 and to restore syntaxin-1 expression and dense-core vesicle secretion (supplementary material Figs S1, S2). These results imply that domain 3a is highly resistant to the introduction of the point mutations. This functional resistance could be explained by the structural flexibility of domain 3a in that the region appears to be unstructured in the unbound solution state while it can possibly adopt a helical structure upon interaction with partners (Hu et al., 2010).
During the process of generating the K332E/K333E mutant in domain 3a using PCR-based site-directed mutagenesis, we unintentionally generated an insertion mutant. This mutant contains 39 extra residues (MPQYQDLSQMLEEMPQYQDLSQMLEEMPQYQDLSQMLEE) following the intended mutation of K332E/K333E (Fig. 1A). Despite the presence of the large insertion, this mutant, KE/39I, expressed well and efficiently restored syntaxin-1A level upon stable re-expression in the DKD cells as shown by immunoblot analysis which also includes EmGFP alone (negative control), Munc18-1 wild-type (WT)–EmGFP (positive control) and E278K mutant (D16 data in Fig. 2A; D7 in Fig. 2B). Note that our rescuing data are consistent between D7 and D16 cells thus have been used interchangeably in some figures. Furthermore, it restored the gross syntaxin-1A localization at the plasma membrane almost as effectively as the wild-type in our immunofluorescence microscopic analysis (Fig. 3, third panel). Nonetheless, this insertion mutant completely failed to restore exocytosis (D16 in Fig. 2C; D7 in Fig. 2D). This was the first indication that there is a potential Munc18-1 mutant that specifically loses the exocytotic ability while retaining syntaxin-1A chaperoning activity.
We further examined whether the KE/39I mutant has a dominant-negative effect by nonspecifically interfering with the secretory processes due to the presence of a large insertion. In addition, it has previously been reported that the transient overexpression of Y337L mutant inhibits secretion of wild-type PC12 cells (Boyd et al., 2008). In our system, we stably overexpressed Munc18-1 WT or its variants fused with EmGFP in the wild-type PC12 cells. EmGFP alone was also stably expressed as a negative control. We found that WT–EmGFP and KE/39I–EmGFP express ∼2–3 times more than the endogenous Munc18-1 while the chaperoning mutant (K46E/E59K)–EmGFP expresses at a similar level to the endogenous Munc18-1 (Fig. 2E). Importantly, we found that the stable overexpression of the KE/39I has no deteriorating effects on secretion from the wild-type PC12 cell (Fig. 2F). These results indicate that the KE/39I is not a dominant-negative mutant, but rather represents a pure loss-of-function mutant.
To minimize the potential artifact that a large insertion of 39 residues may cause on the overall structure of Munc18-1, we reduced the size of the insertion to five residues in the presence of K332E/K333E mutation (Fig. 4A). We found that the insertion of five residues with K332E/K333E mutation (KE/5I) is sufficient to disrupt the exocytotic function of Munc18-1 while retaining the ability to restore the expression level of syntaxin-1 (Fig. 4B,C). In addition, immunofluorescence microscopy demonstrated the restoration of the plasmalemmal localization of syntaxin-1 (Fig. 3, lowest panel). Furthermore, we examined whether the insertion of five residues alone, 5I(MPQEE) or 5I(MPQKK), without K332E/K333E mutations can inhibit exocytosis. We found that in both cases, the insertion of five residues alone strongly reduces the rescuing ability but does not abolish it while retaining their ability to restore syntaxin-1 level (Fig. 4D,E). This result suggests that although K332E/K333E point mutations alone do not cause functional impairment (supplementary material Fig. S2), it exacerbates the secretion rescue ability of the 5I mutants. Taken together, we concluded that the five residue insertion together with K332E/K333E mutation (KE/5I) is the minimal mutation that is necessary to completely abolish the secretion rescuing ability of Munc18-1 among the mutations tested in this study. Our KE/5I and KE/39I mutants represent the novel priming mutants which selectively impair secretion rescue ability while strongly retaining its ability to restore syntaxin-1 chaperoning activity.
KE/5I mutant restores the docking of dense-core vesicles
The loss of dense-core vesicle (DCV) docking has been found to be tightly coupled with the impaired DCV exocytosis in Munc18-1 deficient chromaffin cells (Voets et al., 2001). Our recent analysis of Munc18-1 domain 1 mutants demonstrated that Munc18-1-dependent syntaxin-1 localization is positively correlated with the efficiency of DCV docking to the plasma membrane (Han et al., 2011). For example, the chaperoning mutant (K46E/E59K) failed to restore docking of dense-core vesicles. Based on the observation that the KE/5I mutant effectively restores syntaxin-1 at the plasma membrane, we hypothesized that this mutant will restore DCV docking. Using electron microscopy, we analyzed the effect of the KE/5I mutant on docking of dense-core vesicles in the knockdown cells. The level of DCV docking was compared to the docking efficiency in the DKD (D7) PC12 cells rescued by EmGFP alone (negative control) or wild-type–EmGFP (positive control) (Fig. 5). Dense-core vesicles are defined as docked if they are localized within 50 nm from the plasma membrane. In the D7 cells rescued with EmGFP alone, only 6.6±0.9% (n = 47) of vesicles were docked while 48.3±2.5% (n = 48) of vesicles were docked in D7 cells upon re-expression of wild-type Munc18-1–EmGFP, indicating the clear rescue of docking phenotype (Fig. 5A,B). In D7 cells expressing KE/5I–EmGFP, 35.3±1.7% (n = 46) of vesicles were localized within 50 nm from the plasma membrane. Our one-way ANOVA analysis has indicated the significant difference in the proportion of docked vesicles in these three groups. Furthermore, Post-hoc analysis has suggested the significant difference between each group at the level of P<0.01. These results clearly indicate that despite its complete loss in the ability to rescue secretion, the KE/5I mutant retains the ability to restore docking of DCVs. The size of the cells was not significantly different among three different rescue conditions (Fig. 5D) while the total number of DCVs was slightly (∼20%) increased in the cells rescued by EmGFP compared to the cells rescued by the wild-type Munc18-1 or the KE/5I mutant (Fig. 5C). These findings further confirm that the KE/5I mutant selectively loses the ability to stimulate or prime exocytosis downstream of the docking of dense core-vesicles. To our knowledge, this is the first mutant that demonstrates the selective effect of Munc18-1 on DCV secretion independently from its effect on docking. Our results also suggest that Munc18-1 mediates the DCV docking through syntaxin-1 regulation. This seems to be consistent with the phenotype of adrenal chromaffin cells from the knockin mice of syntaxin-1B ‘open’ conformation mutant (Gerber et al., 2008).
The putative priming mutant and chaperoning mutant exhibit different properties in their genetic interactions with syntaxin-1
If the function of Munc18-1 can solely be explained by its chaperoning function, the secretory phenotypes caused by the loss of Munc18-1 should be at least partially restored in the presence of properly localized syntaxin-1A. To test this hypothesis, we have made an attempt to overexpress syntaxin-1A in Munc18-1/2 DKD cells through the lentiviral infection followed by blasticidin selection. Firstly, we confirmed that syntaxin-1A could indeed be overexpressed in the DKD cells as shown by immunoblot analysis (Fig. 6A). Interestingly, although majority of the overexpressed syntaxin-1A was accumulated in the perinuclear region, some were successfully localized along the plasma membrane (Fig. 6B). Nonetheless, overexpressed syntaxin-1A alone could not effectively restore the defective secretion of Munc18-1/2 DKD cells (D16 in the left panel of Fig. 6E; D7 in the left panel of Fig. 6F). This is another strong indication that the Munc18-1 function is extended beyond its syntaxin-1 chaperoning function.
We took advantage of the syntaxin-1A overexpressed Munc18-1/2 DKD cells to distinguish the phenotype of the ‘chaperoning mutant’ (K46E/E59K) and the ‘priming mutant’ (KE/39I). We hypothesized that if the chaperoning mutant still retains the priming function, this mutant should be able to rescue secretion to certain extent whereas the priming mutant will not be able to restore secretion even in the presence of properly localized syntaxin-1. To test this, we have further expressed K46E/E59K or KE/39I mutant in the presence of the overexpressed syntaxin-1A (Fig. 6C). As expected, the syntaxin-1 localization along the plasma membrane was much more efficiently restored upon introduction of the KE/39I mutant compared to the K46E/E59K mutant (Fig. 6D). In contrast to the impaired ability of the mutants to rescue syntaxin-1 localization, we observed significantly enhanced secretion in the syntaxin-1A overexpressed cells upon re-expressing K46E/E59K mutant compared to our negative control, EmGFP infected syntaxin-1A overexpressing cells (D16 in the right panel of Fig. 6E; D7 in the right panel of Fig. 6F). This indicates that the chaperoning mutant (K46E/E59K) at least partially retains the ability to stimulate/prime exocytosis in presence of plasmalemmal syntaxin-1. In contrast, the priming mutant (KE/39I) has failed to rescue secretion despite its ability to efficiently chaperone syntaxin-1 to the plasma membrane. This is additional evidence suggesting the importance of this region of domain 3a in the post-chaperoning role of Munc18-1. These results indicate that the chaperoning activity and the priming function are mediated through independent mechanisms which involve distinct region of Munc18-1.
KE/5I mutant retains the ability to bind monomeric syntaxin-1A in a stoichiometric manner while it significantly loses the ability to interact with the SNARE complex
To elucidate the underlying mechanisms by which KE/5I mutation leads to the loss of the priming function, we examined the ability of the KE/5I mutant to interact with monomeric syntaxin-1 or the SNARE complex. We first examined the ability of the priming mutants to bind to cytoplasmic syntaxin-1A (residues 1–264) using yeast two-hybrid assays (Fig. 7A). We found that the KE/5I mutant is able to bind to syntaxin-1A as efficiently as the wild-type and the K332E/K333E mutant. This is consistent with our finding that these priming mutants are able to restore syntaxin-1A level and plasmalemmal localization (Figs 2–f03,4). To correlate the syntaxin-1 chaperoning activity of Munc18-1 variants fused with EmGFP in the DKD cells with their ability to bind syntaxin-1, we also examined whether the Munc18-1 variants, including the priming mutant (KE/5I) and the chaperoning mutant (K46E/E59K), expressed in mammalian cell line HEK-293 cells can be quantitatively pulled down by GST-syntaxin-1A (residues 1–264). Using yeast two-hybrid analysis and isothermal titration calorimetry (ITC), we have previously shown that K46E single mutant significantly reduces the binding to syntaxin-1A whereas the K46E/E59K double mutant abolishes the interaction (Han et al., 2011). We found that our pull-down experiments using GST-syntaxin-1A mirrored the previous binding experiments; the pull-down of the K46E–EmGFP is reduced compared to the wild-type Munc18-1–EmGFP while the pull-down of the K46E/E59K-EmGFP is almost completely abolished (Fig. 7B). However, the K332E/K333E mutant and KE/5I mutant fused with EmGFP are quantitatively pulled down, which is comparable to the wild-type Munc18-1 fused with EmGFP (Fig. 7B). All the mutants tested are expressed at the comparable level in immunoblot analysis (Fig. 7C,D). These results demonstrate that the chaperoning mutant (K46E/E59K) loses the ability to bind to syntaxin-1A in a stoichiometric manner whereas the priming mutant (KE/5I) effectively retains it. These results further advocate for the intact chaperoning activity of the priming mutant, which occurs through syntaxin-1 interaction.
We then examined the ability of the priming mutant, KE/5I, to interact with the SNARE complex, as the binding of Munc18-1 to the SNARE complex has been suggested to be the potential mechanisms mediating the priming of exocytosis (Südhof and Rothman, 2009). To ensure the absence of monomeric syntaxin-1A or synaptobrevin-2, which can directly bind to Munc18-1 independently from the SNARE complex, we generated the SNARE complex centering the immobilized SNAP-25B on the column. Since a large tag such as GST can interfere with the formation of the SNARE complex (Dulubova et al., 2007), we expressed the full-length SNAP-25B as a strep (composed of eight amino acids, ∼1 kDa) -tagged protein in E. coli, purified and immobilized onto the strep-tactin Sepharose. Synaptobrevin-2 (residues 1–94) and syntaxin-1A (1–264) were expressed as GST-fusion proteins and GST-tag was removed by thrombin cleavage. Soluble synaptobrevin-2 and syntaxin-1A were incubated with SNAP-25B immobilized on the strep-tactin Sepharose and unbound synaptobrevin and syntaxin-1A were washed out from the Sepharose. SDS-PAGE analysis of the unboiled sample on the strep-tactin Sepharose showed the presence of the SNARE complex as well as the monomeric strep-SNAP-25 whereas the monomeric syntaxin-1A and synaptobrevin-2 were largely absent. Once the sample was boiled, the presence of syntaxin-1A and synaptobrevin-2 was evident. Please note that the boiling of the sample including immobilized SNAP-25B results in a partial elution of monomeric strep-tactin (∼14 kDa band, indicated by *** in Fig. 8A) from the strep-tactin Sepharose (Voss and Skerra, 1997) while the cytoplamic synaptobrevin-2 is present as a ∼13 kDa band in the SDS-PAGE gel stained with Coomassie Blue (Fig. 8A).
We then incubated the SNARE complex immobilized on Sepharose with Munc18-1 variants or BSA (negative control) in the binding solution of PBS containing 0.1% Triton X-100. We found that the wild-type Munc18-1 bound to the preformed SNARE complex while BSA did not, demonstrating the specificity of binding. Moreover, we found that F115E mutant of Munc18-1 could bind to the SNARE complex almost as effectively as the wild-type (Fig. 8). This was unexpected as the previous finding has shown that this mutation in Munc18-1 abolishes the interaction with the SNARE complex (Meijer et al., 2012). Under this condition, our KE/5I mutant exhibited significantly reduced binding to the SNARE complex compared to the wild-type Munc18-1 or F115E mutant (Fig. 8). These observations suggest that the mutation in domain 3a has a more severe effect in binding to the SNARE complex than the mutation (F115E) in the hydrophobic pocket. Our results indicate that the domain 3a of Munc18-1 plays a significant role in binding to the SNARE complex.
We have been strong proponents for the chaperoning function of Munc18-1 (Han et al., 2010; Han et al., 2011; Han et al., 2009). However, this function alone is insufficient to explain the striking phenotype of near complete loss of exocytosis in Munc18-1-deficient neurons, chromaffin cells and Munc18-1/2 DKD PC12 cells (Han et al., 2009; Verhage et al., 2000; Voets et al., 2001). For example, Munc18-1/2 DKD PC12 cells cannot be restored by simple overexpression of syntaxin-1A, even though this allows partial localization of syntaxin-1A at the plasma membrane (Fig. 6). Together with previous studies, this result strongly indicates the possibility of the essential function of Munc18-1 downstream of syntaxin-1 chaperoning and vesicle docking, perhaps by directly stimulating SNARE-dependent membrane fusion/exocytosis. However, due to the difficulty in isolating the ‘priming mutant’ of Munc18-1 that selectively impairs the exocytotic function while preserving the syntaxin-1 chaperoning activity, the direct evidence for its role in the priming stage of exocytosis has been limited. We speculate that this difficulty may be due to the structural flexibility of the domain 3a in which this region is unstructured in unbound state thus is not greatly affected by the point mutations (Hu et al., 2010). A point mutation in Munc18-1 that selectively loses priming activity has not been identified up to present. In this study, we present with the novel priming mutants, KE/5I and KE/39I, which have a combination of point and insertion mutations within domain 3a of Munc18-1 (Figs 1, 4). Similarly, a companion study published in this issue of Journal of Cell Science, revealed that a partial deletion of domain 3a results in impaired secretion without affecting syntaxin-1 trafficking to the plasma membrane (Martin et al., 2013). Together, these results clearly demonstrate the importance of domain 3a at the priming stage of exocytosis.
A comparison in the phenotypes of the newly isolated priming mutants with the previously characterized chaperoning mutant (K46E/E59K) allowed us to recognize distinct functions of Munc18-1 that are independent of each other. The observation that the chaperoning mutant can partially rescue exocytosis in the presence of overexpressed syntaxin-1A indicates that this mutant retains the priming activity. In contrast, the priming mutant which fails to rescue secretion defect still retains the abilities to restore syntaxin-1A expression, localization and dense-core vesicles docking (Figs 2–Fig. 3,Fig. 4,Fig. 5,6). Therefore, we suggest that these two functions of Munc18-1 are largely independent of each other. Our results that the priming mutant supports the docking of dense-core vesicles without supporting exocytosis (Figs 2, 4 and. 5) indicate the post-vesicle docking function of Munc18-1. Furthermore, it supports the notion that Munc18-1 indirectly regulates vesicle docking through syntaxin-1 regulation (Han et al., 2011).
The novel KE/5I mutant retains the ability to quantitatively bind to the monomeric syntaxin-1 while significantly impairing the ability to bind to the SNARE complex (Figs 7, 8). This is in agreement with the hypothesis that the priming function of Munc18-1 may be mediated through the direct interaction with the SNARE complex (Südhof and Rothman, 2009). However, it should be noted that the ability of the mutant to bind to the SNARE complex is not completely impaired (Figs 7, 8) while its ability to support secretion is entirely abolished (Figs 2, 4). Recent structural analysis of domain 3a suggested that this region can undergo structural changes upon binding of Munc18-1 with syntaxin N-terminal peptide (Hu et al., 2010). It is possible that the KE/5I mutant not only reduces the binding to the SNARE complex but also impairs the capability of this domain to undergo conformational change which enables the interaction between Munc18-1 and syntaxin N-peptide. Unexpectedly, we did not observe a strong reduction in the binding of Munc18-1 hydrophobic pocket mutant (F115E) with the SNARE complex in our experimental conditions. This is at odds with the previous studies showing a striking reduction of the same or the corresponding mutant in binding to the SNARE complex (Meijer et al., 2012). However, syntaxin N-peptide binding to Munc18-1 is not highly selective, suggesting that other parts of the SNARE complex are involved in binding to Munc18-1 (Hu et al., 2010). If the binding between Munc18-1 and the SNARE complex is mediated by more than one mechanism, the mutation in Munc18-1 hydrophobic pocket is expected to have a limited impact on the interaction with the SNARE complex. Our observation that the F115E mutant can still interact with SNARE complex may explain why the same mutant has exhibited none or limited effects on the ability to rescue neurotransmitter exocytosis in Munc18-1 deficient neurons (Meijer et al., 2012) or Munc18-1/2 DKD PC12 cells (Han et al., 2009; Malintan et al., 2009). Indeed, accumulating evidence has suggested that N-peptide of syntaxin plays a crucial role in exocytosis (Johnson et al., 2009; Khvotchev et al., 2007; Zhou et al., 2013). However, this is out of scope for this particular study.
In summary, our KE/5I and KE/39I mutants represent the true priming mutants which selectively impair secretion rescue ability while strongly retaining its abilities to restore syntaxin-1 chaperoning activity and dense-core vesicle docking. Our discovery of these novel mutants will facilitate the process of uncoupling the mechanisms by which Munc18-1 primes/stimulates exocytosis downstream of syntaxin-1 chaperoning and dense-core vesicle docking.
Materials and Methods
Parental pLVX-EmGFP-IRES-blast plasmid for lentivirus-mediated Munc18-1 expression was previously described (Han et al., 2011). pLVX-EmGFP-IRES-hygro was generated by replacing the blatsicidin resistance gene with the hygromycin resistance gene. psPAX2 was purchased from Addgene (Cambridge, MA) and pMD.G was a kind gift from Dr Tomoyuki Mashimo (University of Texas Southwestern Medical Center at Dallas, Dallas, TX). We obtained monoclonal antibodies against syntaxin-1 (clone HPC-1) (Barnstable et al., 1985) from Sigma Chemical (ON, Canada); Munc18-1 from BD Biosciences (Mississauga, ON, Canada), GFP from Clontech (Mountain View, CA) and GAPDH (clone 6C5) from Millipore (Billerica, MA).
Lentivirus-mediated expression of Munc18-1 variants in Munc18-1/2 DKD cells
Wild-type PC12 cells were maintained in DMEM (Invitrogen, Carlsbad, CA) containing 5% calf serum, 5% horse serum (both from HyClone Laboratories, Logan, UT), penicillin (100 U/ml)/streptomycin (0.1 mg/ml) (Sigma Chemical), 250 ng/ml amphotericin B (Sigma Chemical). Two clonal lines of Munc18-1/2 DKD cells (D7, D16) were maintained in the same medium containing puromycin (2.5 µg/ml) and G418 (700 µg/ml).
We have generated the lentivirus-mediated expression constructs of various Munc18-1 mutants so that these proteins stably express in the Munc18-1/2 DKD cells. The Munc18-1 gene with silent nucleotide mutations (SNM) (WT or its indicated mutant) was subcloned into the same site of pLVX-EmGFP-IRES-blast plasmid. This Munc18-1 expression plasmid was co-transfected with psPAX2 and pMD.G into HEK-293FT cells to generate recombinant lentiviruses that express Munc18-1 WT or its variant fused with EmGFP. The Munc18-1/2 DKD cells that were infected with lentiviruses expressing rescue proteins were selected with blasticidin (5 µg/ml).
[3H] Noradrenaline release assays from PC12 cells
PC12 cells were plated in 24-well plates; 3–4 days after plating, the cells were labelled with 0.5 µCi of [3H] noradrenaline (NA) in the presence of 0.5 mM of ascorbic acid for 12–16 hours. The labelled PC12 cells were incubated with the fresh complete DMEM for 1–5 hours to remove unincorporated [3H] NA. The cells were washed once with physiological saline solution (PSS) containing 145 mM NaCl, 5.6 mM KCl, 2.2 mM CaCl2, 0.5 mM MgCl2, 5.6 mM glucose and 15 mM HEPES, pH 7.4, and NA secretion was stimulated with 200 µl of PSS or high K+-PSS (containing 81 mM NaCl and 70 mM KCl). Secretion was terminated after a 15-minute incubation at 37°C by chilling to 0°C, and samples were centrifuged at 4°C for 3 minutes. Supernatants were removed, and the pellets were solubilized in 0.1% Triton X-100 for liquid scintillation counting.
Cell preparation for confocal immunofluorescence microscopy
Sterilized circular glass coverslips (0.25 mm in width, 1.8 cm in diameter) were placed in 2.2-cm wells within 12-well cell culture plates. The coverslips were then coated for 1 hour with poly-d-lysine (0.1 mg/ml) at room temperature. Cells were allowed to adhere to the coverslips overnight and then differentiated on the coverslips for 3–4 days in DMEM that contained 100 ng/ml nerve growth factor (NGF) (Sigma Chemical), 1% horse serum, 1% calf serum and penicillin/streptomycin. The cells were washed with phosphate-buffered saline (PBS), fixed for 15 minutes with PBS containing 4% paraformaldehyde (PFA). PFA was then removed from each well and cells were rinsed three times (10 minutes each time) with 1 ml of PBS per well. The fixed cells were then permeabilized with PBS containing 0.2% Triton X-100 and 0.3% BSA for 5 minutes, followed by three times wash with PBS. Non specific sites were blocked for 1 hour at room temperature in PBS containing 0.3% BSA. Primary antibodies against syntaxin-1 (HPC-1 diluted 1∶1000) were applied to the cell for 1 hour. After three washes in blocking buffer, rhodamine red-x-conjugated anti-mouse antibodies (diluted 1∶1000) (Jackson ImmunoResearch Laboratories, West Grove, PA) were applied for 1 hour. Samples were washed again three times in blocking buffer and mounted in Fluoromount-G reagent (Southern Biotechnology, Birmingham, AL). Immunofluorescence staining was recorded with a laser confocal scanning microscope (LSM510; CarlZeiss, Jena, Germany) with an oil immersion objective lens (63×).
Electron microscopy and analysis of docking of dense-core vesicles
The initial fixation was performed within the 10-cm dishes for 1 hour using a 3.2% glutaraldehyde and 2.5% paraformaldehyde fixative mixture (Karnovsky's Fixative) in 0.1 M cacodylate buffer (pH adjusted to 7.6). Cells were then pelleted in microcentrifuge tubes and fixed overnight with new fixative. The following day, the pellets were fixed in 1 mg/ml (1%) osmium tetroxide for 1 hour and En bloc staining was then performed by incubating with 1% uranyl acetate for 1 hour in dark conditions. Washed pellets were incubated successively in increasing concentrations of ethanol for dehydration and then infiltrated overnight with Spurr's resin (23.6 g NSA, 16.4 g ERL-4221, 5.72 g DER-736 and 0.4 g DMAE). After transferring the cell pellets to Beem capsules the capsules were incubated for 48 hours at 65°C. The plasticized pellets were sliced to ultrathin 80-nm sections, which were then mounted on copper grids for subsequent staining and viewing.
Grids mounted with the ultrathin cell sections were first etched by exposing the grids to 3% uranyl acetate for 45 minutes at room temperature. Grids were then washed and stained with lead citrate for 20 minutes. Grids were washed and dried again before loading onto Hitachi H7000 transmission electron microscope for viewing. Electron micrographs were taken of individual Munc18-1/2 DKD cells expressing EmGFP (negative control), wild-type Munc18-1–EmGFP (positive control) or KE/5I–EmGFP. These images were then used for analyzing the docking of dense-core vesicles in the control or the Munc18-1 variants expressing PC12 cells. Dense-core vesicles were identified within the single-cell electron micrographs as dark spots of radius between 60 and 120 nm. The distance of each vesicle from the plasma membrane was then calculated for each individual cell. The data from multiple single cell images (n = 46–48) within each control or Munc18-1 variants has been collated.
Lentivirus-mediated expression of syntaxin-1A and Munc18-1 variants in Munc18-1/2 DKD cells
Munc18-1/2 double-knockdown cells (clones D7 and D16) were infected with lentivirus containing syntaxin-1A expression plasmid in pLVX-IRES-blast. Infected cells were selected by blasticidin to establish stable syntaxin-1A overexpressed D7 or D16 clones. Expression of syntaxin-1A was confirmed by immunoblotting. Syntaxin-1A overexpressed DKD cells were further infected by the lentivius expressing Munc18-1 K46E/E59K–EmGFP, KE/39I–EmGFP or EmGFP alone from pLVX-IRES-hygro plasmids followed by hygromycin B selection. Munc18-1/2 DKD cells.
Yeast two-hybrid assays
Full-length WT Munc18-1with SNM (see above) or indicated mutant Munc18-1 (SNM) was subcloned into SmaI-PstI site of a bait vector, pLexN. A cytoplasmic domain (residues 1–264) of rat syntaxin-1A was subcloned into EcoRI-BglII site of a prey vector, pVP16-3 (Okamoto and Südhof, 1997). Yeast strain L40 (Vojtek et al., 1993) was transfected with bait and prey vectors by using the lithium acetate method (Schiestl and Gietz, 1989). Transformants were plated on selection plates lacking uracil, tryptophan and leucine. After 2 days of incubation at 30°C, colonies were inoculated into supplemented minimal medium lacking uracil, tryptophan and leucine and placed in a shaking incubator at 30°C for 2 days.
β-Galactosidase assays were performed as follows. Yeast cells were chilled on ice and harvested by centrifugation (2000 rpm for 5 minutes). The collected yeast cells were resuspended in 250 µl of breaking buffer [100 mM Tris-Cl, pH 8.0, 1 mM dithiothreitol (DTT) and 20% glycerol]. Then, glass beads (0.45–0.5 mm; Sigma Chemical) were added to the yeast suspension to a level just below the meniscus of the liquid, followed by 12.5 µl of phenylmethylsulfonyl fluoride stock solution (40 mM in 100% isopropanol stored at 4°C). The mixture was then vortexed six times at top speed in 15-seconds bursts. After that, another 250 µl of breaking buffer was added, mixed well and centrifuged for 1 minute. The liquid extract was withdrawn and transferred to new tubes. The extracted liquid was further clarified by centrifuging for 15 minutes in a micro-centrifuge. To perform the assay, 80 µl of the extract was added to 720 µl of Z buffer (60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4 and 2.7 ml/l β-mercaptoethanol, pH 7.0). The mixture was then incubated in a water bath at room temperature for 5 minutes. The reaction was initiated by adding 0.16 ml of stock solution (4 mg/ml o-nitrophenyl-β-D-galactoside in Z buffer; 4°C), and the reaction mixture was incubated at room temperature. The reaction was precisely terminated at the end of 7-minute incubation by addition of 0.4 ml of 1 M Na2CO3 stock solution in distilled water, and the optical density of the reaction mixture was measured at 420 nm by using a spectrophotometer. At the same time, the protein concentration in the extract was measured using Bradford dye-binding assay. A standard curve was prepared using serial dilutions of BSA dissolved in breaking buffer. 10 µl of the extract was added to 1 ml of the Bradford reagent (Bio-Rad Laboratories, Hercules, CA), and the change in colour was measured at 595 nm by using a spectrophotometer. The specific activity of β-galactosidase in the extract was calculated according to the following formula: (OD420×1.36)/[0.0045 × protein concentration (mg/ml)]×extract volume (0.08 ml)×7 minute], where OD420 is the optical density of the product o-nitrophenol at 420 nm. The factor 1.36 corrects for the reaction volume, and the factor 0.0045 is the optical density of 1 nmol/ml solution of o-nitrophenol. The unit of β-galactosidase-specific activity is therefore expressed as nanomoles per minute per milligram of protein.
Binding between Munc18-1 variants and monomeric syntaxin-1A
pGex-KG-rat syntaxin-1A (1–264) was transformed into BL21 (DE3) cells and the bacteria were grown at 37°C until confluent. Recombinant protein expression was induced by adding 50 µM isopropylthio-β-D-galactoside (IPTG) at 30°C for 3 hours. Cells were lysed using sonications in PBS containing 0.1% Triton X-100, 1 mM EDTA and protease inhibitors (10 µg/ml leupeptin and 10 µg/ml aprotinin). Supernatant portion was then mixed with glutathione agarose beads (Pierce Biotechnology, Rockford, IL). Next day, mixtures were washed extensively with PBS containing 20% sucrose, PBS containing 0.1% Triton X-100, and PBS in order. Samples were saved for binding with Munc18-1–EmGFP variants. EmGFP, wild-type Munc18-1–EmGFP, K46E–EmGFP, K46E/E59K–EmGFP, K332E/K333E–EmGFP and KE/5I–EmGFP were transfected into HEK-293FT cells. After 3 days, cells were lysed with KGlu buffer (20 mM HEPES, pH 7.2, 120 mM potassium glutamate, 20 mM potassium acetate, 2 mM EGTA) containing 0.1% Triton X-100 as a detergent. Portions were subjected to SDS-PAGE followed by western blot using anti-GFP antibody to confirm comparable expressions. After centrifugation, KGlu buffer plus detergent containing solubilized lysates were mixed with GST-syntaxin-1A overnight. Next day, mixtures were washed extensively with KGlu buffer containing 0.1% Triton X-100. Samples were then dried, and 2× SDS-PAGE sample buffer was added and subjected to SDS-PAGE followed by Coomassie Blue staining.
Binding between Munc18-1 variants and the SNARE complex
The expression plasmid called pET-strep was generated as follows. cDNA encoding GST and the following polylinker region of pGex-KG (Guan and Dixon, 1991) was amplified by PCR. The resulting PCR product (digested with NdeI and XhoI) was ligated to the pET-21a (Novagen) that was digested with NdeI and SalI, generating pET-GST plasmid. Then a NdeI-BamHI fragment encoding GST was replaced with the anneal oligos that encode strep-tag (WSHPQFEK) following the initial methionine, generating the pET-strep plasmid. cDNA encoding full-length human SNAP-25B was subcloned into the NcoI-SalI site of pET-strep. cDNAs encoding rat synaptobrevin-2 (1–94), rat syntaxin-1A (1–264) and full-length rat Munc18-1 variants were subcloned into the pGex-KG expression vector. The fusion proteins were expressed and purified as follows: A 50-ml preculture of BL21 (DE3) cells transformed with plasmids encoding recombinant proteins was grown overnight at 37°C in LB medium containing ampicillin. After overnight growth, this culture was used to seed 500 ml of LB medium containing ampicillin. The cells were further grown for 3 hours at 37°C then were induced with 100 µM IPTG at 30°C for 3 hours. After centrifugation, the cell pellet was resuspended in PBS containing 0.25 mM EDTA, PMSF, 10 µg/ml leupeptin and 10 µg/ml aprotinin. The cells were lysed by sonication. Cell debris was removed by centrifugation in a JA-20 rotor for 30 minutes at 15,000 rpm. To this lysate, 0.5 ml of strep-tactin Sepharose (IBA, GmbH, Germany) or glutathione-agarose (Pierce) were added and were rotated overnight at 4°C. Recombinant proteins immobilized onto the strep-tactin Sepharose or glutathione agarose were washed with PBS containing additional 0.15 M NaCl followed by final PBS wash. The GST-fused recombinant proteins were removed from the GST moiety by thrombin cleavage for 2–4 hours at room temperature. After centrifugation, the cleaved proteins were eluted in PBS containing PMSF.
The SNARE complex was produced by combining strep-SNAP-25, immobilized on the strep-tactin Sepharose, and soluble cytosolic syntaxin-1A and synaptobrevin-2 proteins in a 1∶1∶2 molar ratio and incubating overnight at room temperature in PBS containing PMSF, protease inhibitors (10 µg/ml leupeptin and 10 µg/ml aprotinin). The complex was isolated on the strep-tactin Sepharose via the strep-SNAP-25.
Binding experiment was performed by incubating molar excess of cleaved Munc18-1 variants or BSA (negative control) with SNARE complex immobilized on strep-tactin Sepharose in PBS containing 0.1% Triton X-100, PMSF and protease inhibitors for 2–4 hours at room temperature or overnight at 4°C. After extensive washing with PBS containing 0.1% Triton X-100, proteins bound to beads were analyzed by SDS-PAGE using standard procedures. The proteins were visualized by Coomassie Blue staining or Ponceau S staining. Immunoblot blot analysis was performed to detect Munc18-1 using anti-Munc18-1 antibody.
We thank Drs Herbert Gaisano (Toronto) and Tomoyuki Masimo (Dallas) for reagents used in this study. We also thank Drs Rory Duncan (UK), Frederic Meunier and Brett Collins (Australia) for critically reading the manuscript.
G.H. designed and performed the experiments and data analyses, made figures and took part in writing the manuscript. N.B. performed the experiments and made a figure. S.K. and L.H. performed the experiments. S.S. designed experiments, and was mainly responsible for writing the manuscript.
This research was supported by the Canada Research Chair Program, the Heart and Stroke Foundation [grant number T6700], Natural Sciences and Engineering Research Council of Canada [grant number 456042] and the Canadian Institute of Health Research [grant number MOP-93665].