The SNARE proteins VAMP/synaptobrevin, SNAP-25 and syntaxin are core components of the apparatus that mediates neurotransmitter release. They form a heterotrimeric complex, and an undetermined number of SNARE complexes assemble to form a super-complex. Here, we present a radial model of this nanomachine. Experiments performed with botulinum neurotoxins led to the identification of one arginine residue in SNAP-25 and one aspartate residue in syntaxin (R206 and D253 in Drosophila melanogaster). These residues are highly conserved and predicted to play a major role in the protein–protein interactions between SNARE complexes by forming an ionic couple. Accordingly, we generated transgenic Drosophila lines expressing SNAREs mutated in these residues and performed an electrophysiological analysis of their neuromuscular junctions. Our results indicate that SNAP-25-R206 and syntaxin-D253 play a major role in neuroexocytosis and support a radial assembly of several SNARE complexes interacting via the ionic couple formed by these two residues.
One of the best regulated physiological processes is the conversion of the electrical signals travelling along the nerve axon into chemical signals at the axon terminals in the form of neurotransmitters released in the intersynaptic space (Kandel et al., 2012). This process is termed neuroexocytosis and it is operated by a multiprotein nanomachine that mediates the fusion of the neurotransmitter-containing small synaptic vesicles (SSV) with the presynaptic membrane leading to neurotransmitter release. This nanomachine is triggered by cytosolic calcium and within hundreds of microseconds forms a membrane fusion pore (Kandel et al., 2012; Kasai et al., 2012). Its core components are the three SNARE proteins: VAMP/synaptobrevin, an integral membrane protein of SSV, SNAP-25 and syntaxin, localized on the cytosolic face of the presynaptic membrane (Jahn and Scheller, 2006; Südhof and Rothman, 2009; Sørensen, 2009; Südhof and Rizo, 2011). VAMP/synaptobrevin is present in nearly 70 copies per SSV (Takamori et al., 2006) and this high concentration is matched by the many copies of SNAP-25 and syntaxin present on the presynaptic membrane, as indicated by the fact that they account for as much as 9% of total synaptosomal proteins (Walch-Solimena et al., 1995). Furthermore, SNAP-25 and syntaxin form large clusters (50–70 copies) at the docking site of granules in PC12 cells (Knowles et al., 2010). In addition to the SNAREs, complexin and calcium-activated proteins, including synaptotagmin, Doc2 and Munc13 and Munc18 play a major role in the correct assembly, synchronization and triggering of the membrane fusion nanomachine (Jahn and Scheller, 2006; Südhof and Rothman, 2009; Sørensen, 2009; Südhof and Rizo, 2011; Neher, 2010; Malsam et al., 2012; Südhof, 2012a; Groffen et al., 2010; Boswell et al., 2012; Ma et al., 2013). Another set of proteins including Liprins, Rabs and Rab-interacting proteins, promote the tethering and priming of the SSV to the active zones of neurotransmitter release and may influence the assembly and regulation of the membrane fusion nanomachine (Jahn and Scheller, 2006; Südhof and Rothman, 2009; Sørensen, 2009; Südhof and Rizo, 2011; Südhof, 2012b; Momboisse et al., 2011). SSV are drawn close to the cytosolic face of the presynaptic membrane by the exergonic pairing of the coil-coiled domains of the three SNARE proteins, which form the SNARE complex in a stepwise mode (Sutton et al., 1998; Sørensen et al., 2006; Weber et al., 1998; Stein et al., 2009; Ellena et al., 2009; Hernandez et al., 2012; Gao et al., 2012). This folding process provides the proximity and the energy necessary for membrane fusion. The common view, based mainly on the use of in vitro model systems, is that few SNARE complexes are sufficient to drive membrane fusion via the formation of a hemifused membrane intermediate (Jahn and Scheller, 2006; Südhof and Rothman, 2009; Sørensen, 2009; Südhof and Rizo, 2011; Hua and Scheller, 2001; Shi et al., 2012; Mohrmann and Sørensen, 2012; Chernomordik and Kozlov, 2008). However, it is likely that a variable number of SNARE complexes is implicated in different events of membrane fusion in different cells and synapses, to adjust for different vesicle compositions and sizes, and different physiological requirements in the control of exocytosis.
The central role of the three SNARE proteins in neuroexocytosis is demonstrated by the fact that tetanus and botulinum neurotoxins (BoNT/A to BoNT/G) cause the neuroparalytic syndromes typical of tetanus and botulism by proteolytically cleaving these three proteins; in particular, BoNT/A removes nine residues from the SNAP-25 C-terminus, whereas BoNT/E removes 26 residues (Schiavo et al., 2000).
The structure of the neuroexocytosis nanomachine is not known and, in particular, there is no information on the critical protein–protein contact regions between the single SNARE complexes. Here, we have refined a molecular model based on the radial assembly of several SNARE complexes and have identified two residues that could play a major role in protein–protein contacts between adjacent SNARE complexes by forming an ionic couple. These residues, arginine 206 (R206) of SNAP-25 and aspartic acid 253 (D253) of syntaxin, were mutated in Drosophila and the synaptic functional consequences were analysed by electrophysiological methods. The results indicate that these residues play a major role in neuroexocytosis and support the multimeric model for the neuroexocytosis nanomachine where they are suggested to form an essential ionic couple.
A radial model for the SNARE supercomplex that mediates neuroexocytosis
A multimeric radial assembly of several SNARE complexes was proposed to account for the different outcome of the intoxication with BoNT/A and BoNT/E in neurons (Montecucco et al., 2005). In this model the SNARE complexes interact with one another mainly via the C-terminus of SNAP-25 and the segment included between amino acids 248 and 252 of syntaxin, thus explaining the strong dominant negative effect exerted by the cleavage of the C-terminus of SNAP-25 by BoNT/A (Keller et al., 2004; Raciborska et al., 1998; Huang et al., 1998). This model was refined as detailed in the experimental section and shown in Fig. 1. A close inspection of this model shows that R198 (the P1′ residue of the mouse SNAP-25 cleavage site by BoNT/A, which is homologous to R206 in D. melanogaster) and D250 of mouse syntaxin 1A (corresponding to D253 in Drosophila) are optimally positioned to form an ionic couple (Fig. 1B). Furthermore, these sites are highly conserved in metazoans (Fig. 1C). The energy released by the formation of an ionic couple in aqueous solvent at physiological ionic strength can be estimated to be in the range of 1–2 kcal/mol (Horovitz et al., 1990). In a SNARE super-complex, multiple ionic couples form and they do so on the two-dimensional membrane plane. This dimensional restriction greatly increases the probability of their formation. In addition, it should be considered that the major energy contribution to the formation of the SNARE super-complex is likely to be provided by the interaction with the additional proteins (i.e. complexin, synaptotagmin, etc.). Indeed, it is rather well established that zippering of the SNARE motif proceeds until nearly half of the four-helix bundle is formed and that, at this point, complexin binds and fixes and stabilizes the multimeric SNARE apparatus (Hernandez et al., 2012; Kümmel et al., 2011; Malsam et al., 2012; Li et al., 2011). Then SNARE coiling completes with the binding and positioning of synaptotagmin and the formation of the Arg–Asp ionic couple. The role proposed here for this ionic couple would be that of gearing neighboring SNARE complexes. On this basis, it can be predicted that the replacement of these residues will lower the functionality of SNARE super-complex with ensuing decreased neurotransmitter release.
Generation of Drosophila transgenic lines expressing either SNAP-25R206D or syntaxinD253R
To test this possibility, we decided to use conditions as close as possible to normal physiology by working in living animals and recording data from one of the most tightly controlled synapses: the neuromuscular junction (NMJ) (Kandel et al., 2012; Jan and Jan, 1976). We generated transgenic Drosophila lines bearing the targeted mutated forms of SNAP-25 or syntaxin (SNAP-25R206D and syntaxinD253R) under the control of the binary UAS/Gal4 system in a wild-type background. In particular, we employed an elav-Gal4 driver to express each of the mutated SNARE proteins in the fly nervous system. We did not attempt to express the mutant SNARE in a SNAP-25 null background because this would have led to the expression of SNAP-24, which would have interfered with our analysis (Vilinsky et al., 2002). The double SNAP-25 and syntaxin mutant, with a swapped ionic couple (i.e. negative charge on SNAP-25 and positive charge on syntaxin) predicted to be generate a fully functional NMJ was not generated owing to intrinsic genetic difficulties. The presence of SNAP-25R206D and syntaxinD253R does not hamper the formation of the SNARE complex, as control experiments showed that wild-type and mutated proteins are equally able to form SNARES in vitro (supplementary material Fig. S2); this result was expected since the mutated positions are external to the SNARE motif (Sutton et al., 1998; Sørensen et al., 2006).
Electrophysiological recordings at the neuromuscular junctions of wild-type and mutant Drosophila larvae
Neurotransmitter release was detected by electrophysiological recordings at the NMJs of 3rd instar larval body wall preparations. Fig. 2A shows that the presence of the mutant SNAP-25R206D in a wild-type SNAP-25 background substantially decreases the frequency of the miniature end plate potentials (mEPPs). A similar result was obtained from the analysis of the syntaxinD253R mutant in a wild-type background. The expression of either mutated isoform is not accompanied by a reduction of the quantum amplitude of mEPPs (Fig. 2B).
As expected from the results on mEPPs, also the evoked junctional potentials (EJPs) are reduced in animals carrying either SNAP-25R206D or syntaxinD253R along with their wild-type counterparts (Fig. 3A). This reduction of EJPs could be due to a reduced probability of assembly of a functional SNARE supercomplex and/or to a reduced lifetime of the pore through which the neurotransmitter is released or to a reduced quantal content. It is noteworthy that the expression of the wild-type proteins along with the endogenous ones does not alter the EJP amplitudes (Fig. 3A). The analysis of the 10–90% rise time of EJPs indicates that the lifetime of the pore is quite similar, if not identical, in all genotypes (Fig. 3B). At the same time an analysis of the mEPPs amplitude distribution indicates that the neurotransmitter content of the SSV is similar in the different animals tested here (Fig. 2B). Therefore, the present results indicate that SNAP-25R206D or syntaxinD253R do not reduce the probability of formation of the SNARE supercomplex, but its capability to mediate vesicle fusion.
The presence of SNAP-25R206D or syntaxinD253R at the neuromuscular junction of Drosophila does not alter its calcium sensitivity
The reduction in neurotransmitter release may result from alterations in the calcium dependency of the process. This possibility is suggested by the fact that the BoNT/A-induced blockade of neuroexocytosis can be partially rescued by increasing extracellular calcium (Sellin et al., 1983; Molgó et al., 1989; Lawrence and Dolly, 2002). To test this possibility, we performed measurements of EJPs at various external [Ca2+]. Fig. 4 shows that NMJs expressing SNAP-25R206D or syntaxinD253R display a [Ca2+] dependency equal to that of the wild-type NMJs. It is noteworthy that for each external [Ca2+] tested the same reduction of EJPs amplitude between the wild-type and each of the two mutants was recorded. This result is in keeping with the reported mapping of the interaction of the Ca2+ sensor synaptotagmin with SNAP-25 residues which was shown to be mediated by D179, D186, D193 of SNAP-25, residues which are at a distance from D198 (mouse numbering) (Sørensen et al., 2002; Zhang et al., 2002).
Another possible explanation for the reduced neurotransmitter release in the NMJ expressing SNAP-25 mutated at the C-terminus, or BoNT/A-cleaved SNAP-25, is that this segment is involved in endocytosis and its alteration would rapidly lead to decreased EJPs (but not mEPPs). However, this cannot be the case since BoNT/A cleavage of the SNAP-25 C-terminus blocks exocytosis, but not endocytosis (Neale et al., 1999).
The present identification of SNAP-25 R206 and of syntaxin D253 as residues essential for neuroexocytosis is an important step forward in the definition of the molecular structure of the neuroexocytosis nanomachine. Its importance is general and not confined to the radial model that has originated it, as one can conceive other oligomeric assemblies for the SNARE super-complex (Risselada and Grubmüller, 2012; Jahn and Fasshauer, 2012). Nor do the present data formally prove the existence of a radial assembly of SNARE complexes only, as other proteins could be implicated in the interaction with R206 and D253 within the NMJ. However, the present data are strictly pertinent to the studies using cell-free membrane fusion model systems (Ellena et al., 2009; Gao et al., 2012; Hua and Scheller, 2001; Shi et al., 2012; Pobbati et al., 2006; Lu et al., 2008; Domanska et al., 2009; Giraudo et al., 2009; Karatekin et al., 2010; van den Bogaart et al., 2011; Diao et al., 2012).
The radial model for the core of the neuroexocytosis nanomachine shown in Fig. 1, which seeded the present work, includes the possibility of a very rapid formation of a neurotransmitter conduit in its central part. In fact, this would result from a change of the lipid–protein interactions, triggered by a conformational switch caused by Ca2+ binding to synaptotagmin, which is known to interact directly with SNAREs and phosphatidylinositols (Sudhof, 2012a; Schiavo et al., 1996; Chapman, 2008). An important aspect of the molecular machines which drive exocytosis is the number of SNARE complexes involved in one membrane fusion event which has a bearing on the size of the initial pore (Hua and Scheller, 2001; Shi et al., 2012; Mohrmann and Sørensen, 2012). Studies performed with model membrane systems provide figures ranging from one to eight (Hua and Scheller, 2001; Mohrmann and Sørensen, 2012; Molgó et al., 1989; Domanska et al., 2009; Giraudo et al., 2009; Karatekin et al., 2010; van den Boogart et al., 2011; Diao et al., 2012), while experiments that used cracked or transfected chromaffin cells suggest that three/four SNARE complexes are involved in the release of catecholamines (Hua and Scheller, 2001; Mohrmann and Sørensen, 2012; Mohrmann et al., 2010). The present work does not address this point directly, but we cannot but mention that an optimal positioning of the SNAP-25 R206/syntaxin D253 ionic couple is obtained with an octameric radial SNARE super-complex (supplementary material Fig. S2B). This arrangement is in keeping with the high concentration of VAMP/synaptobrevin on SSVs (Takamori et al., 2006) and of SNAP-25 and syntaxin at the SSV docking sites (Walch-Solimena et al., 1995; Knowles et al., 2010) and fits well with a previous suggestion of five to eight SNARE complexes based on the systematic mutation of the transmembrane domain of syntaxin (Han et al., 2004). However, at variance from this work, the dimensions of the central part of the radial oligomer assembly are dictated, in the present model, by the packing of the individual SNARE complexes. In the central part and between the transmembrane portions of the SNARE complex units (Fig. 1) there is room for hundreds of lipid molecules which could well hemifuse as suggested by recent experimental evidence (Chernomordik and Kozlov, 2008; Hernandez et al., 2012; Jahn and Fasshauer, 2012). In addition, it has been shown that the highly positively charged segments connecting the SNARE domain to the transmembrane domains of VAMP/synaptobrevin and of syntaxin interact with negatively charged phospholipids (Chasserot-Golaz et al., 2010), and that phosphoinositides are essential per neuroexocytosis (Di Paolo and De Camilli, 2006). Thus the neuroexocytosis nanomachine is to be considered in its key central part as a lipid–protein complex, which, when triggered by a conformational change of a Ca2+-binding protein, rapidly opens a lipid–protein pore. The dimensions of the pore in the model of Fig. 1 are compatible with experimental estimates (Breckenridge and Almers, 1987). With the radial model proposed here it is straightforward to envisage both the ensuing pore expansion, which is known to take place during neurotransmitter release (Breckenridge and Almers, 1987), and the recruitment of the proteins involved in the process of endocytosis that follows (Haucke et al., 2011).
Materials and Methods
The analysis of primary structure conservation among the three SNARE proteins was performed on a set of sequences retrieved by a BLAST search on the UNIPROT database (www.uniprot.org) for all the SNAP-25 and syntaxin 1 proteins belonging to the metazoan kingdom. The sequences identified were aligned using T-Coffee (Di Tommaso et al., 2011). A conservation score was calculated using a Bayesian method as implemented in the ConSurf server (Ashkenazy et al., 2010). Not surprisingly, most of the conserved residues belong to the protein–protein interface within single SNARE complexes. However, some conserved amino acids point towards the external side of the four-helix bundle, among them, Asp250 and Arg198 (mouse numbering in syntaxin 1 and SNAP-25, respectively).
Construction of the super SNARE complex radial model
Analysis of dose/response measurements of neurotransmitter release as a function of SNAP-25 cleavage by Botulinum type A and E neurotoxins suggested a radial arrangement for SNARE complexes during neuroexocytosis (Montecucco et al., 2005). Therefore, a model was generated using the cytosolic domain of a SNARE complex solved by X-ray crystallography (Chen et al., 2002). Transmembrane segments were modeled as alpha-helices according to structure predictions and connected to the cytosolic domain by unstructured peptides. This model of the complete SNARE complex was used as a starting structure (supplementary material Fig. S1A).
Different radial arrangements containing an increasing number of single SNARE complexes (4, 6 and 8) were generated by symmetry operations. The only constraint imposed on the models was to minimize the distance between SNARE complexes without generating steric clashes. The potential energy of the resulting models was optimized performing 100 minimization steps using the steepest descent algorithm followed by 1000 steps of conjugated gradient using the AMBER99 force field (Case et al., 2012). Electrostatic interactions were calculated using a cutoff of 2 nm within the framework of the Generalized Born model for implicit solvation considering the electrostatic screening of a monovalent salt at a concentration of 150 mM as implemented in AMBER (Case et al., 2012).
For each of these models we checked for possible inter-SNARE interactions between conserved residues. As a result of the radial arrangement, the distance between D250 and R198 (mouse numbering in syntaxin 1A and SNAP-25, respectively) decreases from 1.4 nm in the tetrameric arrangement to 0.5 nm in the octameric one (supplementary material Fig. S1B).
Site-directed mutagenesis and expression of SNARE proteins
pGEX2T-SNAP-25 and pGEXKg-syntaxin1a (mouse sequences) were used as templates in a PCR amplification performed using the QuickChange site-directed mutagenesis kit (Stratagene). For the SNAP-25 R198D mutation (mouse numbering) the forward primer was 5′-ATTGATGAAGCCAACCAAGACGCAACAAAGATGCTGGGAAGTGGT-3′, for syntaxin1a D250R the forward primer was 5′-CGTGGAGAGGGCCGTGTCTCGAACCAAGAAGGCCGTCAAGTACC-3′. PCRs were performed under the following conditions: initial denaturation for 60 seconds at 95°C, amplification for 18 cycles of 50 seconds at 95°C, 50 seconds at 60°C and 6 minutes 20 seconds at 68°C, final extension for 7 minutes at 68°C. After digestion of the parental DNA for 1 hour at 37°C with DpnI, the amplified plasmids were transformed into E. coli XL-1 Blue competent cells. The presence of the mutations was confirmed by DNA sequencing. The mutated constructs were transformed into E. coli BL21 DE3 and the recombinant proteins expressed as GST fusion proteins and isolated as previously described (Tonello et al., 1999). Briefly, expression of the GST fusion proteins was induced for 3 hours at 30°C with 1 mM IPTG (isopropyl-β-D-thiogalactoside) and proteins were isolated on a glutathione–Sepharose 4B affinity column (Amersham Biosciences) according to the manufacturer's instructions. Resin-bound GST–SNAP-25R198D and GST–SYXD250R were eluted in a buffer containing 20 mM reduced glutathione and dialysed overnight in 10 mM HEPES buffer pH 7.4 with 150 mM NaCl. Protein purity was checked by SDS-PAGE on a 4–12% gel (Invitrogen). Recombinant syntaxin 1A cytoplasmic domain (1–261) and VAMP2 cytoplasmic domain (1–96), a kind gift from B. Davletov (LMB, Cambridge, UK), were expressed and purified as described previously (Hu et al., 2002). GST-SNAP-25WILDTYPE was expressed and purified as detailed before (Tonello et al., 1999).
RNA extraction and PCR
SNAP-25 and syntaxin wild-type (WT) and mutants (SNAP-25R206D and syntaxinD253R) mRNA expression levels were assessed in ElavGAL4/UAS–SNAP-25WT, ElavGAL4/UAS–SNAP-25R206D, ElavGAL4/UAS–syntaxinWT and ElavGAL4/UAS–syntaxinD253R third instar larvae by semi-quantitative PCR. Total RNA was extracted from 30 brains (from each line) of third instar larvae with TRIzol (Life Technologies) following the manufacturer's instructions and then resuspended in RNase-free water. For each sample, 1 µg of RNA was used for the first-strand cDNA synthesis, employing oligonucleotides dT and SuperScript III (Life Technologies), according to the manufacturer's instructions.
PCR reactions were performed with the following oligonucleotides: sense SNAP-25WT, 5′-ATATGGGCTCTGAGCTGGAA-3′; antisense SNAP-25WT, 5′-TTACTTTAATAGTTGATGTGCCCT-3′; antisense SNAP-25R206D, 5′-TTACTTTAATAGTTGATGTGCGTC-3′; sense syntaxinWT, 5′-GGCCATGCTGGTGGAGT-3′; antisense syntaxinWT, 5′-GAGCGCCTTCTTGGTGTC-3′; antisense syntaxinD253R, 5′-AGCGCCTTCTTGGTGCG-3′; sense rp49, 5′-ATCGGTTACGGATCGAACAA-3′; antisense rp49, 5′-GACAATCTCCTTGCGCTTCT-3′.
The size of the PCR products were 119 bp in the case of SNAP-25 isoforms, 109 bp for syntaxin isoforms and 164 bp for rp49 (housekeeping gene). Samples were analysed by electrophoresis (1.7% agarose gel) and the intensities of the bands were quantified using Image J (v 1.45).
Wild-type, SNAP-25R206A isoform expression was calculated relative to the median values of the rp49 housekeeping mRNA. Four different evaluations were performed and the average value for SNAP-25R206D transgene expression was 2.5±0.3 times that of the SNAP-25WT present in the same transgenic line, whereas syntaxinD253R transgene was found to be 0.4±0.1 times the expression of syntaxinWT.
In vitro assembly of the SNARE complex
The reaction for assembly of the SNARE complex was performed as described previously (Fasshauer et al., 1998). Recombinant VAMP2 and syntaxin 1A were incubated with either GST-SNAP-25WT or GST–SNAP-25R198D, recombinant VAMP2 and GST-SNAP-25WT with GST-syntaxinD250R for different time periods at room temperature in 20 µl of 20 mM HEPES buffer pH 7.2, 150 mM NaCl, 1 mM Na2EDTA, 1 mM dithiothreitol, 0.8% β-octylglucoside. 25 pmol of each SNARE protein were employed. The reactions were stopped by the addition of sample loading buffer. The solutions were then incubated at room temperature or boiled at 100°C for 15 minutes (boiling disrupts the SNARE complex). Samples were then subjected to SDS-PAGE using NuPAGE 4–12% Precast gels (Invitrogen), followed by Coomassie Blue staining; images were collected with ChemiDoc™ XRS (BioRad).
Construction of transgenic Drosophila lines
SNAP-25 [SNAP-25R206D] and syntaxin isoforms [syntaxinD253R and syntaxinWT] were produced by PCR using primers carrying point mutations as necessary. The general strategy is the one used before (Megighian et al., 2010). The ORFs encoding for SNAP-25 and for syntaxin were amplified from the full-length cDNA derived from w1118 flies using the primer pairs listed below. Snap-25 and syntaxin isoforms (wild-type and mutant) were produced by PCR using primers carrying point mutations as required (see primer pair sequences listed below). In the case of syntaxin, in order to introduce the D253R point mutation, two sequential PCR reactions were performed because the amino acid to substitute (Asp253) is in a central position; so two fragments of the gene were generated, one from the 5′-terminus to the region of Asp253 and the other one from this region to the 3′-terminus. The two products were then ligated to generate the syntaxinD253R-encoding and the syntaxinWT-encoding sequences.
Primer pairs used for the amplification of wild-type sequences and for site directed mutagenesis of SNAP-25 and syntaxin are reported below. Site-specific mutagenesis: sense and antisense primers have at their 5′ ends the restriction sites for EcoRI and XhoI, respectively (underlined nucleotides).
Oligonucleotides are in 5′–3′ orientation; restriction sites are underlined; mutated sites are in bold:
EcoRI_SNAP-25_WILDTYPE_F: cggaattcATGCCAGCGGATCCATCTGAAG; XhoI_SNAP-25_WILDTYPE_R: ctctcgagTTACTTTAATAGTTGATGTGCC; XhoI_SNAP-25_R206D_R: ctctcgagTTACTTTAATAGTTGATGTGCGTCTTGATTAGC; XhoI_SNAP-25_delta_R: ctctcgagTTATTGATTAGCAACTGCTATCCGCGCT; NotI_syntaxin1A_WILDTYPE_F: ccgcggccgcATGACTAAAGACAGATTAGCCGCTC; XhoI_syntaxin1A_WILDTYPE_R: ggctcgagTTACATGAAATAACTGCTAACATATGAGG; Syntaxin1A_D253R_F: CAACTCAGCGCACCAAGAAGG; Syntaxin1A_D253R_R: CCTTCTTGGTGCGCTGAGTTG.
In addition to the above pairs of primers, we also employed the following pair of primers to amplify a sequence encoding for a C-terminally truncated isoform of SNAP-25 which corresponds to the product generated by BoNT/A cleavage of the C-terminal 9 amino acids of SNAP-25:
EcoRI_SNAP-25_WILDTYPE_F: cggaattcATGCCAGCGGATCCATCTGAAG; XhoI_SNAP-25_delta_R: ctctcgagTTATTGATTAGCAACTGCTATCCGCGCT.
Amplification of the desired target sequence was obtained as follows. After the initial denaturation step at 94°C for 3 minutes, amplification was achieved through 35 cycles at 94°C for 30 seconds, 55°C for 30 seconds, and 72°C for 2 minutes. A final extension reaction was carried out for 7 minutes at 72°C. The obtained 655 bp PCR products were ligated into the pCR2.1-TOPO vector (Invitrogen, USA) which was used to transform One Shot TOP10 E. coli cells. Positive clones were detected by β-galactosidase screening and sequencing (BRM Genomics, Italy). Using conventional restriction enzyme digestion techniques, sequences were extracted with EcoRI and XhoI and ligated into the pUAST Drosophila transformation vector. The resulting plasmids, harboring each of SNAP-25 and syntaxin wild-type and mutated isoforms were again sequenced to ensure that the mutations occurred only at the position of interest and to verify the proper orientation of each ORF inside the vector. These plasmids were then used for embryo transformation.
P-element-mediated germline transformation was performed using Drosophila embryo injection services (Best Gene Inc., CA, USA; Transflyer, Ferrara, Italy). Briefly, the constructs with the w+ marker were injected into w1118 embryos. Several independent transformant lines were established for each construct. Lines carrying the UAS–SNAP-25WT, UAS–SNAP-25R206D, UAS–SNAP-DELTA, UAS–syntaxinWT and UAS–syntaxinD253R transgenes were chosen on the basis of their homozygous viability. The different SNAP-25 and syntaxin allelic isoforms were expressed in a SNAP-25 and syntaxin wild-type background using the GAL4/UAS system. In particular, pan-neuronal expression of the desired SNARE isoform was obtained in the progeny of crosses involving elav-Gal4 virgin females and males homozygous for one of the above constructs (supplementary material Fig. S3).
Pan-neuronal expression of the SNAP-25 isoform bearing a C-terminal deletion of nine amino acids proved to be embryonic lethal and was not investigated further. The double SNAP-25 and syntaxin mutant, with a swapped ionic couple (i.e. negative charge on SNAP-25 and positive charge on syntaxin) was not generated owing to intrinsic genetic difficulties.
Flies were raised on a standard yeast–glucose–agar medium and were maintained at 23°C, 70% relative humidity, in a 12-hour light∶12-hour dark cycle. Experiments were carried out on F1 third instar larvae.
Experiments were performed at 20–22°C on third instar larval body wall preparations dissected in Ca2+-free HL3 saline (Stewart et al., 1994) and pinned on the silicone-coated surface (Sylgard 184; Dow Corning, USA) of a 35 mm Petri dish (Bolatto et al., 2003). After dissection, Ca2+-free HL3 saline was replaced with 0.75 mM Ca2+ HL3 (see below) and body walls were left to incubate in this saline for at least 15 minutes before starting intracellular recordings.
Electrophysiological recordings were performed on fibres 6 or 7 of abdominal segment 3 or 4 using intracellular glass microelectrodes (1.2 mm o.d.; 0.69 mm i.d.; 10–12 MΩ resistance; Science Products, Germany) filled with a 1∶2 solution of 3 M KCl and 3 M CH3COOK. Fibres with a membrane resting potential lower than −60 mV were discarded. In each fibre both spontaneous and evoked neurotransmitter release were recorded. No more than one fibre for each larval body wall was utilized for electrophysiological recording. Signals were amplified in current-clamp mode by a voltage-clamp amplifier (NPI, Germany).
Both spontaneous and evoked neurotransmitter release were recorded under current-clamp conditions. Resting membrane potential was clamped at −70 mV. Spontaneous neurotransmitter release was analyzed by recording miniature end-plate potentials (mEPPs); spontaneous release events were recorded for 240 seconds (supplementary material Fig. S4). As evoked release can influence spontaneous neurotransmitter release, each activity was recorded in a separate set of experiments.
Evoked neurotransmitter release was analysed by intracellularly recording evoked junction potentials (EJPs). EJPs were elicited by stimulating at 0.5 Hz (stimulus duration 0.1 milliseconds, 1.5 threshold voltage) the single segmental nerve innervating abdominal segment A3 or A4 (supplementary material Fig. S5). The segmental nerve was stimulated using a suction microelectrode, filled with extracellular bathing solution and connected to a stimulator (S88, Grass, USA) via a stimulus isolation unit (SIU5, Grass, USA) in a capacitative coupling mode.
Ca2+ dependence of EJPs was analysed by replacing Ca2+-free dissection saline with a new one containing 0.5 mM Ca2+. After the appropriate incubation time (at least 15 minutes), EJPs were recorded for 60 seconds. Then, leaving the microelectrode in the same fibre, the bath saline was again replaced with a new one containing 0.75 mM Ca2+, leaving body walls to incubate for the appropriate time (see above) before starting EJPs recording. The procedure was repeated for each of the tested [Ca2+] (0.5–0.75–1.5–3 mM). In each fibre, fifteen EJPs were evoked for each of tested [Ca2+]. Particular care was taken to avoid the microelectrode exiting from the fibre or fibre membrane damage; whenever this happened, the sample was discarded.
Amplified signals were digitized using a digital A/C interface (National Instruments, USA) and then fed to a PC for both on-line visualization and off-line analysis using appropriate software (WinEDR, Strathclyde University; pClamp, Axon, USA). Stored data, were analyzed off-line using appropriate software (pClamp, Axon, USA). Statistical analysis and graph construction were carried out using Prism software (GraphPad, USA).
We thank Dr B. Davletov (University of Sheffield, UK) for the kind gift of SNARE proteins.
↵‡ Present address: Department of Cellular Neurobiology, University of Goettingen, Goettingen 37077, Germany
A.M., M.Z., S.P., O.R., C.M. designed the project and the experiments, S.P. performed the computational biology experiments, M.Z. and D.Z. designed mutants and prepared mutant Drosophila, M.R. performed molecular biology experiments, A.M. and M.S. performed electrophysiology and analysed data together with the other authors, A.M., M.Z., S.P., O.R., C.M. wrote the paper.
This work was supported by the grant ‘Synaptic functions and role of glial cells in brain and muscle diseases’ from Fondazione Cariparo and from the University of Padova, Progetto Strategico to C.M. and Progetti di Ateneo and Progetti di Ricerca di Interesse Nazionale (PRIN) to O.R., and by PRIN to A.M.
Supplementary material available online at http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.123802/-/DC1
- Accepted April 23, 2013.
- © 2013. Published by The Company of Biologists Ltd