Drosophila synaptotagmin aspartate residue 229 was mutated to glutamate. Oligonucleotides (cttggtctcgaacttcttcttcttgtcgggcagcaagtacaccttgacatagggctccgaggtac and ctcggagccctatgtcaaggtgtacttgctgcccgacaagaagaagaagttcgagac) were used to create a mutant double stranded DNA fragment with Kpn I and Sty I overhangs which was ligated into a wild-type synaptotagmin cDNA construct in pBluescript II KS (Stratagene, Agilent Tech Inc., Santa Clara, CA), sequenced, subcloned into a pUAST vector to place the mutant syt gene under the control of the UAS promoter (Brand and Perrimon, 1993) and these were sent to Best Gene, Inc. (Chino Hills, CA) to transform Drosophila.

Expression of the transgene was localized to the nervous system using elavGal4 to drive pan neuronal expression of the UAS-syt transgenes (Brand and Perrimon, 1993; Yao and White, 1994). The syt^null mutation used was syt^AD4 (DiAntonio et al., 1993). Standard genetic techniques were used to cross the transgenes into the syt^null background in order to express the transgene in the absence of endogenous synaptotagmin 1 (Loewen et al., 2006a). No gender selection was employed, thus a mix of male and female larvae were used in all experiments. Experimental flies were: yw; syt^nullelavGAL4/syt^null; P[UAS syt^A-D229E]/+ (P[syt^A-D2E], transgenic mutant) and yw; syt^nullelavGAL4/syt^null; P[UAS syt^WT]/+ (P[syt^WT], transgenic control).

A ClustalW2 sequence alignment was performed on the C₂A domain of the following Ca²⁺-binding synaptotagmin isoforms: syt 1 from Drosophila melanogaster (NP_523460.2), Apis mellifera (NP_001139207.1), Manduca sexta (AAK01129.1), Loligo pealei (BAA09866.1), Caenorhabditis elegans (NP_495394.3), Gallus gallus (NP_990502.1), Mus musculus (NP_033332.1), Rattus norvegicus (NP_001028852.2), and Homo sapiens syt 1 (NP_001129277.1), syt 2 (NP_001129976.1), syt 3 (NP_001153801.1), syt 5 (NP_003171.2), syt 6 (NP_995320.1), syt 7 (NP_004191.2), syt 9 (NP_445776.1), and syt 10 (NP_945343.1).

The D2E mutation in Fig 1C was modeled using the mutagenesis plugin in PyMOL (The PyMOL Molecular Graphics System, Version 1.3, Schrödinger, LLC). Asp-178, from the high resolution rat synaptotagmin 1 C₂A X-ray structure (1RSY), was changed to a glutamate. The rotamer with the fewest number of collisions was selected for the figure.

EJPs and mEJPs were recorded using standard techniques (Reist et al., 1998; Paddock et al., 2011) from L3 muscle fiber 6 of abdominal segments 3 and 4 in HL3 saline containing 70 mM NaCl, 5 mM KCl, 20 mM MgCl₂, 10 mM NaHCO₃, 5 mM Trehalose, 115 mM sucrose, 5 mM HEPES, pH 7.2, and 1.5 mM CaCl₂ unless otherwise indicated. Dissections were performed in Ca²⁺-free HL3 saline. Fibers were impaled with a 10–40 MΩ recording electrode containing 3 parts 2 M potassium citrate to 1 part 3 M potassium chloride. The resting membrane potential of each fiber was maintained at −55 mV by passing a bias current. To evoke EJPs, segmental nerves were stimulated with a suction electrode filled with 1.5 mM Ca²⁺ HL3. The Ca²⁺ dependence curve was generated by averaging 10 EJPs recorded at 0.5 Hz from fibers bathed in HL3 containing Ca²⁺ concentrations ranging from 0.6 to 5.0 mM. Recordings in multiple Ca²⁺ concentrations were made from each muscle fiber. 10–13 fibers were recorded at each Ca²⁺ concentration in each genotype with the exception of 1.5 mM Ca²⁺. 28–34 fibers were recorded at 1.5 mM Ca²⁺ since most recording sessions included this concentration. All events were collected using an AxoClamp 2B (Molecular Devices) and digitized using a MacLab4s A/D converter (ADInstruments). The Ca²⁺-dependence data were fit with the Hill equation using Kaleidagraph software. The Ca²⁺ cooperativity coefficient was estimated from the slope of a double-log plot of EJP amplitude versus Ca²⁺ concentration. EJPs were recorded in Scope software and mEJPs were recorded in Chart Software (ADInstruments).

Western analysis was used to determine levels of transgene expression. The central nervous systems of individual third instar larvae were loaded 1 CNS/lane and blots were probed with an anti-synaptotagmin antibody [Dsyt-CL1 (Mackler et al., 2002)] and an anti-actin antibody (MAB 1501, Chemicon) using standard techniques (Loewen et al., 2001). Actin levels were used to normalize for equal protein loading; the synaptotagmin:actin signal ratio was determined for each lane, then normalized to the mean synaptotagmin:actin ratio of the P[syt^WT] lanes on each blot to allow comparison of signal between multiple blots. Transgenic synaptotagmin was localized by immunolabeling third instar larvae with Dsyt-CL1 using standard techniques (Mackler and Reist, 2001). Neuromuscular junctions were visualized on a Zeiss Axioplan 2 upright digital imaging microscope.

CD spectra were measured with an AVIV stop flow Circular Dichroism spectropolarimeter at 192 to 260 nm using a 1 mm path-length cell. Samples containing 0.2 mg/ml of either wild type or mutant C₂A domains were assayed at 22 °C. For corrections of baseline noise, the signal from a blank run of buffer (50mM sodium phosphate) was subtracted from all the experimental spectra.

Isothermal titration calorimetry (ITC) data was generated using a GE Healthcare MicroCal iTC₂₀₀ which can in principle detect dissociation constants from 10 mM to 1 nM. Wild type and mutant C₂A domains were dialyzed overnight in ITC buffer (50 mM HEPES, pH 7.4, 200 mM NaCl, and 10% glycerol). ITC buffer was further used to make calcium chloride stock and protein dilutions. Prior to each experiment, samples were degassed and cooled to experimental temperature. Heat of binding was measured from thirty 1.1 μl injections of calcium chloride into sample cells containing 1.6 mg/ml of proteins of interest at 15° C. Baseline corrections, for heat of dilution, were made by subtracting the signal of calcium chloride injections into buffer from all experimental traces. Data was analyzed using GE Healthcare MicroCal iTC₂₀₀ Origin data software package.

A Student t-test was used to determine whether any statistically significant differences existed between the two independent P[syt^A-D2E] lines (line 3 and line 5). One-way ANOVA and Tukey range tests were used for statistical comparisons of P[syt^WT] and the two P[syt^A-D2E] lines.

C₂ domains are a common functional motif found in multiple proteins. Although not all C₂ domains are regulated by Ca²⁺, many C₂ domains mediate a Ca²⁺-dependent translocation of the protein to membranes (Nalefski and Falke, 1996; Cho and Stahelin, 2006). In these proteins, Ca²⁺ is coordinated by five negatively-charged residues located in loops 1 and 3 of the C₂ domain beta-sandwich structure [e.g., see Fig. 1A,B: D1–5 (Sutton et al., 1995; Nalefski and Falke, 1996; Ubach et al., 1998)]. Both aspartate and glutamate residues are utilized in the Ca²⁺-binding motifs of diverse C₂ domains (Nalefski and Falke, 1996). To assess whether Ca²⁺ binding by C₂A could be supported by either aspartate or glutamate residues, we compared the sequence of C₂A domains of synaptotagmin isoforms that bind Ca²⁺. A sequence alignment revealed that glutamate is excluded from these key positions. In the C₂A Ca²⁺-binding motifs of synaptotagmin 1 from many species, as well as of all of the human synaptotagmin isoforms that bind Ca²⁺, all 5 aspartate residues are 100% conserved (Fig. 1A) suggesting that glutamate residues in these positions would not provide full function.

Examination of the crystal structure of rat syt 1 C₂A demonstrates that the deepest parts of this Ca²⁺-binding pocket are spatially quite restricted (Sutton et al., 1995). Glutamate, like aspartate, is negatively charged, but glutamate possesses a significantly larger molecular volume [91 ų for Asp vs. 109 ų for Glu (Creighton, 1994)]. Coupled with the exclusion of glutamate from C₂A Ca²⁺-binding motifs, this observation suggested that a D→E mutation may impair Ca²⁺ binding. In the crystal structure of rat syt 1 C₂A, both aspartate residue 178 (Fig. 1A,B: D2) and aspartate residue 230 (Fig. 1A,B: D3) are well ordered and located deep in the Ca²⁺-binding pocket (Sutton et al., 1995). To assess whether a D→E mutation in either of these locations may occlude the Ca²⁺-binding pocket, we modeled the C₂A^D2E and C₂A^D3E mutations in silico. Results suggested that the C₂A^D2E mutation might occlude Ca²⁺-binding site 1 (Fig. 1C: Ca1 site). If true, this novel mutation would directly assess the importance of the electrostatic repulsion provided by C₂A in inhibiting vesicle fusion since it would inhibit Ca²⁺ binding without reducing the negative charge of the pocket (Fig. 1C: red regions).

To directly measure Ca²⁺ binding to the C₂A domain of Drosophila synaptotagmin 1, we utilized isothermal titration calorimetry (ITC). The titration data indicate three Ca²⁺-binding sites in Drosophila C₂A (Fig. 2A, Table 1). The intrinsic Ca²⁺ affinities of the three sites measured by ITC (Table 1, K_Ds = 20.7 ± 3.9 μM, 83.4 ± 19 μM, and 766 ± 302 μM, respectively) were in the same range, though the K_Ds were somewhat lower, than those in previous reports on murine synaptotagmin using NMR (Shao et al., 1996; Ubach et al., 1998). As predicted, Fig. 2A demonstrates that the C₂A^D2E mutation inhibited Ca²⁺ binding by the C₂A domain. We assessed protein folding by circular dichroism (CD) spectroscopy to exclude misfolding of the mutant C₂A domain. As shown in Fig. 2B, the C₂A^D2E mutation does not alter the CD spectrum as compared to wild-type C₂A. Thus, our novel C₂A mutation is correctly folded and largely inhibits Ca²⁺ binding by the C₂A domain. D2 only directly coordinates the first Ca²⁺ site (Fig. 1B: D2---Ca1). If the C₂A^D2E mutation does in fact remove all Ca²⁺ binding, this would support a model in which Ca²⁺ binding to site 1 is requisite for Ca²⁺ binding to sites 2 and 3.

To determine the importance of electrostatic repulsion by the C₂A domain for synchronous synaptic transmission at an intact synapse, we examined evoked transmitter release at Drosophila neuromuscular junctions expressing our mutant transgenic synaptotagmin protein (syt^A-D2E) in the absence of any wild-type synaptotagmin 1. To indicate its transgenic origin, we will refer to the mutant as P[syt^A-D2E] and the transgenic controls as P[syt^WT]. We found that the syt^A-D2E Ca²⁺-binding motif mutation, which maintains the negative charge of the C₂A pocket, decreased synchronous evoked release by >80% (Fig. 3A,B). The amplitude of the excitatory junction potential (EJP) in P[syt^A-D2E] was 5.16 ± 0.96 mV (line 3, n=11) or 2.84 ± 0.41 mV (line 2, n=8) compared with 34.19 ± 3.97 (n=8) in P[syt^WT] controls [Fig. 3A (line 3 shown) and Fig. 3B, p≪0.001]. There was no significant difference between the independent mutant lines (p>0.07) demonstrating that the decrease in evoked release is not due to the insertion sites of the transgene, but rather is a direct result of the mutation.

Since analogous aspartate to asparagine mutations in C₂A, which also inhibit Ca²⁺ binding but partially neutralize the negative charge of the pocket, did not significantly impair synchronous evoked release at this same synapse (Robinson et al., 2002; Yoshihara et al., 2010), or at cultured excitatory synapses (Fernandez-Chacon et al., 2002; Stevens and Sullivan, 2003), our findings demonstrate that the key function of Ca²⁺ binding to the C₂A domain is to neutralize the negative charge of the C₂A Ca²⁺-binding pocket. Thus, Ca²⁺ binding to the C₂B domain is necessary and sufficient for synchronizing synaptic vesicle fusion to Ca²⁺ influx (Mackler et al., 2002; Robinson et al., 2002), as seen by the low level of fast, synchronous, evoked release that remains in our P[syt^A-D2E] mutants (Fig. 3A,B). But it is not sufficient to efficiently trigger the electrostatic switch. Efficient, synchronous release requires Ca²⁺ binding to the C₂A and C₂B domains to neutralize both negatively-charged Ca²⁺-binding pockets and flip the electrostatic switch resulting in fast, synchronous vesicle fusion.

The C₂A Ca²⁺-binding motif mutation had no significant effect on either the amplitude or frequency of spontaneous transmitter release. The amplitude of miniature excitatory junction potentials (mEJPs) in P[syt^A-D2E] was 0.69 ± 0.03 mV (line 3, n=12) or 0.68 ± 0.03 mV (line 2, n=12) compared to 0.70 ± 0.02 mV (n=12) in P[syt^WT] (Fig. 3C, p>0.7). The constant mEJP amplitude demonstrates that synaptic vesicle filling and the postsynaptic response to neurotransmitter are unimpaired. The frequency of mEJPs in the C₂A mutants was also not significantly different at 3.71 ± 0.52 Hz (line 3), or 4.45 ± 0.37 Hz (line 2) in P[syt^A-D2E] compared to 3.19 ± 0.48 Hz in P[syt^WT] controls (Fig. 3D, p>0.15). Since this C₂A Ca²⁺-binding motif mutation results in a large decrease in evoked release (Fig. 3A,B), yet does not affect the rate of spontaneous release (Fig. 3A,D), the increase in spontaneous release seen in other synaptotagmin point mutants (Mackler and Reist, 2001; Mackler et al., 2002; Paddock et al., 2008) cannot be explained as an indirect developmental artifact resulting from the decrease in evoked release. Rather, our findings support the hypothesis that synaptotagmin plays a direct role in regulating the rate of spontaneous release (Broadie et al., 1994; Morimoto et al., 1995; Mace et al., 2009) and that the negative charge of the C₂A Ca²⁺-binding pocket plays a key role in this process. Indeed, when Ca²⁺ binding is inhibited by D→N mutations of the C₂A Ca²⁺-binding pocket, the rate of spontaneous release is increased 6-fold (Yoshihara et al., 2010). This dramatic difference in the effect on spontaneous fusion frequency between D→N mutations and our D→E mutation demonstrates that the electrostatic repulsion created by the negative charge in the C₂A Ca²⁺-binding pocket must be neutralized to enhance any fusion.

To assess whether the decrease in evoked release resulted from changes in either the apparent Ca²⁺ affinity or cooperativity of release, we measured EJP amplitudes in extracellular Ca²⁺ concentrations ranging from 0.6 mM to 5 mM. P[syt^A-D2E] had a significantly reduced evoked response compared to the control at every extracellular Ca²⁺ concentration (Fig. 3E). To facilitate comparison, the data were plotted as a percentage of the maximal response (Fig. 3F). The apparent Ca²⁺ affinity of release in vivo was decreased in the P[syt^A-D2E] mutants; 45% more Ca²⁺ was required to trigger a half maximal response (EC₅₀ = 1.92 ± 0.26 mM) compared to P[syt^WT] controls (EC₅₀ = 1.33 ± 0.18 mM). By plotting the mean EJP amplitude against the extracellular Ca²⁺ concentration at non-saturating levels on a double log plot (Fig. 3G), we found that the Ca²⁺ cooperativity of release was not affected by the syt^A-D2E mutation [n = 2.52 for P[syt^A-D2E] and n = 2.69 for P[syt^WT], similar to previously reported values at wild type neuromuscular junctions in Drosophila (Stewart et al., 2000; Okamoto et al., 2005). The shift in the Ca²⁺ affinity of release with no effect on the cooperativity of release is consistent with the stochastic model of cooperativity (Dodge and Rahamimoff, 1967; Stewart et al., 2000; Fernandez-Chacon et al., 2001; Mackler et al., 2002). However some synaptotagmin mutations alter the cooperativity of release which would favor the stoicheiometric model (Dodge and Rahamimoff, 1967; Yoshihara and Littleton, 2002; Tamura et al., 2007). Further work will be necessary to discriminate between these models. Regardless, the decrease in the apparent Ca²⁺ affinity of evoked release is consistent with the finding that the syt^A-D2E mutation specifically inhibits Ca²⁺ binding by the C₂A domain and demonstrates that Ca²⁺ binding by C₂A is essential for fast, synchronous synaptic transmission.

Since a decrease in synchronous evoked release could also result from protein misexpression, we assessed transgenic synaptotagmin expression levels in each transgenic line. Western blot analysis of third instar larval central nervous systems with an anti-synaptotagmin antibody demonstrated that the C₂A Ca²⁺-binding motif mutant lines expressed similar levels of transgenic synaptotagmin as the transgenic control (Fig. 4A,B). In addition, the mutant synaptotagmin was appropriately localized to the neuromuscular junction (Fig. 4C). Therefore, the deficits in evoked release are not due to insufficient protein expression or protein mislocalization.