The Ca2+ sensor synaptotagmin-1 (syt-1) regulates neurotransmitter release by interacting with anionic phospholipids. Here we test the idea that the intrinsic kinetics of syt–membrane interactions determine, in part, the time course of synaptic transmission. To tune the kinetics of this interaction, we grafted structural elements from the slowest isoform, syt-7, onto the fastest isoform, syt-1, resulting in a chimera with intermediate kinetic properties. Moreover, the chimera coupled a physiologically irrelevant metal, Sr2+, to membrane fusion in vitro. When substituted for syt-1 in mouse hippocampal neurons, the chimera slowed the kinetics of synaptic transmission. Neurons expressing the chimera also evinced rapid and efficient Sr2+ triggered release, in contrast to the weak response of neurons expressing syt-1. These findings reveal presynaptic sensor–membrane interactions as a major factor regulating the speed of the release machinery. Finally, the chimera failed to clamp the elevated spontaneous fusion rate exhibited by syt-1 KO neurons, indicating that the metal binding loops of syt-1 regulate the two modes of release by distinct mechanisms.
SIGNIFICANCE STATEMENT In calcium, synaptotagmin-1 triggers neurotransmitter release by interacting with membranes. Here, we demonstrate that intrinsic properties of this interaction control the time course of synaptic transmission. We engineered a “chimera” using synaptotagmin-1 and elements of a slower isoform, synaptotagmin-7. When expressed in neurons, the chimera slowed the rate of neurotransmitter release. Furthermore, unlike native synaptotagmin-1, the chimera was able to function robustly in the presence of strontium–a metal not present in cells. We exploited this ability to show that a key function of synaptotagmin-1 is to penetrate cell membranes. This work sheds light on fundamental mechanisms of neurotransmitter release.
The fusion of synaptic vesicles (SVs) with the presynaptic plasma membrane is catalyzed by SNARE proteins: vesicular (v-) and target membrane (t-) SNAREs assemble into four-helix bundles that form the core of a conserved membrane fusion machine. A SV protein, p65/synaptotagmin-1 (syt-1; Matthew et al., 1981; Perin et al., 1990), binds Ca2+ via tandem C2 domains (Brose et al., 1992) and has been identified as a Ca2+ sensor that triggers rapid SV exocytosis, as deletion of syt-1 results in a loss of synchronous synaptic transmission (DiAntonio and Schwarz, 1994; Geppert et al., 1994; Littleton et al., 1994; Mackler et al., 2002; Nishiki and Augustine, 2004), and Ca2+ · syt-1 accelerates SNARE-catalyzed fusion in vitro (Fig. 1; Tucker et al., 2004). There are 17 known isoforms of syt (Dean et al., 2012), and the isoforms that trigger fusion (Bhalla et al., 2008) exhibit a wide range of Ca2+ binding affinities (Bhalla et al., 2005) and membrane binding kinetics (Hui et al., 2005). Whether these differences in intrinsic kinetics have functional consequences is not known.
The nonphysiological divalent cation Sr2+ has been used to probe the kinetics of neurotransmitter release, because replacement of extracellular Ca2+ with equimolar Sr2+ yields substantially reduced EPSCs with a largely asynchronous time course (Goda and Stevens, 1994; Rumpel and Behrends, 1999; Xu-Friedman and Regehr, 2000; Lau and Bi, 2005). Sr2+ readily enters presynaptic boutons through voltage-gated channels, but endogenous Ca2+ binding proteins are largely ineffective at buffering Sr2+, leading to a Sr2+ transient that can persist for seconds (Rumpel and Behrends, 1999; Xu-Friedman and Regehr, 2000; Babai et al., 2014).
The goal of this study was twofold: first, to determine whether the intrinsic kinetics of syt determine the rate of neurotransmitter release, and second, to use a novel chemical genetic approach to engineer a sensor that responds to Sr2+. Therefore, a chimera between the fastest syt (syt-1) and a slower isoform, which responds to Sr2+, was generated. This chimera exhibited slower intrinsic membrane binding kinetics and slowed the rate of neurotransmitter release in neurons, arguing that EPSC decay kinetics are partially governed by the intrinsic kinetics of the presynaptic Ca2+ sensor. Furthermore, our modifications conferred upon syt-1 the ability to both penetrate membranes in vitro and trigger synchronous release in vivo in response to Sr2+, consistent with the notion that penetration of the target membrane by syt-1 is a key step in SV exocytosis. Additionally, we dissociate evoked exocytosis from spontaneous single vesicle fusion, identifying a role for the metal binding loops of syt-1 in suppressing “miniature” release.
Materials and Methods
The following lipids were purchased from Avanti Polar Lipids: 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine [phosphatidylethanolamine (PE)]; 1,2-dioleoyl-sn-glycero-3-phospho-l-serine [phosphatidylserine (PS)]; 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine [phosphatidylcholine (PC)]; 1,2-dipalmitoyl-sn-glycero-3-phospho-ethanolamine-N-(7-nitro-2–1,3-benzoxadiazol-4-yl) (NBD-PE);N-(lissamine rhodamine B sulfonyl)-1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (rhodamine-PE); and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(5-dimethylamino-1-naphthalenesulfonyl) (dansyl-PE). N,N′-dimethyl-N-(iodoacetyl)-N′-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)ethylenediamine (IANBD-amide) was purchased from Invitrogen. Accudenz was purchased from Accurate Chemical & Scientific Corporation. Glutathione Sepharose and Ni2+ Sepharose High Performance affinity media were obtained from GE Healthcare Bio-Sciences.
Recombinant proteins and protein purification.
cDNA encoding the cytoplasmic domain (denoted C2AB) of rat syt-1 (residues 96–421; the amino acid residues for all other syt constructs below are indicated in parentheses) was provided by T. C. Südhof (Department of Molecular and Cellular Physiology, Howard Hughes Medical Institute, Stanford University, Stanford, CA); the D374 mutation was corrected by replacement with a glycine. cDNA for syt-2 (C2AB, 139–423), syt-6 (C2AB, 143–426), syt-7 (C2AB, 134–403), and syt-9 (C2AB, 104–386) were provided by M. Fukuda (Department of Developmental Biology and Neurosciences, Tohoku University, Sendai, Miyagi, Japan). cDNA encoding syt-3 (C2AB, 290–569) was provided by S. Seino (Department of Physiology and Cell Biology, Kobe University Graduate School of Medicine, Kobe, Japan). W. A. Catterall (Department of Pharmacology, University of Washington, Seattle, WA) provided α1B-synprint in a pTrcHis vector.
Syt-1 C2AB was subcloned into pGEX vectors to generate GST-tagged fusion proteins. Syt-2, syt-6, syt-7 and syt-9 C2AB were subcloned into a pTrcHis vector, and syt-3 C2AB was subcloned into a pET28a vector, to generate His6-tagged fusion proteins.
Chimeras and cysteine mutant forms of syt-1 and chimera1,3 were generated using PCR and a QuikChange Site-Directed Mutagenesis Kit (Stratagene). Chimeras were generated by grafting the metal binding loops of syt-7 C2A (loop 1, 162–172; loop 2, 192–198; loop 3, 225–233) onto syt-1 C2B. For cysteine mutants, first the lone native cysteine, C277, was mutated to an alanine. Then, a single cysteine was introduced at M173 and F234 of C2A and V304 and I367 of C2B for syt-1 and M173 and F234 of C2A and F304 and F367 of C2B for chimera1,3.
cDNA for SNAP-25B was provided by M. C. Wilson (Department of Neurosciences, University of New Mexico School of Medicine, Albuquerque, NM). Full-length synaptobrevin-2 (syb) and syntaxin-1A (syx) were provided by J. E. Rothman (Department of Cell Biology, School of Medicine, Yale University, New Haven, CT). All individual SNARE proteins were subcloned into pTrcHis vectors to generate His6 fusion proteins. Full-length SNAP-25B and syx were subcloned into a pRSFDuet vector to generate His6-tagged t-SNARE heterodimers (Chicka et al., 2008). All GST and His6-tagged fusion proteins were expressed in Escherichia coli and purified as described previously (Liu et al., 2014).
Full-length syt-1 and chimera1,3 were subcloned into pLox Syn-DsRed-Syn-GFP lentiviral vector (provided by F. Gomez-Scholl, Facultad de Medicina, Departamento de Fisiología Médica y Biofísica, Universidad de Sevilla, Sevilla, Spain) for electrophysiological recordings. Proteins were inserted with BamH1-Not1 to replace the existing DsRed, and GFP was left intact to visualize transfected cells.
Protein-free liposomes were prepared as described previously (Liu et al., 2014). Lipid compositions were 15% PS, 30% PE, and 55% PC for penetration assays; 25% PS, 30% PE, and 55% PC for cosedimentation assays; and 25% PS, 25% PE, 5% dansyl-PE, and 45% PC for stopped-flow rapid-mixing assays.
SNARE-bearing vesicles were prepared as described previously (Chicka et al., 2008). Lipid compositions for the vesicles used in the in vitro fusion assays were as follows: 15% PS, 27% PE, 55% PC, 1.5% NBD-PE, and 1.5% rhodamine-PE for v-SNARE vesicles; 15% PS, 30% PE, and 55% PC for heterodimer t-SNARE vesicles; and 25% PS, 30% PE, and 45% PC for syx-only t-SNARE vesicles. For coflotation assays, t-SNARE heterodimer vesicles were made from a lipid composition of 30% PE and 70% PC.
In vitro fusion assays.
Fusion, between v-SNARE vesicles and heterodimer, or syx-only t-SNARE, vesicles was monitored using a Synergy HT multidetection microplate reader (Bio-Tek) as described previously (Chicka et al., 2008; Liu et al., 2014). For standard assays, 1 μm cytoplasmic domain (C2AB) of the indicated syt isoform or chimera was also used, and for split t-SNARE assays, 7 μm soluble SNAP-25B was also used. During each run, 1 mm or the indicated final free [cation] was added and the reaction was monitored for an additional 60 or 120 min. Traces were normalized to the first time point and the maximum fluorescence signal, determined from the addition of n-dodecyl-β-d-maltoside, to determine the %Fmax.
The cysteine mutant forms of syt-1 or chimera1,3 were labeled with IANBD-amide as described previously (Hui et al., 2011). NBD-labeled proteins (0.3 μm) were combined with liposomes (0.1 mm total lipid) in 50 mm HEPES-NaOH, 100 mm NaCl, pH 7.4, buffer plus 0.2 mm EGTA or 1 mm free cation. Fluorescence measurements were performed using a QM-1 fluorometer (Photon Technology International). NBD was excited at 470 nm, and emission spectra were acquired from 490 to 630 nm. Spectra were corrected by subtracting the background; all spectra were normalized to the signal in EGTA.
The ability of syt-1 or chimera1,3 to bind to t-SNAREs was monitored via a coflotation assay using a Optima L-90K ultracentrifuge (Beckman Coulter), as described previously (Tucker et al., 2004). The cytoplasmic domains of syt-1 or chimera1,3 (10 μm) were incubated with PS-free t-SNARE heterodimer vesicles in the presence of Ca2+ or Sr2+ (1 mm), or EGTA (0.2 mm). After 60 min at room temperature, samples were centrifuged, collected, and subjected to SDS-PAGE; gels were stained with Coomassie blue. Data were quantified using densitometry and normalized to the syx band in each lane.
The membrane-binding activity of the syt-1 or chimera1,3 was monitored via a cosedimentation assay using a Optima MAX-E tabletop ultracentrifuge (Beckman Coulter) as described previously (Liu et al., 2014). Data were analyzed using Prism software (GraphPad) to determine the Hill coefficients and [cation]1/2 values.
Stopped-flow rapid mixing.
Divalent cation regulated interactions between syt constructs and membranes were monitored via FRET using an SX.18MV stopped-flow spectrometer (Applied Photophysics) as described previously (Hui et al., 2005). Complexes were formed by mixing 44 nm liposomes, 4 μm C2AB, and 0.2 mm cation, and were subsequently disassembled by rapid mixing with 2 mm EGTA. Traces were fit using the Applied Photophysics Pro Data SX software package. A single exponential function was used to determine the rate of disassembly, kdiss.
Synprint binding assay.
Purified synprint (3 μg) was incubated with 30 μg syt-1 or chimera1,3 GST-fusion proteins, immobilized on 30 μl of glutathione Sepharose beads, as described previously (Chapman et al., 1998) and immunoblotted with anti-T7 tag mouse monoclonal antibody (1:10,000; Novagen) and a goat-anti-mouse HRP secondary antibody (1:10,000; Abcam).
Cell culture and immunostaining.
All procedures were performed under the guidance of the Animal Care and Use Committee (Protocol M01221-0-06-14) at the University of Wisconsin–Madison. Hippocampal neurons from syt-1 KO mice, of either sex, were obtained at postnatal day 0 as described previously (Liu et al., 2014). The cells were infected at 5 d in vitro (DIV) with lentivirus, prepared in HEK293T cells as described previously (Liu et al., 2014), to express syt-1 or chimera1,3, and GFP.
Immunocytochemistry was performed at 14 DIV. Cells were stained with anti-syt-1 (1:500; Developmental Studies Hybridoma Bank), anti-synaptophysin (1:1000; Synaptic Systems), and anti-Map2 (1:500; Millipore) antibodies, DAPI, and Alexa Fluor secondary antibodies (1:500; Life Technologies). Cells were mounted in Fluoromount (Southern Biotechnology Associates) and imaged on an FV1000 upright confocal microscope with a 60×, 1.4 numerical aperture oil-immersion objective (Olympus).
At 14–18 DIV, cells were moved to a recording chamber perfused with a bath solution containing the following (in mm): 128 NaCl, 5 KCl, 25 HEPES, 30 d-glucose, 1 MgCl2, and 5 Ca2+ or Sr2+, pH 7.4 (adjusted to 300–310 mOsm with d-glucose). To ensure that residual Ca2+ did not confound the Sr2+ recordings, Milli-Q water was treated with Chelex-100 resin (Bio-Rad) before use in bath solution to remove residual divalent cations. AMPA receptor currents were isolated with 20 μm bicuculline (Tocris Bioscience) and 50 μm d-APV (Tocris Bioscience). Borosilicate glass pipettes were pulled to a resistance of 3–5 MΩ and filled with an intracellular solution containing 130 mm K-gluconate, 10 mm HEPES, 5 mm Na-phosphocreatine, 2 mm Mg-ATP, 1 mm EGTA, 5 μm QX-314 (Tocris Bioscience), and 0.3 mm Na-GTP, pH 7.4. All experiments were conducted at room temperature.
Whole-cell patch-clamp recordings were made with an Axon MultiClamp 700b amplifier (Molecular Devices) in voltage-clamp mode, sampled at 10 kHz and filtered at 2 kHz. Typical series resistances were 15 MΩ, with 70% of this resistance compensated, and neurons were clamped at −70 mV. Neurons showing significant changes in series resistance over the course of recording were excluded from analysis, as were cells that exhibited a large degree of recurrent excitation that precluded analysis of EPSC kinetics. For measurement of evoked EPSCs, neurons near (within several hundred microns of) the patched (i.e., postsynaptic) cell were stimulated with a bipolar electrode in theta glass tubing. This electrode was placed directly adjacent to the soma of the presynaptic neuron. In this configuration, EPSCs from individual presynaptic cells are isolated (i.e., moving the stimulating electrode >10 microns away from the soma of the presynaptic neuron abolished the response). Neurons whose somata made contact with the somata of other neurons were not stimulated, to avoid evoking overlapping EPSCs from multiple presynaptic neurons. Occasional indirect or polysynaptic responses were easily identified by the lack of smooth rise and decay kinetics and were excluded from analysis. Presynaptic neurons were stimulated with 20 V square-wave pulses every 30 s, and the first artifact-free EPSC (typically the first) was used for analysis. For measurement of paired-pulse facilitation, two EPSCs were evoked (with a 50 ms interstimulus interval) and quantified as the ratio of the second response over the first. Data were analyzed offline in Clampfit (Molecular Devices) and MATLAB (MathWorks) software. Rise and decay times were computed as 20–80% of maximum, and cumulative charge transfer functions (computed across 500 ms) were fit with double exponentials to quantify fast and slow components of the charge transfer (see Fig. 5C).
For recordings of mEPSCs, 1 μm TTX was added to the bath solution immediately before recording, and no series resistance compensation was used. Sixty seconds of data were analyzed for each cell. mEPSCs were quantified with the Clampfit template matching algorithm and custom-written MATLAB software with a 5 pA threshold.
Hypertonic sucrose was applied to measure the RRP of SVs (Rosenmund and Stevens, 1996; Liu et al., 2014). After patching a neuron, a second pipette containing extracellular solution, with 500 mm sucrose added, was positioned at the edge of the of field of view under a 40× objective (upstream of solution flow). The sucrose solution was puffed onto patched cells with a Picospritzer III (Parker Hannifin) such that all boutons contacting a given neuron were stimulated. Sucrose was applied for 10 s, yielding a response with distinct fast and slow (steady-state) phases (see Fig. 7A). The fast component of the response was integrated to measure the RRP.
All recordings were made from a minimum of three coverslips each from three independent litters of animals. The number of cells, N, is indicated in the figure legends (N = 10–20 per condition for each electrophysiology experiment). Statistical significance was assessed with Student's t tests or Mann–Whitney tests as appropriate.
We first identified syt isoforms that couple Sr2+ to fusion, using a well-characterized “standard” in vitro membrane fusion assay that utilizes preassembled t-SNARE heterodimers (Fig. 1A; Tucker et al., 2004). Six isoforms were screened: syt-6, syt-7, and syt-9 coupled both Ca2+ and Sr2+ to fusion; syt-1, syt-2, and syt-3 were efficiently activated by Ca2+ but not Sr2+ (Fig. 1B–G; Bhalla et al., 2005). Because syt-7 has the slowest membrane disassembly kinetics within this family (Hui et al., 2005) and is absent from SVs (Dean et al., 2012), we performed further analysis of this isoform. Syt-7 was rescreened using a more stringent variant of the fusion assay in which syt must first fold soluble SNAP-25B onto syx to trigger fusion (Fig. 1H; Bhalla et al., 2006). In this “split” t-SNARE fusion assay, Sr2+ failed to activate syt-1 (Fig. 1I), but both metals activated syt-7 (Fig. 1J). Isothermal titration calorimetry (ITC) confirmed that syt-1 binds one Sr2+ ion via its C2B domain (data not shown; Cheng et al., 2004). These findings might explain why Sr2+ fails to efficiently activate syt-1, as mutagenesis studies indicate the C2B domain must bind two metal (Ca2+) ions to trigger exocytosis (Nishiki and Augustine, 2004). Ca2+-ligand mutations in C2B completely disrupt function, whereas analogous mutations in C2A have little effect (Chapman, 2008); hence, our efforts to engineer syt-1 focused on the C2B domain.
In an effort to tune the kinetics of syt-1, we replaced its metal binding loops with those of syt-7 (Fig. 2A,B). To determine which C2 domain of syt-7 coupled Sr2+ to fusion, both C2A and C2B were screened in a standard fusion assay. Since both C2 domains of syt-7 stimulated fusion in response to Sr2+ (data not shown), chimeras were constructed by grafting, individually and in combinations, the metal binding loops of either syt-7 C2A or C2B onto syt-1 C2B (Fig. 2A,B). Of these two sets of chimeras, only the proteins that harbored loops from syt-7 C2A, but not C2B (data not shown), responded to Sr2+ in the standard fusion assay (Fig. 2C,D). These four constructs, which all contained loop 3, were therefore rescreened in the split t-SNARE fusion assay and only one, chimera1,3, efficiently coupled Sr2+ to fusion (Fig. 2E,F). The cation (Ca2+ or Sr2+) sensitivities were greater for chimera1,3 than for syt-1, in terms of both driving fusion (Fig. 2G,H; Table 1) and binding to membranes (Fig. 3A; Table 2), consistent with the higher affinity of syt-7 for metals as compared to syt-1 (Bhalla et al., 2005). Syt-1 also regulates fusion in vitro, by interacting with t-SNAREs (Bhalla et al., 2006), so we directly measured binding using PS-free heterodimer t-SNARE vesicles. In the presence of Ca2+, chimera1,3 bound more efficiently than syt-1; in Sr2+, no differences were observed (Fig. 3B). It should be noted, however, that the relevance of this interaction during secretion from cells remains the subject of debate (Zhang et al., 2002, 2010).
Upon binding Ca2+, metal binding loops 1 and 3, from each C2 domain of syt-1, penetrate membranes, with a small contribution from loop 2 (Hui et al., 2011). To monitor penetration, loops 1 and 3, were individually labeled with an environmentally sensitive fluorophore, NBD. In the presence of Ca2+, all labeled loops of syt-1 penetrated membranes. However, Sr2+ failed to trigger efficient penetration (Fig. 3C,D). Importantly, both C2 domains of chimera1,3 efficiently penetrated membranes in the presence of either Ca2+ or Sr2+ (Fig. 3C,D). These results support the emerging view that membrane penetration by syt-1 is a crucial step in Ca2+-triggered membrane fusion (Paddock et al., 2011; Liu et al., 2014) and provide an explanation as to why Sr2+ drives synaptic transmission less efficiently than Ca2+. Interestingly, although the C2A domain of chimera1,3 was not engineered to bind Sr2+, it efficiently penetrated membranes (Fig. 3C,D); ITC was not possible with this construct. So, either an active C2B domain “pulls” C2A into the bilayer, or C2A is activated via contact with the engineered C2B domain (Bai et al., 2002; Liu et al., 2014).
To determine whether these alterations in syt-1 led to changes in membrane interactions, we performed stopped-flow rapid-mixing experiments. To mimic the decay of Ca2+ transients in nerve terminals, sensor–cation–membrane complexes were rapidly mixed with excess chelator. In Ca2+, chimera1,3 disassembled much more slowly than syt-1 (Fig. 3E, left), a point we return to below. Similarly, the disassembly rates of chimera1,3 were slower in Sr2+ than syt-1 (Fig. 3E, right).
To determine whether chimera1,3 can change the rate of neurotransmitter release, we expressed it, or syt-1, in dissociated hippocampal neurons derived from syt-1 KO mice. Both proteins were properly localized to nerve terminals, as colocalization (Pearson's correlation coefficient) with synaptophysin did not differ (p > 0.1; Fig. 4). The number of synapses (quantified as the number of synaptophysin puncta) also did not differ across conditions (p > 0.1; data not shown).
We performed patch clamp recordings of EPSCs in the presence of 5 mm Ca2+ or Sr2+. Individual EPSCs were evoked with a bipolar electrode positioned near the soma of a nearby cell. Approximately 60% of stimulated neurons had monosynaptic connections with the patched (postsynaptic) neuron, and this did not differ across conditions (p > 0.1; data not shown). The characteristics of these EPSCs were comparable to what has been found previously in this preparation (Liu et al., 2009, 2014; Yao et al., 2011).
Chimera1,3 did not simply rescue rapid evoked release, but exhibited a gain-of-function relative to syt-1. In the presence of Ca2+, chimera1,3 increased the total charge transfer without altering the peak current (Fig. 5A; chimera1,3 charge, 14.66 ± 1.94 pC; syt-1 charge, 7.12 ± 1.10 pC; p < 0.01; N = 10–20 per condition). Importantly, in Ca2+, chimera1,3 yielded a slower rise of evoked release (Fig. 5B; chimera1,3 rise slope, 60.48 ± 16.8 pA/ms; syt-1 rise slope, 132.9 ± 26.7 pA/ms; p < 0.05) and a decay time twice as long as that of neurons expressing syt-1 (Fig. 5B; chimera1,3 decay time, 32.90 ± 2.12 ms; syt-1 decay time, 14.45 ± 1.49 ms; p < 0.01; N = 10–20 per condition). When cumulative charge transfers were fit with double exponential functions, we found that EPSC decays of neurons expressing chimera1,3 had a significantly larger slow component of release (Fig. 5C; chimera1,3 slow charge amplitude, 4.45 ± 0.79 pC; syt-1 slow charge amplitude, 2.09 ± 0.74 pC; p < 0.05), and the time constant (τ) of the fast component was significantly lengthened (Fig. 5C; chimera1,3 fast charge τ, 20.1 ± 1.49 ms; syt-1 fast charge τ, 12.8 ± 1.45 ms; p < 0.01). Hence, grafting the loops of the slowest syt isoform, syt-7 (Hui et al., 2005), onto syt-1 slows the kinetics of synaptic transmission.
A key finding was that in Sr2+, chimera1,3 significantly increased both the total charge transfer and peak current relative to syt-1 (Fig. 6A; chimera1,3 charge, 9.21 ± 1.37 pC; syt-1 charge, 5.94 ± 1.06 pC; p < 0.05; chimera1,3 amplitude, 262.5 ± 38.5 pA; syt-1 amplitude, 141.0 ± 21.7 pA; p < 0.05; N = 10–20 per condition). In neurons expressing chimera1,3, the rising phase of the EPSC in Sr2+ was steeper than in cells expressing syt-1 (Fig. 6B; chimera1,3 rise slope, 66.17 ± 13.3 pA/ms; syt-1 rise slope, 35.49 ± 7.76 pA/ms; p < 0.05). There were no observable differences in the decay phase of EPSCs regulated by chimera1,3 and syt-1 in Sr2+ (Fig. 6B), presumably because of the slow rate (seconds) at which Sr2+ is cleared from nerve terminals (Xu-Friedman and Regehr, 2000; Babai et al., 2014). As such, EPSC decays in Sr2+ were not analyzed further. Consistent with previous reports, Sr2+ was ineffective at triggering release in syt-1 KO neurons (Fig. 6A; Shin et al., 2003; Babai et al., 2014), indicating that any remaining Ca2+ sensors do not efficiently sense Sr2+.
It should be noted that the increase in charge transfer could in principal be due to an increase in the size of the readily-releasable pool (RRP) of vesicles (Rosenmund and Stevens, 1996), rather than an effect on the release machinery during the final stages of exocytosis. To address this, we measured the RRP in KO, syt-1, and chimera1,3-expressing neurons with hypertonic sucrose. We found that chimera1,3 rescued the size of the RRP as efficiently as syt-1 in either Ca2+ (Fig. 7A, left panels) or Sr2+ (right panels). To the best of our knowledge, these experiments provide the first example of a mutant form of syt-1 that alters the kinetics of functional synaptic transmission (i.e., without impairing the RRP).
Because overexpression of wild-type and mutant forms of syt do not alter Ca2+ entry (Wang et al., 2001; Young and Neher, 2009), it is unlikely that our kinetic effects are due to changes in metal influx. Nonetheless, we examined paired-pulse facilitation, a form of short-term plasticity sensitive to changes in presynaptic Ca2+ dynamics (Fioravante and Regehr, 2011). The paired-pulse ratio was similar between neurons expressing syt-1 and chimera1,3 (Fig. 7B). Moreover, the ability of chimera1,3 to bind synprint, the Ca2+ channel domain that mediates interactions with the release machinery (Sheng et al., 1997), was indistinguishable from that of syt-1 (Fig. 7C). Collectively, these results argue against the kinetic effects in Figure 6 being due to an alteration in the coupling of release machinery to Ca2+ channels.
Finally, recordings of “mini” excitatory currents (mEPSCs), reflecting spontaneous release of individual SVs, showed no difference in amplitude or kinetics among conditions (Fig. 8), arguing against a postsynaptic effect. Loss of syt-1 results in an increase in the frequency of mEPSCs, suggesting that syt-1 has a second function as a fusion clamp (Chicka et al., 2008; Liu et al., 2014). Interestingly, chimera1,3 failed to clamp minis, showing a frequency equivalent to KO (Fig. 8B). Thus, metal binding loops 1 and 3 of syt-1 C2B not only govern the kinetics of evoked release, but also play roles in suppressing spontaneous transmission.
Upon binding Ca2+, syt-1 penetrates membranes that harbor anionic phospholipids (Chapman and Davis, 1998; Chapman, 2008). In a reconstituted system, syt-1 preferentially penetrated t-SNARE-bearing membranes to accelerate fusion (Bai et al., 2004; Chicka et al., 2008). Moreover, the abilities of mutant forms of syt-1 to penetrate membranes was correlated with their abilities to drive synchronous synaptic transmission in cultured neurons (Liu et al., 2014). This latter finding suggests that membrane penetration constitutes an essential step in excitation–secretion coupling. The work presented here provides a more direct demonstration that penetration by syt-1 is a key step in SV exocytosis. Specifically, we used a chemical genetic approach to engineer syt-1 to respond to a nonphysiological metal, Sr2+. In the presence of Sr2+, syt-1 binds membranes (Fig. 3A), but fails to penetrate (Fig. 3C,D), and so only weakly triggers exocytosis in neurons (Fig. 6A). In contrast, chimera1,3 bound (Fig. 3A) and penetrated membranes (Fig. 3C,D) in response to Sr2+, resulting in robust synchronous neurotransmitter release (Fig. 6A). These results prompt the question as to whether the kinetics of the crucial interaction between syt-1 and membranes affect the time course of synaptic transmission.
EPSC decays are governed by a myriad of presynaptic and postsynaptic mechanisms, including presynaptic action potential waveform (Taschenberger and von Gersdorff, 2000), AMPA receptor desensitization (Wall et al., 2002), and clearance of glutamate from the synaptic cleft (Takahashi et al., 1995). The effects reported here reveal a novel factor that determines the kinetics of transmission, namely, chimera1,3 slowed the sensor–membrane disassembly rate upon chelation of Ca2+ (Fig. 3E, left) and also slowed the rate at which EPSCs decay (Fig. 5B,C). Once activated by metal, the chimera “holds onto” membranes longer, thus widening the window in which vesicles can fuse. It is unlikely that these kinetic changes are the result of changes in Ca2+ affinity, as it has been shown that simply increasing the affinity of syt-1 for Ca2+ does not alter the time course of transmission (Rhee et al., 2005). Rather, the longer EPSC decays reported here likely reflect longer-lived sensor–membrane complexes that continue to drive release. Thus, we propose that the decay phase of EPSCs are governed, in part, by the dwell time of syt-1–membrane complexes.
Previous work has shown that syt-1 KO neurons exhibit a significantly enhanced rate of spontaneous “miniature” release (DiAntonio and Schwarz, 1994; Littleton et al., 1994; Liu et al., 2009). This led to the suggestion that during basal (i.e., low Ca2+) conditions, syt-1 reduces the rate of spontaneous fusion by serving as a “clamp,” either by inhibiting SNARE assembly directly (Chicka et al., 2008) or by inhibiting a second Ca2+ sensor (Xu et al., 2009; Kochubey and Schneggenburger, 2011). Interestingly, while chimera1,3 was able to efficiently trigger evoked exocytosis, it failed to rescue this clamping function of syt-1. The mechanism of this clamping function is unclear, because it can be disrupted by mutating either the metal/membrane binding loops (the present work) or the linker connecting the two C2 domains of syt-1 (Liu et al., 2014). It should also be noted that spontaneous release appears to involve a somewhat distinct pool of SVs (Kavalali, 2015), so the differential regulation of these two modes of release likely involves numerous other factors.
Finally, during synaptic transmission, chimera1,3 restored the total charge transfer in Sr2+ (Fig. 6A) to levels comparable to syt-1 in Ca2+ (Fig. 5A), demonstrating the functionality of this engineered sensor. Future work will use this chimera as a tool to dissect aspects of syt-1 function from other Ca2+ triggered processes in nerve terminals (e.g., in the context of spontaneous release), and will take advantage of its unique kinetic properties to probe the effect of glutamate release rates on network behavior.
This work was supported by NIH Grant MH061876. C.S.E was supported by a PhRMA Foundation predoctoral fellowship and University of Wisconsin–Madison Molecular and Cellular Pharmacology Training Grant 5T32-GM008688. D.A.R. was supported by a National Science Foundation Graduate Research Fellowship Program Grant DGE-1256259 and Neuroscience Training Program Grant T32-GM007507. E.R.C. is an investigator at the Howard Hughes Medical Institute.
The authors declare no competing financial interests.
- Correspondence should be addressed to Edwin R. Chapman, Howard Hughes Medical Institute, University of Wisconsin, 1111 Highland Avenue, Madison, WI 53705-2275.