Article |
Address correspondence to J. Troy Littleton, The Picower Center for Learning and Memory, Massachusetts Institute of Technology, E18-672, 50 Ames St., Cambridge, MA 02139. Tel.: (617) 452-2605. Fax: (617) 452-2249. email: troy{at}mit.edu
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Abstract |
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Key Words: synaptic transmission; exocytosis; Drosophila; membrane trafficking; C2 domain
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Introduction |
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Apart from Syt 1, more than a dozen additional synaptotagmins have been identified in mammals (Südhof, 2002), whereas the Caenorhabditis elegans and Drosophila melanogaster genomes encode eight and seven synaptotagmin genes, respectively (Lloyd et al., 2000; Adolfsen and Littleton, 2001). Several observations suggest that different synaptotagmin isoforms might cooperate to regulate the same exocytotic process, including dense core vesicle fusion in PC12 cells (Saegusa et al., 2002; Tucker et al., 2003). Heterooligomerization of distinct synaptotagmins has also been hypothesized to regulate the calcium sensitivity of neurotransmitter release (Littleton et al., 1999; Desai et al., 2000; Wang et al., 2001). Alternatively, each synaptotagmin isoform may participate in distinct membrane trafficking pathways. Supporting this model, several synaptotagmin isoforms do not colocalize with Syt 1 (Butz et al., 1999; Martinez et al., 2000; Ibata et al., 2002). To investigate the possibility that other synaptotagmins are involved in regulating neurotransmitter release, we characterized the seven synaptotagmins encoded in the Drosophila genome. We find that synaptotagmin isoforms localize to nonoverlapping subcellular compartments, suggesting that they participate in the regulation of distinct membrane trafficking steps in vivo.
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Results |
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Expression analysis of Drosophila synaptotagmin mRNAs
To characterize the expression profile of the Drosophila synaptotagmin family, we assayed their mRNA abundance and localization. The abundance of mRNA transcripts and their temporal expression in embryos was determined from developmental microarray expression experiments performed by the Berkeley Drosophila Genome Project. Embryonic mRNA was isolated at 1-h windows throughout the first 12 h of development and used to probe Affymetrix Drosophila genome arrays that include all seven Drosophila synaptotagmin isoforms (Fig. 2 A). The onset of expression of syt 1 coincides with formation of the nervous system. Similarly, none of the remaining synaptotagmins show a peak of expression before 11 h AEL, making it unlikely that they function at earlier stages of development. syt 4, syt 7, and syt ß mRNA showed a similar developmental regulation, with increased expression from 10 to 12 h during nervous system development. The mRNAs for syt 12, syt 14, and syt were expressed at very low levels throughout embryonic development.
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To identify the expression patterns for the synaptotagmin family, we performed RNA in situ hybridization experiments on 022-h embryos using RNA probes specific to each isoform (Fig. 2, CH). Similar to syt 1, syt 4 and syt 7 mRNAs were abundantly expressed throughout the central nervous system (CNS; Fig. 2, D and E). In addition to CNS staining, syt 7 mRNA was expressed in several tissues outside the nervous system, indicating a more ubiquitous expression pattern. syt 14 was expressed at low levels in the CNS (Fig. 2 F). syt and syt ß displayed a highly restricted expression pattern in subsets of CNS cells. syt ß was expressed in a few bilaterally symmetrical large cells found in each segment of the ventral nerve cord (VNC; Fig. 2 G). Expression was also detected in peritracheal cells and within a small population of neurons in each brain lobe. The syt
isoform was found in a distinct population of smaller cells within each VNC segment, and in a subset of neurons within each brain lobe (Fig. 2 H). syt 12 mRNA was not detected by microarray or in situ analysis, suggesting it is expressed at levels below detection. Together with the microarray analysis, our data indicate that Syt 1 and Syt 4 are expressed in most, if not all, neurons. Syt 7 is also abundantly expressed, but in a ubiquitous pattern both within and outside of the nervous system. The remaining synaptotagmins display restricted expression in the nervous system, labeling only specific subpopulations of cells.
Generation of antisynaptotagmin antisera and characterization of compartmental localization
We had previously generated antisera to Drosophila Syt 4 and found the antisera recognized an antigen copurifying with synaptic vesicles and coIPing with Syt 1, leading us to conclude that Syt 4 was a synaptic vesicle protein (Littleton et al., 1999). The lack of a mutant in syt 4 prevented us from confirming the antibody was isoform specific. It is now clear that the original Syt 4 antisera is not isoform specific, as the antigen detected by the antisera is not removed in animals lacking the syt 4 locus, resulting in cross reactivity with the synaptic vesicle-localized Syt 1 protein. Therefore, to define the subcellular localization of the Drosophila synaptotagmins, we generated isoform-specific antisera to each synaptotagmin using multiple host animals and performed control experiments to confirm their specificity. The reactivity of the purified antisera to Drosophila synaptotagmins is shown in Fig. 3 A. The synaptotagmin antisera uniquely recognize their respective recombinant proteins and do not cross-react with other isoforms. In addition, preincubation of the antisera with excess recombinant antigen abolished the signal obtained on Westerns (Fig. 3 E) and immunostaining (Fig. S1, available at http://www.jcb.org/cgi/content/full/jcb.200312054/DC1). Further confirmation that the antiSyt 7 antibody is specific was obtained by generating UAS-syt 7 transgenic animals and overexpressing the protein using elav-GAL4. Overexpression of Syt 7 resulted in up-regulation of the signal detected by antiSyt 7 antisera. The most definitive confirmation of isoform specificity is to demonstrate that immunoreactivity is lost in mutant animals. This has been determined for our antisera to Syt 1 and Syt 4, proving that these antisera display isoform specificity (Fig. 3 D and Fig. 4 B). We have not yet generated mutations in the remaining synaptotagmins, so their specificity is still tentative using these rigorous requirements. However, as shown in Fig. 6 and Fig. 7, the localization patterns for each isoform are unique and correspond with in situ results.
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Subcellular localization of Drosophila synaptotagmins
To characterize the subcellular distribution of the synaptotagmins, we examined their localization in Drosophila embryos and larvae using immunocytochemistry. Syt 1 has been previously localized to synaptic vesicles at presynaptic terminals (Littleton et al., 1993). Similar to Syt 1 and consistent with our in situ localization data, the Syt 4 protein was found concentrated in the neuropil of the larval CNS (Fig. 4 A), suggesting localization to mature synapses. This immunostaining is abolished in mutants that remove the syt 4 locus (Fig. 4 B). During embryonic development, the subcellular localization of Syt 4 is clearly distinct from Syt 1. As shown in Fig. 4 C, Syt 4 is abundant in neuronal cell bodies in the developing CNS at a time in which Syt 1 and other axonal markers such as Fas II have already trafficked to axons, indicating Syt 4 is differentially sorted during the establishment of neuronal polarity. The consequences of this differential sorting are apparent at mature third instar larval neuromuscular junctions (NMJs), where Syt 4, in contrast to Syt 1, is found postsynaptically (Fig. 4, DI). Syt 4 antiserum labels the postsynaptic side of NMJs (Fig. 5, A and B), surrounding the outside of presynaptic terminals (labeled by anti-HRP) in a punctate pattern, suggesting Syt 4 resides in a postsynaptic vesicular compartment. Double labeling experiments performed in animals overexpressing a myc-tagged postsynaptic glutamate receptor subunit, mycGluRIIA, reveals that Syt 4 localizes to regions adjacent to postsynaptic receptor clusters (Fig. 5, C and D). Costaining with Syt 1 antisera demonstrates no overlap in the distribution of the two proteins, confirming Syt 4 is not a synaptic vesicle protein. In addition, overexpression of Syt 4 presynaptically in syt 4 mutant animals does not shift its localization to synaptic vesicles (Fig. 5, E and F). Immunostaining of Syt 4 and Syt 1 revealed a nonoverlapping pattern of expression, with Syt 4 excluded from Syt 1positive synaptic vesicle microdomains (Fig. 5 F). Although there is no overlap between Syt 4 and Syt 1 staining (Fig. 5, E and F), and the majority of Syt 4 labeling is postsynaptic (Fig. 5, A and B), a smaller fraction of Syt 4 may also localize presynaptically. However, synaptic defects present in syt 4 mutants (unpublished data) are rescued by postsynaptic expression, suggesting that Syt 4 functions in postsynaptic trafficking steps required for synaptic growth and plasticity.
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Overexpression of Syt 4 and Syt 7 cannot rescue syt 1 null mutants
The localization of synaptotagmin isoforms to distinct subcellular compartments suggests that they function in unique trafficking pathways. To test this hypothesis, we examined if Syt 4 and Syt 7, which are coexpressed in neurons with Syt 1, could rescue the synaptic transmission and behavioral defects of syt 1 mutants when overexpressed in the nervous system. We overexpressed Syt 4 in the syt 1 null background (sytAD4 in trans to Df(2L)N13) using C155elav-GAL4 (Fig. 8, A and B). To obtain quantitative information on the behavioral rescue, we performed larval locomotion assays to examine the output of the central motor pattern generator. Representative traces of crawling patterns from the control line C155elav-GAL4 and the syt 1 null mutant (sytAD4/Df(2L)N13) are shown in Fig. 8 C. In contrast to the robust locomotion observed in control animals, the lack of Syt 1 dramatically slows larval locomotion. In addition to a decrease in distance traveled and locomotor cycle number (Fig. 8 D), syt 1 null mutants display an increase in the duration of a single locomotor cycle from 1 s to 6 s (Fig. 8 E). Transgenic expression of the syt 1 gene in the null background was able to partially restore all the behavioral defects observed (Fig. 8, CE). In contrast to Syt 1, Syt 4 (Fig. 8, CE) or Syt 7 (not depicted) overexpression did not rescue any aspect of the behavioral defects.
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Discussion |
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Syt 4 was found in the postsynaptic compartment, suggesting it regulates a postsynaptic membrane trafficking pathway. We cannot rule out that a small fraction of Syt 4 may also be present in some presynaptic compartments, though it does not localize to Syt 1positive synaptic vesicles. The detection of the Syt 4 protein by Western analysis and immunocytochemistry with our new antisera is abolished in syt 4 null mutants, confirming the antisera accurately reflects the subcellular localization of Syt 4. These results indicate that previous detection of Syt 4 on synaptic vesicles (Littleton et al., 1999) reflected cross-reactivity of the old antisera with Syt 1. Given that Syt 4 does not colocalize on Syt 1positive synaptic vesicles, the reduction of neurotransmitter release by Syt 4 up-regulation observed in Drosophila (Littleton et al., 1999) is unlikely to be due to heteromultimerization of the two proteins on vesicles and may instead reflect competitive binding to Syt 1 effectors or altered presynaptic calcium buffering.
In terms of Syt 4's postsynaptic localization, there is evidence in several experimental systems for a regulated form of postsynaptic vesicular trafficking (Ludwig et al., 2002). Studies in hippocampal culture neurons indicate that long-term labeling with FM143 loads dendritic organelles that undergo rapid calcium-triggered exocytosis that is blocked by tetanus toxin (Maletic-Savatic and Malinow, 1998). In addition, pharmacological blockage of postsynaptic membrane fusion reduces LTP (Lledo et al., 1998), suggesting postsynaptic vesicle trafficking contributes to synaptic plasticity. Mammalian Syt 4 has been localized within dendrites and soma (Ibata et al., 2002), suggesting Syt 4 and the related homologue Syt 11 may also function postsynaptically. Although the exact role for regulated postsynaptic fusion remains unclear, possibilities include the release of retrograde signals, trafficking of postsynaptic receptors, and/or trafficking of synaptic cell adhesion proteins.
The remaining synaptotagmins were not ubiquitously localized to synapses. Unlike Syt 1 or Syt 4, we could not detect Syt 7 at synapses, but found it was expressed in both neuronal and nonneuronal tissues. Mammalian Syt 7 has been found in secretory lysosomes (Martinez et al., 2000) and in synaptic active zones where it has been postulated to function as a plasma membrane calcium sensor (Sugita et al., 2001). Genetic studies of Syt 7 will be required to determine if it also functions at Drosophila active zones. Peripheral Syt ß staining was restricted to muscle fiber 8 synapses that are known to release the neuropeptide leukokinin (Cantera and Nassel, 1992). In the CNS, Syt ß was observed in a pair of bilateral neurons that may be the DPM neurosecretory neurons known to secrete the amnesiac neuropeptide. The only staining outside the nervous system is detected at tracheal branch points, where a group of myomodulin-releasing neurosecretory cells are located (O'Brien and Taghert, 1998). These localization studies suggest Syt ß is a candidate calcium sensor for mediating dense core vesicle fusion and release of neuropeptides. Similar to Syt ß, Syt showed specific expression in another set of putative CNS neuropeptide-releasing neurons, as well as within the mushroom bodies. In the periphery, staining was restricted to the LBD neurosecretory neuron, which is consistent with a role in neuropeptide release. In addition, the localization of Syt
in mushroom bodies and the possible localization of Syt ß in DPM neurons makes these isoforms attractive candidates for potential roles in vesicular trafficking pathways contributing to neuronal plasticity. We were unable to localize the two remaining synaptotagmins, Syt 12 and Syt 14. It is likely that the proteins are below the detection level of our antisera, which is consistent with the microarray and in situ experiments, indicating that these isoforms are expressed at low levels in embryos and adults. Unlike the other synaptotagmins, these two isoforms lack most of the calcium coordination residues in C2A and C2B in both vertebrates and flies, indicating that they may function in trafficking pathways not regulated by calcium.
In summary, Drosophila synaptotagmin isoforms identify unique membrane-trafficking compartments. A summary of the expression of both the mRNA and protein for each synaptotagmin family member is shown in Fig. 9. Our data indicate that only the Syt 1 isoform is found on synaptic vesicles and so argue against heterooligomerization models. In addition, we find that Syt 4 and Syt 7 cannot rescue the behavioral or physiological defects in syt 1 mutants, suggesting that synaptotagmins define unique membrane trafficking pathways within neurons. It is possible synaptotagmins function in an analogous manner to control vesicle fusion, but do so in distinct compartments. Given that Syt 4 localizes to the postsynaptic compartment, our findings indicate that calcium-dependent membrane trafficking occurs on both sides of the synapse.
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Materials and methods |
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Cluster analysis and dendrogram
Synaptotagmin protein sequences were collected from Drosophila, C. elegans, A. gambiae, F. rubripes, M. musculus, and H. sapiens genomes. Sequences were identified by BLAST analysis of Drosophila synaptotagmin protein sequences against the corresponding genomes deposited in GenBank. Collected sequences were then clustered based on homology using the ClustalW program (http://www.ch.embnet.org/software/ClustalW.html). Results were displayed as a tree diagram using the Phylodendron program (http://iubio.bio.indiana.edu/treeapp/treeprint-form.html).
In situ hybridization
Embryos aged 022 h were collected and processed according to standard procedures. Probes of 500700 bp long were designed to the C2 domain region of each synaptotagmin gene.
Microarray analysis
Microarrays were performed with Affymetrix Drosophila Genechips using biotinylated cRNA using the laboratory methods described in the Affymetrix genechip expression manual (Affymetrix, Inc.). RNA was isolated from heads or heads and bodies of Canton-S males aged 34 d after eclosion at RT. All flies were killed between 12 and 2 p.m. to reduce any circadian-dependent transcriptional changes. Affymetrix high-density oligonucleotide arrays were probed, hybridized, stained, and washed in MIT's Biopolymers Facility according to the manufacturer's instructions. Microarray analysis was performed using Microarray Suite Vs.5 and Data Mining Tool Vs.3 statistics-based analysis software (Affymetrix, Inc.).
Western analysis
Western blots were done using standard laboratory procedures. All synaptotagmin antibodies were used at a 1:1,000 dilution and detected using a goat antirabbit antibody conjugated to HRP (Jackson ImmunoResearch Laboratories). Visualization of HRP was accomplished using a SuperSignal ECL kit (Pierce Chemical Co.).
Gradient centrifugation
Isolation of Canton-S head homogenates was performed as described previously (Littleton et al., 1999). For rate-zonal sedimentation experiments, a post-nuclear extract was layered onto a 1030% sucrose gradient and centrifuged at 50,000 RPM for 1 h in a NVT65 rotor (Beckman Coulter). 1-ml fractions were collected beginning from the bottom of the gradient and proceeding to the top. After collection, fractions were mixed with an equal volume of 2x SDS-PAGE loading buffer and probed by Western analysis. For equilibrium sedimentation experiments, post-nuclear extract was combined with a 26% Optiprep (Axis Shield) solution. The mixed sample was centrifuged at 60,000 RPM for 3.5 h in a NVT65 rotor and fractions were collected as for velocity experiments.
Protein expression and antibody purification
Polyclonal antibodies were generated in rabbits (Invitrogen). For the Syt 4, Syt 7, Syt , and Syt ß isoforms, we generated antisera to recombinant proteins encompassing the C2 domains of each protein. For Syt 14, we prepared antisera to a recombinant protein that encompassed the linker between the TM domain and C2A. For Syt 12, we generated antisera against a peptide derived from the linker domain between C2A and C2B. Each respective sequence was cloned into pGEX vectors. Recombinant GST fusion proteins were expressed and processed in E. coli (BL21) according to standard laboratory protocols. The fusion proteins were purified in batch using glutathione-sepharose (Amersham Biosciences). To remove the GST affinity tag, protein samples were incubated with thrombin for 1 h at RT. Antisera was purified using affinity chromatography. The domain of each synaptotagmin was coupled to a 1-ml NHS-activated sepharose column (Amersham Biosciences). Antisera (2 ml) injection, subsequent washes, and elution from the columns were all performed on an AKTA FPLC (Amersham Biosciences). Columns were washed in 20 mM sodium phosphate and eluted with 0.1 M glycine, pH 2.7. To minimize denaturation of the antibody at low pH, the eluted fractions were immediately mixed with 1 M Tris, pH 9. Fractions containing the desired peak were concentrated using Amicon ultra centrifugal filter devices (Millipore), aliquoted, and stored at 80°C.
Immunostaining
Embryos and larvae were immunostained as described previously (Yoshihara and Littleton, 2002; Rieckhof et al., 2003). The dilution of primary antibodies was as follows: Syt 1 (1:1,000) Syt 4 (1:500), Syt 7 (1:1,000), Syt (1:2,000), and Syt ß (1:500). To decrease background, antibodies were preabsorbed to 011-h embryos. Samples were washed and mounted in 70% glycerol. Cy2-conjugated goat
rabbit secondary antibodies (Jackson ImmunoResearch Laboratories) were used at 1:200. Visualization was performed under light microscopy using a 40x oil-immersion lens. Images were taken using confocal microscopy on a microscope (model Axoplan 2; Carl Zeiss MicroImaging, Inc.) and processed with PASCAL software (Carl Zeiss MicroImaging, Inc.).
Electrophysiology analysis
Electrophysiological analysis of wandering stage third instar larva was performed in Drosophila saline (70 mM NaCl, 5 mM KCl, 4 mM MgCl2, 10 mM NaHCO3, 5 mM Trehalose, 115 mM sucrose, and 5 mM Hepes-NaOH, pH 7.2) supplemented with either 1.5 mM or 5.0 mM CaCl2 using an Axoclamp 2B amplifier (Axon Instruments, Inc.) at 22°C as described previously (Rieckhof et al., 2003).
Larval locomotion analysis
To quantify larval locomotion, late third instar larvae grown at 25°C were collected and placed on a flat layer of 2.9% agar supplemented with grape juice. Quantification of larval locomotion parameters was performed as described previously (Saraswati et al., 2004). For quantification of cycle duration, video recording of locomotion was performed using a digital video camera (model XL1S; Canon) attached to a 16x zoom lens (field of 3 cm2). Cycle duration was reconstructed offline by digitizing frame-by-frame locomotor contractions.
Online supplemental material
The specificity of the Syt and Syt ß antibodies for immunocytochemistry is shown in Fig. S1. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200312054/DC1.
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Acknowledgments |
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This work was supported by grants from the National Institutes of Health, the Human Frontiers Science Program Organization, the Packard Foundation, and the Searle Scholars Program. J.T. Littleton is an Alfred P. Sloan Research Fellow.
Submitted: 5 December 2003
Accepted: 10 June 2004
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References |
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![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Adams, M.D., S.E. Celniker, R.A. Holt, C.A. Evans, J.D. Gocayne, P.G. Amanatides, S.E. Scherer, P.W. Li, R.A. Hoskins, R.F. Galle, et al. 2000. The genome sequence of Drosophila melanogaster. Science. 287:21852195.
Adolfsen, B., and J.T. Littleton. 2001. Genetic and molecular analysis of the synaptotagmin family. Cell. Mol. Life Sci. 58:393402.[Medline]
Aparicio, S., J. Chapman, E. Stupka, N. Putnam, J.M. Chia, P. Dehal, A. Christoffels, S. Rash, S. Hoon, A. Smit, et al. 2002. Whole-genome shotgun assembly and analysis of the genome of Fugu rubripes. Science. 297:13011310.
Butz, S., R. Fernandez-Chacon, F. Schmitz, R. Jahn, and T.C. Südhof. 1999. The subcellular localizations of atypical synaptotagmins III and VI. Synaptotagmin III is enriched in synapses and synaptic plasma membranes but not in synaptic vesicles. J. Biol. Chem. 274:1829018296.
Cantera, R., and D.R. Nassel. 1992. Segmental peptidergic innervation of abdominal targets in larval and adult dipteran insects revealed with an antiserum against leucokinin I. Cell Tissue Res. 269:459471.[Medline]
C. elegans Sequencing Consortium. 1998. Genome sequence of the nematode C. elegans: a platform for investigating biology. Science. 282:20122018.
Desai, R.C., B. Vyas, C.A. Earles, J.T. Littleton, J.A. Kowalchyck, T.F. Martin, and E.R. Chapman. 2000. The C2B domain of synaptotagmin is a Ca2+-sensing module essential for exocytosis. J. Cell Biol. 150:11251136.
DiAntonio, A., R.W. Burgess, A.C. Chin, D.L. Deitcher, R.H. Scheller, and T.L. Schwarz. 1993. Identification and characterization of Drosophila genes for synaptic vesicle proteins. J. Neurosci. 13:49244935.[Abstract]
Fernàndez-Chacòn, R., A. Königstorfer, S.H. Gerber, J. Garcìa, M.F. Matos, C.F. Stevens, N. Brose, J. Rizo, C. Rosenmund, and T.C. Südhof. 2001. Synaptotagmin I functions as a calcium regulator of release probability. Nature. 410:4149.[CrossRef][Medline]
Geppert, M., Y. Goda, R.E. Hammer, C. Li, T.W. Rosahl, C.F. Stevens, and T.C. Südhof. 1994. Synaptotagmin I: a major Ca2+ sensor for transmitter release at a central synapse. Cell. 79:717727.[Medline]
Holt, R.A., G.M. Subramanian, A. Halpern, G.G. Sutton, R. Charlab, D.R. Nusskern, P. Wincker, A.G. Clark, J.M. Ribeiro, R. Wides, et al. 2002. The genome sequence of the malaria mosquito Anopheles gambiae. Science. 298:129149.
Ibata, K., T. Hashikawa, T. Tsuboi, S. Terakawa, F. Liang, A. Mizutani, M. Fukuda, and K. Mikoshiba. 2002. Non-polarized distribution of synaptotagmin IV in neurons: evidence that synaptotagmin IV is not a synaptic vesicle protein. Neurosci. Res. 43:401406.[CrossRef][Medline]
Jahn, R., T. Lang, and T.C. Südhof. 2003. Membrane fusion. Cell. 112:519533.[Medline]
Koh, T.W., and H.J. Bellen. 2003. Synaptotagmin I, a Ca2+ sensor for neurotransmitter release. Trends Neurosci. 26:413422.[CrossRef][Medline]
Lander, E.S., L.M. Linton, B. Birren, C. Nusbaum, M.C. Zody, J. Baldwin, K. Devon, K. Dewar, M. Doyle, W. FitzHugh, et al. 2001. Initial sequencing and analysis of the human genome. Nature. 409:860921.[CrossRef][Medline]
Littleton, J.T., H.J. Bellen, and M.S. Perin. 1993. Expression of synaptotagmin in Drosophila reveals transport and localization of synaptic vesicles to the synapse. Development. 118:10771088.
Littleton, J.T., M. Stern, M. Perin, and H.J. Bellen. 1994. Calcium dependence of neurotransmitter release and rate of spontaneous vesicle fusions are altered in Drosophila synaptotagmin mutants. Proc. Natl. Acad. Sci. USA. 91:1088810892.
Littleton, J.T., T.L. Serano, G.M. Rubin, B. Ganetzky, and E.R. Chapman. 1999. Synaptic function modulated by changes in the ratio of synaptotagmin I and IV. Nature. 400:757760.[CrossRef][Medline]
Lledo, P.M., X. Zhang, T.C. Südhof, R.C. Malenka, and R.A. Nicoll. 1998. Postsynaptic membrane fusion and long-term potentiation. Science. 279:399403.
Ludwig, M., N. Sabatier, P.M. Bull, R. Landgraf, G. Dayanithi, and G. Leng. 2002. Intracellular calcium stores regulate activity-dependent neuropeptide release from dendrites. Nature. 418:8589.[CrossRef][Medline]
Lloyd, T.E., P. Verstreken, E.J. Ostrin, A. Phillippi, O. Lichtarge, and H.J. Bellen. 2000. A genome-wide search for synaptic vesicle cycle proteins in Drosophila. Neuron. 26:4550.[Medline]
Lloyd, T.E., R. Atkinson, M.N. Wu, Y. Zhou, G. Pennetta, and H.J. Bellen. 2002. Hrs regulates endosome membrane invagination and tyrosine kinase receptor signaling in Drosophila. Cell. 108:261269.[Medline]
Maletic-Savatic, M., and R. Malinow. 1998. Calcium-evoked dendritic exocytosis in cultured hippocampal neurons. Part I: trans-Golgi network-derived organelles undergo regulated exocytosis. J. Neurosci. 18:68036813.
Martinez, I., S. Chakrabarti, T. Hellevik, J. Morehead, K. Fowler, and N.W. Andrews. 2000. Synaptotagmin VII regulates Ca2+-dependent exocytosis of lysosomes in fibroblasts. J. Cell Biol. 148:11411149.
O'Brien, M.A., and P.H. Taghert. 1998. A peritracheal neuropeptide system in insects: release of myomodulin-like peptides at ecdysis. J. Exp. Biol. 201(Pt 2):193209.
Perin, M.S., V.A. Fried, G.A. Mignery, R. Jahn, and T.C. Südhof. 1990. Phospholipid binding by a synaptic vesicle protein homologous to the regulatory region of protein kinase C. Nature. 345:260263.[CrossRef][Medline]
Rieckhof, G.E., M. Yoshihara, Z. Guan, and J.T. Littleton. 2003. Presynaptic N-type calcium channels regulate synaptic growth. J. Biol. Chem. 278:4109941108.
Robinson, I.M., R. Ranjan, and T.L. Schwarz. 2002. Synaptotagmins I and IV promote transmitter release independently of Ca2+ binding in the C2A domain. Nature. 418:336340.[CrossRef][Medline]
Saegusa, C., M. Fukuda, and K. Mikoshiba. 2002. Synaptotagmin V is targeted to dense-core vesicles that undergo calcium-dependent exocytosis in PC12 cells. J. Biol. Chem. 277:2449924505.
Salzberg, A., N. Cohen, N. Halachmi, Z. Kimchie, and Z. Lev. 1993. The Drosophila Ras2 and Rop gene pair: a dual homology with a yeast Ras-like gene and a suppressor of its loss-of-function phenotype. Development. 117:13091319.
Saraswati, S., L.E. Fox, D.R. Soll, and C.F. Wu. 2004. Tyramine and octopamine have opposite effects on the locomotion of Drosophila larvae. J. Neurobiol. 58:425441.[CrossRef][Medline]
Schulze, K., K. Broadie, M. Perin, and H.J. Bellen. 1995. Genetic and electrophysiological studies of Drosophila syntaxin-1A demonstrate its role in nonneuronal secretion and neurotransmission. Cell. 80:311320.[Medline]
Stevens, C.F., and J.M. Sullivan. 2003. The synaptotagmin C2A domain is part of the calcium sensor controlling fast synaptic transmission. Neuron. 39:299308.[Medline]
Südhof, T.C. 2002. Synaptotagmins: why so many? J. Biol. Chem. 277:76297632.
Sugita, S., W. Han, S. Butz, X. Liu, R. Fernandez-Chacon, Y. Lao, and T.C. Südhof. 2001. Synaptotagmin VII as a plasma membrane Ca2+ sensor in exocytosis. Neuron. 30:459473.[CrossRef][Medline]
Sugita, S., O.H. Shin, W. Han, Y. Lao, and T.C. Südhof. 2002. Synaptotagmins form a hierarchy of exocytotic Ca2+ sensors with distinct Ca2+ affinities. EMBO J. 21:270280.
Tucker, W.C., J.M. Edwardson, J. Bai, H.J. Kim, T.F. Martin, and E.R. Chapman. 2003. Identification of synaptotagmin effectors via acute inhibition of secretion from cracked PC12 cells. J. Cell Biol. 162:199209.
Wang, C.T., R. Grishanin, C.A. Earles, P.Y. Chang, T.F. Martin, E.R. Chapman, and M.B. Jackson. 2001. Synaptotagmin modulation of fusion pore kinetics in regulated exocytosis of dense-core vesicles. Science. 294:11111115.
Wang, C.T., J.C. Lu, J. Bai, P.Y. Chang, T.F. Martin, E.R. Chapman, and M.B. Jackson. 2003. Different domains of synaptotagmin control the choice between kiss-and-run and full fusion. Nature. 424:943947.[CrossRef][Medline]
Waterston, R.H., K. Lindblad-Toh, E. Birney, J. Rogers, J.F. Abril, P. Agarwal, R. Agarwala, R. Ainscough, M. Alexandersson, P. An, et al. 2002. Initial sequencing and comparative analysis of the mouse genome. Nature. 420:520562.[CrossRef][Medline]
Yamada, W.M., and R.S. Zucker. 1992. Time course of transmitter release calculated from simulations of a calcium diffusion model. Biophys. J. 61:671682.[Abstract]
Yoshihara, M., and J.T. Littleton. 2002. Synaptotagmin I functions as a calcium sensor to synchronize neurotransmitter release. Neuron. 36:897908.[Medline]
Yoshihara, M., B. Adolfsen, and J.T. Littleton. 2003. Is synaptotagmin the calcium sensor? Curr. Opin. Neurobiol. 13:315323.[CrossRef][Medline]