Elsevier

Molecular and Cellular Neuroscience

Volume 56, September 2013, Pages 244-254
Molecular and Cellular Neuroscience

Molecular mechanisms of COMPLEXIN fusion clamp function in synaptic exocytosis revealed in a new Drosophila mutant

https://doi.org/10.1016/j.mcn.2013.06.002Get rights and content

Abstract

The COMPLEXIN (CPX) proteins play a critical role in synaptic vesicle fusion and neurotransmitter release. Previous studies demonstrated that CPX functions in both activation of evoked neurotransmitter release and inhibition/clamping of spontaneous synaptic vesicle fusion. Here we report a new cpx mutant in Drosophila melanogaster, cpx1257, revealing spatially defined and separable pools of CPX which make distinct contributions to the activation and clamping functions. In cpx1257, lack of only the last C-terminal amino acid of CPX is predicted to disrupt prenylation and membrane targeting of CPX. Immunocytochemical analysis established localization of wild-type CPX to active zone (AZ) regions containing neurotransmitter release sites as well as broader presynaptic membrane compartments including synaptic vesicles. Parallel biochemical studies confirmed CPX membrane association and demonstrated robust binding interactions of CPX with all three SNAREs. This is in contrast to the cpx1257 mutant, in which AZ localization of CPX persists but general membrane localization and, surprisingly, the bulk of CPX–SNARE protein interactions are abolished. Furthermore, electrophysiological analysis of neuromuscular synapses revealed interesting differences between cpx1257 and a cpx null mutant. The cpx null exhibited a marked decrease in the EPSC amplitude, slowed EPSC rise and decay times and an increased mEPSC frequency with respect to wild-type. In contrast, cpx1257 exhibited a wild-type EPSC with an increased mEPSC frequency and thus a selective failure to clamp spontaneous release. These results indicate that spatially distinct and separable interactions of CPX with presynaptic membranes and SNARE proteins mediate separable activation and clamping functions of CPX in neurotransmitter release.

Introduction

It is widely accepted that SNARE proteins function at the core of the neurotransmitter release apparatus, where they promote exocytotic fusion of neurotransmitter-filled synaptic vesicles with the presynaptic plasma membrane (Jahn and Scheller, 2006). However, defining the mechanisms which provide precise and rapid regulation of synaptic vesicle fusion remains among the foremost problems in cellular and molecular neuroscience. The identification of CPX as a protein which binds and regulates SNARE complexes (Ishizuka et al., 1995, McMahon et al., 1995) has advanced our understanding of these mechanisms (Brose, 2008, Neher, 2010, Rizo and Rosenmund, 2008, Stein and Jahn, 2009, Südhof and Rothman, 2009). Notably, CPX can both promote SV fusion evoked by a presynaptic action potential and suppress or “clamp” spontaneous vesicle fusion. Recent models suggest that specific domains of CPX (Fig. 1A) contribute to different aspects of synaptic vesicle fusion (Hobson et al., 2011, Martin et al., 2011, Maximov et al., 2009, Reim et al., 2001, Strenzke et al., 2009, Tang et al., 2006, Xue et al., 2007, Xue et al., 2009, Xue et al., 2010, Yang et al., 2010). Whereas a “central helix” which binds SNARE complexes (Bracher et al., 2002, Chen et al., 2002) is absolutely required for CPX function, other domains appear to mediate specific aspects of CPX activity (Rizo and Rosenmund, 2008, Stein and Jahn, 2009). For example, recent studies have shown that the CPX C-terminus is specifically required for the clamping function (Buhl et al., 2013, Cho et al., 2010, Kaeser-Woo et al., 2012, Martin et al., 2011, Xue et al., 2009). Of particular relevance to the present study is a specific CaaX motif found at the extreme C-terminus of several mammalian and Drosophila CPX isoforms. This motif has been shown to mediate CPX prenylation [a form of lipid modification; (Omer and Gibbs, 1994, Resh, 2006)] and has been implicated in both targeting CPX to membranes (Reim et al., 2005) and the CPX clamping function (Cho et al., 2010, Xue et al., 2009). The form of prenylation demonstrated for mammalian CPX isoforms (CPX3 and 4) is farnesylation (Reim et al., 2005), consistent with previous studies indicating that one of several specific residues in the X position of the CaaX motif (A,C,M,Q,S) selectively mediates farnesylation (Omer and Gibbs, 1994).

This progress is extended by new insights gained from the present study, in which the isolation and characterization of a new cpx mutant further define the in vivo molecular basis of CPX functions and interactions within the neurotransmitter release apparatus. This study reveals a specific subcellular distribution for CPX within the presynaptic terminal and a role for C-terminal farnesylation in mediating both association of CPX with presynaptic membranes and CPX clamping of spontaneous synaptic vesicle fusion.

Section snippets

Genetic and molecular characterization of a new cpx mutant

Further genetic analysis to examine the in vivo molecular mechanisms of CPX function was pursued through a forward genetic screen for new mutant alleles of the single Drosophila cpx gene. To complement a previously reported cpx null mutant (Huntwork and Littleton, 2007), this screen was intended to recover hypomorphic and conditional alleles that may further define the in vivo molecular determinants of CPX function. A screen was performed using chemical mutagenesis and subsequent screening for

Discussion

Through characterization of a new Drosophila cpx mutant, the present study advances our understanding of the molecular mechanisms mediating CPX function in neurotransmitter release. Our findings provide new information about the subcellular distribution of CPX with respect to presynaptic membranes, as well as its molecular basis, and implicate CPX membrane association as a critical element in its clamping function and in vivo interactions with SNARE proteins.

As shown in Fig. 8, our working

Drosophila strains

Appl-GAL4 and w;;Ly/TM6c were from our laboratory stock collection. The cpxSH1 null mutant and the UAS-cpx transgenic line were generously provided by Troy Littleton (MIT, Cambridge, MA). Deficiency lines, Df(3L)GN34 and Df(3R)Exel6140, were obtained from the Bloomington Stock Center. UAS-EGFP-cpx and UAS-EGFP-cpx1257 transgenic lines were generated in the current study (see “Generation of transgenic lines”). Stocks and crosses were cultured on a conventional cornmeal–molasses–yeast medium at 20

Conflict of interest

The authors have no conflict of interest in submitting this manuscript.

Acknowledgments

A cpx null mutant stock, a UAS-cpx transgenic line and an anti-CPX antibody were generously provided by Troy Littleton (MIT, Cambridge, MA). We are also grateful to Noreen Reist (Colorado State University, Fort Collins, CO) and David Deitcher (Cornell University, Ithaca, NY) for providing anti-SYT and anti-SNAP25 antibodies, respectively. We thank Richard Ordway (Penn State University) for his continuous encouragement and invaluable discussion throughout this work. This study was supported by

References (51)

  • R.A. Mathias et al.

    Triton X-114 phase separation in the isolation and purification of mouse liver microsomal membrane proteins

    Methods

    (2011)
  • H.T. McMahon et al.

    Complexins: cytosolic proteins that regulate SNAP receptor function

    Cell

    (1995)
  • E. Neher

    Complexin: does it deserve its name?

    Neuron

    (2010)
  • S. Pabst et al.

    Rapid and selective binding to the synaptic SNARE complex suggests a modulatory role of complexins in neuroexocytosis

    J. Biol. Chem.

    (2002)
  • K. Reim et al.

    Complexins regulate a late step in Ca2 +-dependent neurotransmitter release

    Neuron

    (2001)
  • A.T. Reinicke et al.

    A Salmonella typhimurium effector protein SifA is modified by host cell prenylation and S-acylation machinery

    J. Biol. Chem.

    (2005)
  • F. Seiler et al.

    A role of complexin–lipid interactions in membrane fusion

    FEBS Lett.

    (2009)
  • A. Stein et al.

    Complexins living up to their name—new light on their role in exocytosis

    Neuron

    (2009)
  • J. Tang et al.

    A complexin/synaptotagmin 1 switch controls fast synaptic vesicle exocytosis

    Cell

    (2006)
  • K. Weninger et al.

    Accessory proteins stabilize the acceptor complex for synaptobrevin, the 1:1 syntaxin/SNAP-25 complex

    Structure

    (2008)
  • R.T. Wragg et al.

    Synaptic vesicles position complexin to block spontaneous fusion

    Neuron

    (2013)
  • M. Xue et al.

    Tilting the balance between facilitatory and inhibitory functions of mammalian and Drosophila Complexins orchestrates synaptic vesicle exocytosis

    Neuron

    (2009)
  • X. Yang et al.

    Complexin clamps asynchronous release by blocking a secondary Ca2 + sensor via its accessory α helix

    Neuron

    (2010)
  • W. Yu et al.

    Activity-dependent interactions of NSF and SNAP at living synapses

    Mol. Cell. Neurosci.

    (2011)
  • A. Bracher et al.

    X-ray structure of a neuronal complexin–SNARE complex from squid

    J. Biol. Chem.

    (2002)
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