Review Article
The presynaptic active zone: A dynamic scaffold that regulates synaptic efficacy

https://doi.org/10.1016/j.yexcr.2015.02.011Get rights and content

Highlights

  • Recent progress linking presynaptic active zone ultrastructure to function.

  • Dynamic properties of the active zone.

  • Role of the active zone in the regulation of synaptic efficacy.

Abstract

Before fusing with the presynaptic plasma membrane to release neurotransmitter into the synaptic cleft synaptic vesicles have to be recruited to and docked at a specialized area of the presynaptic nerve terminal, the active zone. Exocytosis of synaptic vesicles is restricted to the presynaptic active zone, which is characterized by a unique and highly interconnected set of proteins. The protein network at the active zone is integrally involved in this process and also mediates changes in release properties, for example in response to alterations in the level of neuronal network activity. In recent years the development of novel techniques has greatly advanced our understanding of the molecular identity of respective active zone components as well as of the ultrastructure of this membranous subcompartment and of the SV release machinery. Furthermore, active zones are now viewed as dynamic structures whose composition and size are correlated with synaptic efficacy. Therefore, the dynamic remodeling of the protein network at the active zone has emerged as one potential mechanism underlying acute and long-term synaptic plasticity. Here, we will discuss this recent progress and its implications for our view of the role of the AZ in synaptic function.

Introduction

Information transfer in the brain, from acute initiation of movement to higher cognitive functions like memory and emotions, takes place at contact sites between neurons and their target cells. At this cell to cell connection, the synapse, information arrives in form of an electrical signal and is transmitted to the target cell via neurotransmitters, chemical signaling molecules. This information transfer is mediated by the fusion of neurotransmitter filled synaptic vesicles (SVs) at a highly specialized area of the presynaptic plasma membrane, the active zone (AZ). The AZ is characterized by an electron-dense protein network, the cytomatrix of the active zone (CAZ), and a precise alignment with the postsynaptic reception apparatus, the postsynaptic density (PSD). To achieve the high spatial and temporal precision of synaptic transmission the process of SV fusion is tightly regulated. Before SVs can fuse with the plasma membrane in response to Ca2+-influx through voltage-gated calcium channels (VGCCs) they have to be recruited to and docked at the AZ followed by a maturation process (“priming”) that renders them fusion-competent (reviewed in [1]). Presynaptic active zones are integrally involved in multiple processes that control and enable the high speed, temporal and spatial control as well as plasticity of synaptic vesicle fusion: (1) Through specific trans-synaptic cell adhesion molecules AZs coordinate the precise opposition of the pre- and postsynaptic specializations; (2) AZs contribute to the spatial restraint of the fusion process as SVs dock and fuse at the AZ; (3) AZ components play a role in the recruitment of VGCCs to the presynaptic plasma membrane and in the efficient coupling of SVs to VGCCs, which in turn is crucial for the high speed of synchronous release; (4) the AZ contributes to the sorting of SV proteins to the endocytic machinery and (5) AZ proteins play important roles in the processes mediating activity-induced presynaptic short- and long-term plasticity. However, there are still many open questions regarding the functional contribution of individual AZ components to the individual steps of the SV cycle. In this review we will focus on recent progress in our understanding of the functioning of the molecular machinery at the AZ, in particular on novel insights into the correlation between ultrastructure, molecular composition and function.

In contrast to the proteomes of synaptosomes, synaptic vesicles and the postsynaptic density (PSD), which have been characterized in multiple studies (reviewed in [2]), the composition of the cytomatrix at the presynaptic active zone has remained less well defined, mainly due to difficulties in obtaining reasonably pure preparations. However, analyses of genetic mutations, mainly in the worm Caenorhabditis elegans, the fruit fly Drosophila and rodents, as well as of protein–protein interactions have revealed five evolutionarily conserved proteins, RIM (Rab3-interacting molecule), Munc13, ELKS/CAST/Bruchpilot (glutamic acid (E), leucine (L), lysine (K), and serine (S)-rich protein; CAZ-associated structural proteins), α-Liprin/syd-2 (synapse-defective 2), and RIM-BP (RIM-binding protein), that are highly enriched at and form the core of the cytomatrix at the active zone (CAZ) (reviewed in [2], [3], [4]). In vertebrates, two additional large homologous proteins, Bassoon and Piccolo/Aczonin, are associated with active zones. Moreover, in mammals the five core active zone proteins are encoded by multiple genes, which are further diversified by alternative splicing. Via their conserved domains, like PDZ, SH3, coiled-coil, SAM, Zn2+-finger, and C2, the core CAZ components can interact with each other, form oligomers and a tight protein network that regulates synaptic vesicle fusion. Furthermore, these multiple connections are thought to contribute to the synapse׳s ability to both alter its efficacy in response to activity (plasticity) and to maintain properties over long periods of time (tenacity). It is important to note that the core active zone proteins are not exclusively localized to this specialized area of the presynaptic plasma membrane but are only strongly enriched at the AZ. They are partially expressed also in neuroendocrine and neurosecretory or even non-neuronal cells. In recent years different strategies have been employed to isolate presynaptic fractions of high purity and yield in order to identify the complete AZ proteome using an unbiased approach. Here by, synaptosomes were treated by hypo-osmotic shock to release two populations of synaptic vesicles, one free and a second one that remained docked to the presynaptic plasma membrane. Both fractions were further purified by immunoisolation using antibodies against SV proteins [5], [6], [7]. In particular two methodological improvements introduced by Boyken et al., separation of the pre- from the postsynaptic compartment by a mild proteolytic digestion of the synaptosomes before the hypo-osmotic lysis and the quantitative analysis of the composition of the “free” versus the “docked” SV fraction, provided for the first time quantitative information about the protein complement of the SV docking complex and the AZ [6]. In addition to the core AZ proteins, ion channels, transporters and cell adhesion molecules with known presynaptic functions 30 so far uncharacterized proteins were identified, of which around 50% were strongly enriched in the “docked” SV fraction. Future studies will be required to determine if these proteins play a role in the regulation of AZ structure and function. An important finding of this study was the observation that glutamatergic and GABAergic docking complexes exhibit only few quantitative differences indicating that the release machineries of these two synapse types are very similar or even identical. This is in contrast to the molecular composition of the postsynaptic scaffolding and signaling complexes, which differ significantly between excitatory and inhibitory synapses [8], [9]. However, synapse-type specific proteins might have been missed in this global proteomics approach due to their low abundance. Furthermore, synapses throughout the brain exhibit considerable morphological and functional heterogeneity [10]. This heterogeneity might be mediated by a diversity in their molecular composition, for example by the exact complement of alternatively spliced variants or family members of AZ proteins present in a specific synapse type or even in individual synapses. The recently reported purification of synaptosomes from distinct synaptic subtypes using the Fluorescence Activated Synaptosome Sorting (FASS) method together with high-resolution imaging techniques for validation and fine mapping of the distribution of distinct proteins will allow to address these open questions [10], [11].

Recently, Wilhelm et al. have reported a first quantitative molecular-scale model of an “average” synapse. They have used an integrative approach combining quantitative immunoblotting and mass spectrometry to determine the abundance and super-resolution-fluorescence microscopy to define the localization of individual proteins in the presynaptic nerve terminal [12]. By analyzing the position of proteins relative to release sites with the resolution of stimulated emission depletion (STED) microscopy in three different preparations, synaptosomes, cultured hippocampal neurons and the levator auris longus neuromuscular junction they obtained the relative spatial distribution of each protein. They observed the tested AZ core proteins to be mostly confined to the active zone area, which their 3D model suggests to be rather crowded. Even though the abundance of proteins correlated if they were involved in the same steps of the SV cycle, differences were detected within groups. For example, of the AZ proteins RIM had the lowest (38 molecules) and Munc13 the highest (1551 molecules) copy number (Bassoon, 446, and Piccolo, 100, in contrast, copy numbers of the non-AZ enriched proteins Munc18 and Rab3 were, 4253 and 18,846, respectively). However, these numbers will have to be validated using an independent approach, for example by super resolution microscopy that allows quantification of individual molecules. As the density and relative spatial distribution within the presynaptic nerve terminal of proteins and SVs impacts their mobility and their access to specific subcellular localizations like the AZ and thereby directly affects the properties of synaptic transmission these results are an important step in improving our understanding of how synapse organization is linked to function. Therefore, models will have to be developed that take heterogeneity of synaptic morphology and molecular composition into account.

Technical advances in recent years have also greatly advanced our view of the AZ ultrastructure and of the correlation between ultrastructure, molecules and function. Electronmicrographs of chemically fixed and stained central nervous system (CNS) mammalian synapses revealed the AZ to be composed of a hexagonal grid of electron dense projections and intercalated SVs [13]. On the other hand, cryo-ET of unfixed, vitrified and frozen-hydrated synapses did not detect dense projections but revealed vesicles to be connected to the plasma membrane (“tethers”, 5–20 nm) and to each other (“connectors”, around 10 nm) via filaments of varying length [14]. Nevertheless, the consensus from all ultrastructural approaches suggests the existence of a structural scaffold at the AZ that mediates the spatial positioning of SVs within the AZ and to the sites of release (reviewed in [3]). Immuno-EM and high-resolution light microscopy have provided first insights into the molecular nature of the electron-dense projections suggesting that RIM and Munc13 are located adjacent to the plasma membrane between the dense projections. Bassoon and Piccolo were detected further away from the membrane and rather on the tips of the electron dense projections [15], [16]. Through high-resolution light microscopy analyses of AZs at the neuromuscular junction of Drosophila ultrastructural elements, like the T-bar, could also be attributed to specific CAZ components (for example Bruchpilot [17], RIM-BP [18], and α-Liprin [19]). Further evidence for the molecular identity of ultrastructural AZ elements has come from the analysis of genetic mutants deficient for specific CAZ proteins. Cryo-ET analyses of vitrified, frozen-hydrated synaptosomes prepared from RIM1α knock-out (KO) mice revealed a reduction in the number of AZ proximal SVs and of tethers per unit AZ surface in about 50% of synapses indicating a critical role for RIM1α in SV tethering [20]. Consistently, classical EM studies of aldehyde-fixed synapses lacking all RIM1 and RIM2 isoforms [21], [22] and EM tomography of C. elegans synapses [23] also detected a reduction in the number of SVs adjacent to the plasma membrane. Deletion of other core AZ scaffold proteins also had an impact on SV tethering as a decrease in the attachment of SVs to AZs was observed in mutants of syd-2, the single α-Liprin gene in C. elegans,[23] and synapses deficient for either Liprin-α2 [24] or Piccolo and Bassoon [25]. However, a classical EM analysis of synapses deficient for ELKS1 and ELKS2 did not find any ultrastructural alterations [26]. The sequential steps in SV AZ recruitment (tethering) and membrane attachment (docking) were recently dissected for the first time in a comparative systematic study analyzing cryo-fixed organotypic hippocampal slice cultures from various mice lacking key synaptic proteins by 3D-electron tomography [27]. Imig et al. propose that SV docking/priming is preceded by a multi-step tethering process (Fig. 1). In their model SVs are first recruited to the AZ with the participation of AZ proteins before being tethered in close proximity to the AZ plasma membrane (around 10 nm). Subsequently, SVs get even closer to the AZ (around 6 nm) with the help of Munc13s, CAPSs, and/or additional yet unidentified proteins before trans-SNARE complex assembly for final docking/priming. Applying this approach to synapses deficient for the remaining AZ core proteins will further advance our understanding of the steps between SV recruitment and SNARE complex formation. Furthermore, it will be interesting to perform these comparative analyses under different physiological conditions, for example after the induction of synaptic plasticity or after neuronal network silencing.

The action-potential induced Ca2+-influx is spatially restricted to a microdomain around the VGCCs (radius of approximately 100 nm). Therefore, the structural organization of the release machinery at the AZ, in particular the precise distance between the Ca2+-sensor on the SV and the VGCCs, directly impacts release properties. Accordingly, the molecular mechanisms underlying this positional priming and the physical coupling of the SVs to the VGCCs at release sites have been a topic of intense research. One mechanism that localizes Ca2+-channels to the sites of release involves a tripartite complex composed of RIM, RIM-BP and Cav2 channels [21], [22]. Support for this functional role of the tripartite complex also comes from analyses at the NMJ of Drosophila, where it was observed that RIM-BP tethers tagged Ca2+-channels to the AZ and promotes Ca2+-influx [18] and normal Ca2+-influx requires RIM [28]. Furthermore, it was shown that at the Drosophila NMJ Bruchpilot promotes Ca2+-channel clustering at the AZ [17]. However, removal of ELKS from inhibitory hippocampal synapses in mice did not dramatically reduce the presynaptic localization of Ca2+-channels but impaired presynaptic Ca2+-influx indicating a role for ELKS in the modulation of channel opening [26]. At the AZ of central synapses Ca2+-influx is mainly mediated by channels containing the Cav2.1 and Cav2.2 pore-forming subunits. The relative contribution of Cav2.1 and Cav2.2 to neurotransmitter release differs between individual synapses and this might impact spontaneous and evoked SV fusion as well as synaptic plasticity. Recently, a first mechanism for the differential recruitment of the two channel types to release sites was reported. Davydova et al. showed that Bassoon via an interaction with RIM-BPs specifically localizes Cav2.1 to AZs [29]. Taken together, these findings suggest that multiple targeting mechanisms coexist to ensure the localization of specific Ca2+-channels to the requisite release sites. Future studies will have to resolve how these different modes of Ca2+-channel tethering are regulated and how they contribute to the plasticity of release properties. Another important factor for the physical coupling of SVs to VGCCs is the topography of VGCCs at the release sites in presynaptic AZs. The availability of improved Ca2+-channel antibodies has finally shed light onto this open question. Immuno-EM analyses of hippocampal [30] and cerebellar [31] synapses showed VGCCs to be clustered at AZs. Furthermore, immunolabelings of detergent-digested freeze-fracture replicas (SDS-FRL) of the calyx of Held also revealed clusters of VGCCs [32]. In order to further resolve synapse-type specific release coupling mechanisms it will be essential to determine the topography of the SV to VGCC cluster relationship in addition to the functional measurement of the coupling distance between SV and the calcium source.

During the last decade the concept of the synapse as a rigid structure with a static molecular composition and organization, which is only driven to change when triggered by physiological stimuli, has evolved to one that views synapses as a dynamic assembly of proteins. This change in concept was put forward based on results obtained from imaging of fluorescently tagged synaptic proteins in live neurons (reviewed in [33]). This approach revealed that SVs and SV-associated molecules shuttle between neighboring presynaptic sites at surprisingly high exchange rates. The exchange rates for CAZ core proteins, which were tightly integrated into the AZ protein matrix were much slower, with time constants (exchange of around two thirds of the molecules) for Bassoon being≥8 h [34] and for Munc13-1 around 1.5 h [35] (preliminary experiments indicate time constants for RIM1 and Liprin-α2 also on the order of several hours (Zürner, Schoch, Ziv, unpublished data)). Therefore, the AZ is a dynamic protein network; however within the even more dynamic presynaptic bouton it represents a relatively stable core, which could define and maintain individual synaptic properties over time scales of hours. This notion is further supported by the observation that at individual synapses the ratio between the levels of Munc13-1 and the PSD molecule PSD-95 are well correlated over the same time scale despite the spontaneous variations in protein amounts [36]. Based on results from fluorescence recovery after photobleaching experiments (FRAP) it has been suggested that AZ proteins exist at presynapses in two pools, a relatively small immobile pool and a larger mobile one, and it was hypothesized that changes in the relative distribution of certain AZ components in the mobile and immobile fraction are directly linked to synaptic efficacy [24] (Fig. 2). Analysis of Liprin-α2 knock-down neurons revealed a decrease in the mobility of its binding partners RIM1 and CASK and a concurrent reduction of synaptic output indicating a role for Liprin-α2 in the modulation of AZ dynamics. Dissociation of Bassoon from the immobile fraction of the CAZ was shown to be regulated by phosphorylation as phosphorylation-dependent binding of Bassoon to the adapter protein 14-3-3 decreased its attachment to the CAZ [37]. Both findings suggest that cooperating and competitive protein–protein interactions form the basis for controlling AZ dynamics and that these interactions in turn are regulated by post-translational modifications like phosphorylation.

Measurement of presynaptic [Ca2+] transients using two-photon imaging and subsequent electron microscopic reconstructions of the presynaptic AZ revealed a tight correlation between AZ area and the release probability [30]. Furthermore, the number of Cav2.1 Ca2+-channels and of RIM1/2 molecules was found to be proportional to the AZ area indicating that the number of functional release sites and thereby synaptic efficacy scales with the size of the AZ. Accordingly, also synaptic protein levels of a fluorescently-tagged Bassoon correlated well with AZ area and release probability and spontaneous alterations in the synaptic abundance of Bassoon were associated with corresponding changes in synaptic strength [38]. Taken together, these findings suggest that both fluctuations in the total levels of CAZ proteins as well as in the relative amounts associated with the mobile/immobile AZ fraction directly affect synaptic release properties. However, it has to be considered that heterozygous knock-out mice of Bassoon, Piccolo, RIM1, and Munc13-1 for example, do not show clear phenotypes, even though the amount of the respective proteins is decreased to around 50%. Therefore, it will be important to resolve the mechanisms that control how the size of the AZ and how the amounts of individual CAZ proteins in the different fractions are adapted in response to specific signals as well as if these mechanisms differ at synapses with diverging properties.

The abundance of a protein also depends on the level of its synthesis versus its degradation. Therefore, it is also important to consider how the exchange dynamics of the CAZ proteins relate to their metabolic turnover rates. This aspect was addressed both in cultured neurons and in live adult mice using state-of-the-art metabolic labeling methods combined with quantitative mass spectrometry. A systematic analysis of the turnover rates of synaptic proteins revealed half-life times in cultured neurons to be between 2 and 5 days [39]. Half-life times were three to four fold slower in brains of adult mice [40]. These results indicate that the synaptic dynamics of AZ components are rather dominated by their exchange rates and not by synthesis and degradation.

The ability of synapses to modify their functional characteristics in response to relevant physiological stimuli is referred to as synaptic plasticity. Presynaptic strength, the probability of SV fusion Pr (release probability) and concomitantly AZ size, have been shown to be regulated during homeostatic and Hebbian (experience-induced) synaptic plasticity (reviewed in [4], [41], [42]). Already a decade ago it was reported that prolonged silencing of global network activity is associated with an increase in the synaptic release probability [43]. Since then the results of numerous studies, in different preparations and using various manipulations of neuronal activity, have led to the concept that presynaptic efficacy is inversely correlated to the network activity that the synapse experiences; it is augmented by a decrease and reduced by an increase in overall network activity (reviewed in [42]). Furthermore, also Hebbian plasticity has been shown to involve presynaptic structural and functional changes (reviewed in [41]). Even though at most synapses in the brain the change in synaptic strength during long-term plasticity (LTP) is due primarily to postsynaptic alterations in receptor trafficking, several synapses also express presynaptic forms of LTP. For example the mossy fiber synapses in the hippocampus and the cerebellar parallel fiber synapses manifest robust sustained potentiation in response to repetitive presynaptic stimulation (reviewed in [44]). In addition to changes in presynaptic strength also structural alterations were observed after the induction of LTP [45].

The presynaptic response to network activity also involves changes in the composition of the AZ. For example, rapidly induced presynaptic strengthening by blocking glutamate receptors at the NMJ of Drosophila with Philanthotoxin resulted in the fast remodeling of the presynaptic cytomatrix at the AZ as an enlargement of the AZ was accompanied by an increase in the level of Bruchpilot and in the size of the readily releasable pool [46]. Changes in AZ composition were also detected in mammalian synapses in response to alterations of synaptic activity, as the total levels of the CAZ core components RIM, Munc13, Bassoon, Piccolo, ELKS and Liprin-α were reduced after global network silencing of cultured neurons [24], [47] and the levels of RIM1 and Munc13-1 were decreased after depolarization-induced presynaptic muting [48]. Taken together, this indicates that changes in AZ composition allow individual synapses to regulate their strength (Fig. 2). However, it is still unresolved how the modulation of AZ protein composition is regulated in response to changes in activity.

Accumulating evidence points to the degradation of CAZ proteins by the ubiquitin-proteasome system (UPS) as an important mechanism in this process. For instance, the downregulation of Bassoon and Liprin-α after prolonged silencing of global network activity was dependent on the activity of the UPS [24], [47]. In addition it was shown that presynaptic silencing/muting, the reduction of release probability after strong depolarization, requires a decrease in the levels of RIM1 and Munc13-1, which could be inhibited by blocking the UPS with MG-132 [48]. Accordingly, both presynaptic muting and the decrease in Munc13-1 levels could be prevented by overexpression of RIM1 [48] supporting a role for RIM1 in maintaining synaptic Munc13-1 levels [49] and in the induction of presynaptic muting [48]. Interestingly, the induction of presynaptic muting and the depolarization-induced decrease in RIM1 and Munc13-1 protein levels could also be inhibited by inducing cyclic-AMP (cAMP) signaling in cultured neurons with forskolin. Further support for a direct link between synaptic RIM levels and the activity of a individual synapse came from the observation that the amount of RIM at a synapse correlates with the synapse׳s activity [47], [48]. Based on these results it was proposed RIM levels determine the synaptic efficacy of an individual synapse and that RIM turnover is regulated by an interplay between degradation and phosphorylation, whereby phosphorylation would protect RIM from degradation. First evidence that degradation of AZ components can be controlled locally by protein interactions of AZ components comes from studies on the E3 ubiquitin ligase Shia-1, which was shown to be negatively regulated by Bassoon and Piccolo [50]. Shia-1 binds to Bassoon and Piccolo and loss of the two proteins promotes excessive ubiquitination and degradation of many presynaptic proteins, among them RIM and Munc13-1. Also the activity-induced changes in the levels of Liprin-α2 were found to be UPS-dependent and inhibition of the proteasome furthermore caused a shift in the mobile and immobile fractions of GFP-Liprin-α2 [24]. As Liprin-α2 levels in turn affect the association dynamics of RIM1 with the AZ the processes regulating Liprin-α2 turnover could also promote changes in synaptic strength.

Blocking network activity results in a compensatory increase in action-potential evoked Ca2+ entry into the presynaptic nerve terminal [51], which could in part be caused by the concurrent increase in the number voltage-gated P/Q type Ca2+-channels [42]. Considering that activity blockade also augments synaptic RIM levels [47] and that RIM plays a role in the localization of VGCCs to the AZ [21], [22] it was proposed that RIM contributes to homeostatic adaptations by recruiting VGCCs and components of the release machinery to AZ and by modulating vesicle priming. Interestingly, it was recently shown that the post-translational modification of RIM1α by SUMOylation is critical for the initial fast phase of SV exocytosis [52]. SUMOylation of RIM1α affects its interaction with Cav2.1 Ca2+-channels and thereby determines the clustering of VGCCs at the AZ. Hence, modulating the amount of SUMOylated RIM1α might represent a further mechanism to control presynaptic efficacy.

A further post-translational modification to which a role in synaptic plasticity has been attributed is phosphorylation. For example, a critical role for Cdk5 activity in presynaptic adaptation of hippocampal CA3 recurrent circuitry in response to chronic inactivity has been reported [53]. It was furthermore shown that binding of Cdk5 to the VGCC Cav2.2 causes an increase in current density as well as channel open probability, and affects the interaction of Cav2.2 with RIM1 and thereby SV release [54]. Protein kinase A (PKA) has long been known to be critically involved in the expression of presynaptic LTP (reviewed in [44]). As RIM1α is both important for most types of presynaptic LTP and is a PKA substrate it had been proposed that RIM1α phosphorylation by PKA on Serine 413 is necessary for presynaptic LTP, however analyses of knock-in mice (S413A) did not support this model (reviewed in [3], [44]). Therefore, PKA targets in presynaptic plasticity still remain to be identified. Recently, protein kinase C (PKC) was found to control the activity-induced dispersion from and reclustering to the AZ of the essential component of the release machinery Munc18-1 [55]. As synaptic Munc18-1 levels correlate with synaptic strength the PKC-dependent dynamic control of synaptic Munc18-1 levels represents a mechanism for individual synapses to modulate their output in response to ongoing activity. Further studies will be required to understand the mechanisms mediating the role of the AZ in synaptic plasticity in more detail.

Section snippets

Conclusions

Technological advances in recent years have greatly improved our understanding and view of AZ composition, ultrastructure and function. Proteomics approaches have identified the global AZ protein complement and based on progress in resolving the AZ ultrastructure basic principles of the structural organization of the protein meshwork at the AZ and the SV release machinery have emerged. Furthermore, first steps have been made to correlate ultrastructural elements to specific molecules and to

Acknowledgments

Work in the authors׳ laboratory is supported by the Deutsche Forschungsgemeinschaft, Germany (DFG, SFB1089), the German Ministry of Research and Education (BMBF, 01GQ0806), the European Union׳s Seventh Framework Programme (FP7/2007–2013) under Grant agreement no.602102 (EPITARGET), and local funding (BONFOR). We would like to thank Ege Kavalali, Noam Ziv and Dirk Dietrich for helpful discussions and critical reading of the manuscript.

References (55)

  • Y. Nakamura et al.

    Nanoscale distribution of presynaptic Ca2+ channels and its impact on vesicular release during development

    Neuron

    (2015)
  • V.N. Murthy et al.

    Inactivity produces increases in neurotransmitter release and synapse size

    Neuron

    (2001)
  • F. Girach et al.

    RIM1α SUMOylation is required for fast synaptic vesicle exocytosis

    Cell Rep.

    (2013)
  • S.C. Su et al.

    Regulation of N-type voltage-gated calcium channels and presynaptic function by cyclin-dependent kinase 5

    Neuron

    (2012)
  • J.J.E. Chua

    Macromolecular complexes at active zones: integrated nano-machineries for neurotransmitter release

    Cell. Mol. Life Sci.

    (2014)
  • M. Morciano et al.

    The proteome of the presynaptic active zone: from docked synaptic vesicles to adhesion molecules and maxi-channels

    J. Neurochem.

    (2009)
  • M. Sheng et al.

    The postsynaptic organization of synapses

    Cold Spring Harb. Perspect. Biol.

    (2011)
  • J.-M. Fritschy et al.

    Molecular and functional heterogeneity of GABAergic synapses

    Cell. Mol. Life Sci.

    (2012)
  • N.A. O’Rourke et al.

    Deep molecular diversity of mammalian synapses: why it matters and how to measure it

    Nat. Rev. Neurosci.

    (2012)
  • C. Biesemann et al.

    Proteomic screening of glutamatergic mouse brain synaptosomes isolated by fluorescence activated sorting

    EMBO J.

    (2014)
  • B.G. Wilhelm et al.

    Composition of isolated synaptic boutons reveals the amounts of vesicle trafficking proteins

    Science

    (2014)
  • K. Pfenninger et al.

    The fine structure of freeze-fractured presynaptic membranes

    J. Neurocytol.

    (1972)
  • R. Fernandez-Busnadiego et al.

    Quantitative analysis of the native presynaptic cytomatrix by cryoelectron tomography

    J. Cell Biol.

    (2010)
  • C. Limbach et al.

    Molecular in situ topology of Aczonin/Piccolo and associated proteins at the mammalian neurotransmitter release site

    Proc. Natl. Acad. Sci. USA

    (2011)
  • T. Matkovic et al.

    The Bruchpilot cytomatrix determines the size of the readily releasable pool of synaptic vesicles

    J. Cell Biol.

    (2013)
  • K.S.Y. Liu et al.

    RIM-binding protein, a central part of the active zone, is essential for neurotransmitter release

    Science

    (2011)
  • W. Fouquet et al.

    Maturation of active zone assembly by Drosophila Bruchpilot

    J. Cell Biol.

    (2009)
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