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RAPID REPORT

Revisiting synaptic vesicle pool localization in the Drosophila neuromuscular junction Annette Denker1,2 , Katharina Kr¨ohnert1 and Silvio O. Rizzoli1 1 2

European Neuroscience Institute, Grisebachstr. 5, G¨ottingen 37077, Germany International Max Planck Research School Molecular Biology, G¨ottingen, Germany

The synaptic vesicles are organized in distinct populations or ‘pools’: the readily releasable pool (the first vesicles released upon stimulation), the recycling pool (which maintains release under moderate stimulation) and the reserve pool (which is called into action only upon strong, often unphysiological stimulation). A major question in the field is whether the pools consist of biochemically different vesicles or whether the pool tag is a spatial one (with the recycling vesicles found next to the release sites, and the reserve ones farther away). A strong and stable spatial segregation has been proposed in the last decade in the Drosophila larval neuromuscular junction – albeit based solely on light microscopy experiments. We have tested here this hypothesis using electron microscopy (EM) photoconversion. We found the recycling and reserve pools to be thoroughly intermixed at the EM level, indicating that spatial location is irrelevant for the functional properties of the vesicle. (Received 13 February 2009; accepted after revision 22 April 2009; first published online 29 April 2009) Corresponding author S. O. Rizzoli: European Neuroscience Institute, Grisebachstr. 5, G¨ottingen 37077, Germany. Email: [email protected] Abbreviation NMJ, neuromuscular junction; RRP, readily releasable pool.

Synaptic vesicle recycling is arguably the best-described membrane trafficking event (see for example S¨udhof, 2004). However, the apparent simplicity of the vesicle cluster is deceiving: while the vesicles are similar on the ultrastructural level, they have different functional properties. They are currently separated into three main populations: the readily releasable pool (RRP), the recycling pool and the reserve pool (Rizzoli & Betz, 2005). The RRP is defined as the synaptic vesicles that are immediately available for release upon neural stimulation. These vesicles are generally thought to be docked to the presynaptic active zone and primed for release. The recycling pool is the pool of vesicles that maintain release on moderate stimulation; physiological frequencies of stimulation cause it to recycle continuously, and it is refilled by newly recycled vesicles – not by recruitment from the reserve pool. Finally, the reserve pool is a depot of synaptic vesicles from which release is triggered only during intense stimulation. These vesicles constitute the majority of vesicles in most presynaptic terminals, and it is possible that they are seldom or never recruited during physiological activity. The RRP constitutes only a small fraction (a sub-pool) of the recycling pool – those recycling pool vesicles which happen to be next to the release sites when an  C 2009 The Authors. Journal compilation  C 2009 The Physiological Society

action potential arises. The recycling and reserve pools, though, are stable entities, with little functional intermixing. What distinguishes these vesicles? It has often been proposed that they are spatially segregated – an intuitive model which, however, has not been confirmed by vesicle labelling experiments in many preparations (frog NMJ, Rizzoli & Betz, 2004; snake NMJ, Teng & Wilkinson, 2000; mammalian hippocampal synapses, Harata et al. 2001; mammalian calyx of Held, de Lange et al. 2003; goldfish bipolar nerve terminals, Paillart et al. 2003). However, the two pools of vesicles are still thought to be spatially separated in one major preparation, which is widely used for studies of synaptic activity – the larval Drosophila neuromuscular junction. Kidokoro, Kuromi and collaborators (Kuromi & Kidokoro, 1998, 1999, 2000, 2002; Kuromi et al. 2004; review by Kuromi & Kidokoro, 2003) have established, using fluorescence imaging, that the exo/endo cycling pool (the equivalent of the recycling pool), and the reserve pool are morphologically separated. The exo/endo cycling pool is proposed to be located at the periphery of the synaptic boutons, with the reserve pool found in the bouton centre. This, however, conflicts with the known ultrastructure of the Drosophila NMJ, where the bouton centres are largely devoid of vesicles (see for example Atwood et al. 1993); also, immunostaining for a DOI: 10.1113/jphysiol.2009.170985

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synaptic vesicle marker revealed no vesicles in the bouton centre even for the Kuromi and Kidokoro group (Kuromi et al. 2004). We decided to test this vesicle localization hypothesis by taking advantage of the high resolution of electron microscopy, rather than relying solely on light microscopy. We surprisingly found that the two Drosophila vesicle pools are intermixed, in contrast to the previous findings, but in agreement with the general understanding of vesicle recycling in most other preparations. Methods Materials

Chemicals were purchased from VWR (Hannover, Germany) or Sigma (Taufkirchen, Germany), unless stated otherwise. FM 1-43 (SynaptoGreen) was purchased from Biotrend (K¨oln, Germany). The following wild-type fly stocks were used: w1 , w1118 . The flies were raised at 21◦ C. Buffers

The standard Drosophila medium contained 130 mM NaCl, 36 mM sucrose, 5 mM KCl, 2 mM CaCl 2 , 2 mM MgCl 2 , 5 mM Hepes, pH 7.3 (Jan & Jan, 1976; Kuromi & Kidokoro, 1999). High potassium medium contained 90 mM KCl (with a corresponding reduction in NaCl). Cyclosporine A was used at a final concentration of 30 μM. In cyclosporine A experiments, a pre-incubation of 20 min was used before high potassium stimulation. Dye loading

The preparations were rapidly dissected according to previous procedures (Jan & Jan, 1976). Images were obtained from body wall muscles 6 and 7, from the fourth and fifth abdominal segments. For FM staining experiments, the preparations were bathed in FM 1-43-containing high potassium medium (10 μM) for 5 min. The same procedure was used for cyclosporine A pre-treated preparations. For electrical stimulation, the larvae were mounted in a chamber provided with a platinum plate electrode (custom made in the workshop of the Max Planck Institute for Biophysical Chemistry, G¨ottingen, Germany). The electrode plates were 8 mm apart; 100 A shocks were delivered using an A385 stimulus isolator and an A310 AccupulserTM stimulator (World Precision Instruments, Berlin, Germany), at the frequency mentioned in the text. To allow for recycling after the end of stimulation, tetanically stimulated preparations were allowed to recover in presence of the dye for 10 min. The preparations were then washed in Drosophila medium for 5–10 min and imaged. Destaining of FM 1-43 labelled preparations was performed by incubation for 5 min in

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high potassium medium in absence of the dye, before briefly washing with normal medium and imaging. Fluorescence imaging

We used an upright epifluorescence microscope (Axioskop 2 FS plus, Zeiss, G¨ottingen, Germany), equipped with a Zeiss 63× 1.0 NA water immersion objective lens, a 100 W Hg lamp (Zeiss), 25% neutral density transmission filters, excitation filter (HQ 470/40), dichroic mirror (495 DCLP), and emission filter (500 long pass, all from AHF, T¨ubingen, Germany). Image acquisition was performed using a Zeiss MRM camera. Image analysis was performed using software custom written in MATLAB (The Mathworks, Natick, MA, USA; Rizzoli & Betz, 2004). Photoconversion and electron microscopy

The procedure was performed essentially as described previously (Rizzoli & Betz, 2004). Briefly, the preparations were stained with FM 1-43 as above, fixed with 2.5% glutaraldehyde in phosphate-buffered saline (PBS) (subsequently quenched with 100 mM ammonium chloride), incubated with 1.5 mg ml−1 di-aminobenzidine, and illuminated using a 20× 0.5 NA objective from Olympus (Hamburg, Germany). The preparations were illuminated until a dark precipitate accumulated, and were then washed, treated with osmium tetroxide, and dehydrated using an ethanol dilution series. They were then embedded in Epon resin (Plano, Wetzlar, Germany), and processed for electron microscopy as described previously (Rizzoli & Betz, 2004). Sections of ∼100 nm thickness were analysed. Electron micrographs were acquired using a Zeiss EM 902A microscope, provided with a 1024 × 1024 CCD detector (Proscan CCD HSS 512/1024; Proscan Electronic Systems, Scheuring, Germany). Image analysis was performed using software custom written in MATLAB (Rizzoli & Betz, 2004). Results Fluorescence imaging of the vesicle pools

To investigate vesicle pool localization in the Drosophila neuromuscular junction, we first labelled the different pools with the styryl dye FM 1-43, as originally described by Kuromi & Kidokoro (1998, 1999, 2000). Styryl dye labelling (see review by Cochilla et al. 1999) is a time-tested technique in which the preparations are bathed in low concentrations of the dye during stimulation, allowing it to get trapped within recycling vesicles. After washing the dye, vesicle cluster positioning can easily be investigated by fluorescence microscopy. Kuromi and Kidokoro proposed  C 2009 The Authors. Journal compilation  C 2009 The Physiological Society

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that high potassium stimulation induces only the recycling of the peripherally located exo/endo cycling pool, which was indeed the case (Fig. 1A). Tetanic stimulation (30 Hz for 5 min) and cyclosporine A pre-treatment have been proposed to label both the exo/endo cycling and the reserve pool (Kuromi & Kidokoro, 1999, 2000). While the two treatments did cause a significantly stronger labelling of the synapses (Fig. 1C),

they did not increase substantially the proportion of boutons with labelled centres (Fig. 1E). Finally, the vesicles labelled by these treatments were quite able to be released upon a second step of high potassium stimulation in the absence of the dye (destaining; Fig. 1B and D) although one would not expect the reserve pool to release with high potassium treatment (Kuromi & Kidokoro, 1998). Moreover, the dye destaining should have left the bouton

Figure 1. Synaptic vesicle pools at the light microscopy level A, typical Drosophila synapses stained by stimulation in the presence of FM 1-43 (upper panels). The expected result (staining pattern) for the different staining conditions (filling of the periphery, or of both the centre and the periphery) is indicated by the cartoons. The images were auto-scaled, to give a good indication of the staining distribution. Insets show the same images, scaled identically; note the lower intensity of the high potassium stimulated preparations. Size bar = 10 μm. B, typical images of preparations stained as in A, and then destained via high potassium application. The expected staining pattern is indicated by the cartoons. C, quantification of the fluorescence intensity in the stained terminals. Bars show averages of at least three independent experiments (± S.E.M.). D, quantification of the fluorescence destaining. The bars show the average residual fluorescence after destaining (as percentage of the initial levels) in at least three independent experiments (± S.E.M.). E, quantification of the fraction of synaptic boutons which present ‘empty’ or ‘filled’ centres. The analysis was performed by determining the intensity of the bouton centres, and comparing it with that of the bouton periphery, in a semi-automated fashion (using a self-written routine in Matlab). Bars show averages of at least three independent experiments (± S.E.M.).  C 2009 The Authors. Journal compilation  C 2009 The Physiological Society

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centres (reserve pool) labelled – which was clearly not the case (Fig. 1B). Thus, our results only partially confirmed the previous findings – stronger stimulation does result in more labelling, occasionally in more filling of the bouton centres, although no separation of the two pools is evident. One should note also that the definition of the bouton centre is quite difficult to apply: even in the high potassium stained preparations (Fig. 1A and E), virtually all of the small boutons appeared to have their centres filled – likely to be due to the low resolution of light imaging missing the (even smaller) empty centre. Ultrastructural investigation of the vesicle pools

Fluorescence imaging is limited by the diffraction of light to an optimal resolution of ∼200 nm (Hell, 2007; this value actually increases substantially for the water-immersion objectives generally used for NMJ imaging, due to their relatively low numerical aperture). Therefore, we decided to employ transmission electron microscopy to investigate the vesicle pools. To be able to investigate the same conditions as described above, we employed a technique termed photoconversion (or photooxidation; Sandell & Masland, 1988; see details in Harata et al. 2001; Rizzoli & Betz, 2004, for styryl dye photoconversion in synaptic vesicles). The principle of the technique is straightforward: the fluorescently labelled preparations are fixed, incubated with di-amino-benzidine (DAB), and illuminated using a normal fluorescence set-up (see online supplemental material, Supplementary Fig. 1A). During illumination, free radicals generated by the dye molecules oxidize the DAB, which forms a stable, insoluble, electron-dense precipitate that has a dark appearance. DAB is oxidized only in the immediate vicinity of the fluorescent molecules, which allows the precipitate to form selectively within the FM-labelled synaptic vesicles. As no membrane permeabilization is involved, the DAB product remains trapped within the vesicle lumen. The reaction is relatively easy to follow (Supplementary Fig. 1B), with the preparations bleaching and accumulating the DAB oxidation product within ∼20–45 min. The preparations are then processed for electron microscopy. The labelled

vesicles are clearly identifiable (Supplementary Fig. 1C and D). We next applied the photoconversion technique to preparations in which we labelled either selectively the exo/endo cycling pool (by high potassium stimulation), or vesicles from both of the pools (tetanic stimulation, cyclosporine A pre-treatment). Typical electron micrographs are shown in Fig. 2A; clearly, the labelled and unlabelled vesicles appear intermixed in all of the conditions. To label selectively the reserve pool, we applied the stronger stimulation protocols, followed by releasing the exo/endo cycling pool via high potassium application in the absence of the dye (Fig. 2B; for comparison purposes, we also investigated preparations where the exo/endo cycling pool was stained and then destained via high potassium application). We quantified the distance of the labelled and unlabelled vesicles to the bouton periphery, as the exo/endo cycling pool vesicles are expected to be located closer to the periphery than the reserve pool (according to the pool model). This was not the case – the vesicle pools appeared intermixed in all conditions (Fig. 2C), even when only the reserve pool was labelled (Fig. 2D). Note that the slight shift towards the bouton centre in the cyclosporine A and tetanically treated preparations (Fig. 2C) is actually reversed after destaining the recycling pool via high potassium application (Fig. 2D) – which is the opposite of what one would expect based on the pool model (i.e. a filled centre, when the reserve pool alone is labelled). To verify whether the definition of bouton periphery was affected by the fact that we relied solely on single EM sections, we decided to analyse also three-dimensionally reconstructed boutons. The largely intermixed status of the pools was confirmed for all conditions (Fig. 3; see also Supplementary Movies 1–3). Bouton centre occupancy

While our quantification of electron microscopy results suggests that the two vesicle pools are largely intermixed, it does not deal specifically with the presence of vesicles (or fluorescent material) in the bouton centres. As indicated by the images presented above (and by the supplementary

Figure 2. Synaptic vesicle pools at the electron microscopy level A, typical images of FM 1-43 labelled and photoconverted nerve terminals, in the different staining conditions. Size bar = 500 nm. The cartoons indicate the expected FM 1-43 staining pattern of the preparations. The insets show examples of labelled and unlabelled vesicles from the respective images (the original position of the vesicles is indicated by the arrowheads). Note that, although the contrast and/or staining intensity differs between the preparations, the labelled and unlabelled vesicles are easily distinguishable. B, the preparations were stained as in A and subsequently destained via high potassium application in the absence of the dye, followed by fixation and photoconversion. The expected staining pattern is indicated by the cartoons. C–D, quantification of the distance from the bouton periphery (defined as the outline of the plasma membrane in the particular micrograph) for the labelled and unlabelled vesicles, in the stained (C) or stained and subsequently destained (D) preparations. The graphs show averages of at least three independent preparations (± S.E.M.).  C 2009 The Authors. Journal compilation  C 2009 The Physiological Society

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movies), large synaptic boutons generally had a centre almost completely devoid of vesicles, while this empty centre was more difficult, or impossible, to observe in the smaller boutons, irrespective of the treatment – an observation which agrees well with the fluorescence data (Fig. 1). The cyclosporine A treated preparations also had a number of labelled vacuoles in the bouton centres (see Fig. 3A), which offers an explanation for the increase in the number of centre-labelled boutons in this condition (Fig. 1D). However, it could still be argued that extremely strong stimulation (which would label almost all vesicles) would also result in the labelling of the bouton centres. We thus checked for synapses where the large majority of the vesicles were labelled (Supplementary Fig. 2). The pattern was the same as before: the larger boutons showed clearly a vesicle-free centre, without any possibility for hosting the reserve pool.

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Discussion The results presented above offer a fairly different view of vesicle pool localization than that expected from the previous styryl dye imaging experiments: the two pools are intermixed, and pool status clearly does not depend upon distance from the bouton periphery. The occupancy of the centre of the boutons (the main argument in favour of a specially localized reserve pool) may be due to the accumulation of recycling intermediates (vacuolar structures), and of a number of vesicles when extremely strong stimulation procedures are used (Fig. 2C). However, this is clearly only a short-lived phenotype (Fig. 2D), with the labelled vesicles being later released or transported to the bouton periphery. While Kuromi & Kidokoro (1998) also used the dynamin mutant shibire ts to label the bouton centres, this observation is again likely to be due to problems concerning recycling

Figure 3. 3D reconstructions of labelled preparations A, exemplary three-dimensional reconstructions of FM 1-43 labelled and photoconverted preparations. Plasma membrane is shown in yellow; active zones are shown in red; labelled synaptic vesicles and vacuoles are shown in purple; unlabelled synaptic vesicles and vacuoles are shown in white. Note mixing of the labelled and unlabelled vesicles. Also, note the vesicle-free centre in the cross-section view of the tetanic stimulation reconstruction. Arrow points to a region where a number of large (labelled and unlabelled) vacuoles are present. Size bar = 500 nm. See Supplementary Movies 1–3 for a more detailed view. The cartoons indicate the expected FM 1-43 staining pattern of the preparations. B, quantification of the vesicle distances from the terminal periphery for the different preparations from A).  C 2009 The Authors. Journal compilation  C 2009 The Physiological Society

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of membrane intermediates, rather than to a special vesicle pool. Although beyond the scope of our study (to investigate wild-type vesicle pools in Drosophila), we did test shibire ts mutants in conditions similar to those published previously, and still found numerous vesicle-free bouton centres, and intermixed labelled and unlabelled vesicles (data not shown). The results connect the Drosophila larval NMJ with the general view of synaptic pools, which predicts that the two different types of vesicles (reserve and recycling) are intermixed, but still maintain their different properties (Rizzoli & Betz, 2005). This is certainly a somewhat counter-intuitive result, as the different vesicles do not look different in any morphological correlate – it is actually their homogenous morphology which has allowed the purification of vesicles via size-exclusion columns (Takamori et al. 2006). Interestingly, the vesicle purification study revealed that while the vesicles contain a high copy number of certain essential proteins, such as the fusion protein synaptobrevin or the (proposed) calcium sensor synaptotagmin, they also contain (on average) a small number of other proteins, such as endosomal markers, as well as a high variety of small Rab GTP-ases (Takamori et al. 2006). It seems impossible to suggest a vesicular function for only a handful of copies per vesicle of, for example, the endosomal SNARE fusion proteins (Takamori et al. 2006). However, if these proteins are found only on a fraction (pool) of vesicles (Rizzoli et al. 2006), it would imply that they are present in a substantial copy number per vesicle, thereby allowing these vesicles to function in a different fashion from the others (i.e. allowing them to recycle through endosomal sorting). This would be all the more the case for the Rab proteins, which are well-known membrane organizers (Zerial & McBride, 2001), understood to impart different functions to the organelles (or membrane domains) which they bind to. The maintenance of two functionally separate but spatially intermixed pools of vesicles would not be difficult if after exocytosis the vesicle molecules remain in clusters accessible to the (clathrin) sorting machinery (Willig et al. 2006). Sorting of each individual vesicle, without mixing with neighbouring ones, could then easily be performed at the plasma membrane, or could even involve a further step of endosomal recycling (see review by Jung & Haucke, 2007). Additionally, the fact that different synaptic vesicle pools appear to use distinct recycling routes (Rizzoli & Betz, 2005), even in the Drosophila NMJ (Koenig & Ikeda, 1996), would be an even stronger argument to the fact that the vesicles are biochemically separated, and can therefore be recycled independently, without a need for spatial segregation. Furthermore, our results are in good agreement with several recent studies of vesicle movement, which indicate that (newly labelled) recycling vesicles are fairly mobile, dispersing from synapse to synapse (Darcy et al. 2006;  C 2009 The Authors. Journal compilation  C 2009 The Physiological Society

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Westphal et al. 2008), or moving at relatively high rate within the synapses (Gaffield & Betz, 2007; Westphal et al. 2008). Finally, a study relying on a genetically encoded vesicle marker demonstrated that the reserve vesicles are also mobile, being transported between synapses in a similar fashion to the recycling ones (Fernandez-Alfonso & Ryan, 2008), underlining the conclusion that vesicle pool spatial segregation would be difficult if not impossible to achieve under such conditions.

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Author contributions A.D. conceived, designed and performed experiments; analysed and interpreted data; drafted the manuscript; and approved the final version of the manuscript. K.K. designed and performed experiments; analysed data; revised the manuscript critically for important intellectual content; and approved the final version of the manuscript. S.O.R. conceived, designed and performed experiments; analysed and interpreted data; drafted the manuscript; and approved the final version of the manuscript. All experiments were performed at the European Neuroscience Institute, Grisebachstr. 5, G¨ottingen, 37077, Germany. Acknowledgements We thank Christina Sch¨afer for technical assistance. We thank Dr Carolin Wichmann for help with Drosophila maintenance and general Drosophila procedures. The work was supported by the DFG Research Center for Molecular Physiology of the Brain (CMPB)/Excellence Cluster 171.

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