The authors thank Dr. Mike Ehlers and his laboratory members for originally demonstrating the protocol for subcellular fractionation of synaptic plasma membranes. We also thank Wendy Horsfall for management of the mouse colonies. This work was supported by operating grants from CIHR (AJR and AS).

Le protocole suivant est conforme aux lignes directrices du Conseil canadien de protection des animaux et a été approuvé par la Faculté de Pharmacie Comité de protection des animaux et la médecine à l&#39;Université de Toronto. 1 Préparer requis réactifs comme indiqué dans le tableau 1 <ol><li> Ajouter protease et de phosphatase (si nécessaire) des inhibiteurs de l&#39;ensemble des solutions de saccharose, des tampons et le trou DDH 2 O à des concentrations indiquées dans le tableau 1. IMPORTANT: Effectuez toutes les étapes à 4 ° C et pré-cool tous les réactifs et le matériel avant le début de l&#39;expérience. Étiqueter tous les tubes, y compris les tubes d&#39;ultracentrifugation, avec un marqueur permanent. </li></ol> 2 Dissection de la région du cerveau appropriée <ol><li> Sacrifier l&#39;animal par dislocation cervicale ou décapitation. Maison et euthanasier les animaux conformément aux politiques institutionnelles et gouvernementales sur les soins aux animaux. </li><li> Retirer rapidement les ee cerveau du crâne et les placer sur un bloc de dissection froide. </li><li> Disséquer la région du cerveau appropriée de tissus frais et passez à l&#39;étape 3. </li></ol> 3. homogénéisation des tissus entraînés par un moteur de verre Téflon Homogenizer <ol><li> Placer 50 à 100 mg (pour SPM), ou 100 à 200 mg (pour PSD) du tissu découpé dans un tube de 13 ml en polypropylene marqué contenant 4 ml de solution 0,32 M de saccharose tamponné à l&#39;HEPES. Transférer l&#39;échantillon dans un 15 ml conique tissu de verre Téflon broyeur / homogénéisateur, réglez la commande de moteur à 900 tours par minute (réglage 7) et homogénéiser l&#39;échantillon avec 12 coups sur une période de 30 secondes. Utilisez un autre verre-téflon homogénéisation entre les échantillons, ou rincer l&#39;homogénéisation avec de l&#39;eau distillée froide entre les échantillons et essuyez avec un Kimwipe. NOTE: jusqu&#39;à 4 g de tissu peut être homogénéisé dans 4 ml de sucrose 0,32 M de solution tampon HEPES, cependant il est préférable d&#39;utiliser moins de tissu pour assurer un fractionnement convenable. </li><li> Transférer l&#39;échantillon homogénéisé dos to le même tube de 13 ml en polypropylene. Réserve 100 pi d&#39;homogénat et conserver à -80 ° C pour la quantification de protéines et d&#39;une analyse western blot de la fraction protéique total. </li></ol> 4. basse vitesse de centrifugation pour éliminer la fraction nucléaire (rendements surnageant S1) <ol><li> Centrifugeuse le broyat dans un rotor à angle fixe à 900 g pendant 10 min à 4 ° C. Transférer le surnageant (S1) pour une nouvelle, marquée 13 ml tube et remettre en suspension le culot nucléaire fraction (P1) dans 500 ul de 0,32 M saccharose tamponné à l&#39;HEPES. NOTE: Le (P1) la fraction peut être conservé à -80 ° C et utilisé pour l&#39;analyse western blot de la fraction nucléaire. </li></ol> 5. enrichissement de la fraction brute Synaptosomal (rendements Pellet P2) <ol><li> Centrifuger le surnageant (S1) à 10 000 g pendant 15 min à 4 ° C. Eliminer le surnageant (S2) et réserve de 500 pl dans un tube à centrifuger de stocker à -80 ° C pour la suite quant protéinefication et l&#39;analyse western blot de l&#39;/ fraction membranaire lumière cytosolique. Remettre en suspension le reste de la fraction synaptosomale brute pastille (P2) dans 1 ml de solution de saccharose 0,32 M tamponnée par HEPES, puis ajouter une autre solution de 3 ml de saccharose 0,32 M tamponné à l&#39;HEPES. </li><li> Centrifugeuse à 10.000 x g pendant 15 min à 4 ° C. Eliminer le surnageant (S2 &#39;) et de la réserve de 500 pl dans un tube à centrifuger pour stocker à -80 ° C pour la quantification de protéines et d&#39;une analyse western blot de l&#39;/ fraction membranaire lumière cytosolique. Enregistrez le synaptosomale brut culot lavé (P2 &#39;) dans le tube en polypropylène. </li></ol> 6. de lyse (hypoosmotiques choc) de la Fraction brut Synaptosomal <ol><li> Lyser les synaptosomes bruts pastille (P2 &#39;) dans le tube en polypropylène par remise en suspension dans 1 ml de trou DDH 2 O. Ajouter un autre 3 ml de ddH 2 O, transférer à une homogénéisation des tissus de verre Téflon, et homogénéiser à la main avec 3 coups. NOTE: Il est importantd&#39;effectuer cette étape aussi rapidement que possible et de procéder rapidement à l&#39;étape 6.2. </li><li> échantillons de transfert de l&#39;homogénéisation de nouveau dans le même 13 ml tube en polypropylène et rapidement ajuster l&#39;échantillon revenir à 4 mM HEPES avec 16 pi de 1 M solution HEPES, renverser pour mélanger. </li><li> Faire tourner les échantillons à 4 ° C pendant 30 min pour assurer la lyse complète. </li></ol> 7. enrichissement de la fraction Synaptosomal membrane (P3) <ol><li> Centrifugeuse à 25 000 x g pendant 20 min à 4 ° C. Retirer le surnageant de la fraction vésiculaire brut (S3) et réserver dans un tube de 5 ml pour stocker à -80 ° C, pour la quantification de protéines et analyse subséquente western blot. Remettre en suspension de la fraction membranaire de synaptosomes (P3) dans 1 ml de solution de saccharose à 0,32 M tamponné à l&#39;HEPES. </li></ol> 8 Préparation du gradient de saccharose discontinu <ol><li> Pipeter exactement 3,5 ml de solution de saccharose 1,2 M tamponné à l&#39;HEPES et 3,0 ml chacune exactement de 1,0 M et 0,8 M HEPESolution de saccharose à S-tampon dans séparées 6 ml accrochage des tubes à bouchon. NOTE: Ceci est fait pour pré-mesurer les volumes des tampons qui sont utilisés pour faire le gradient de saccharose. Mesure exacte du volume est important de veiller à ce que les gradients résultants seront équilibrées par ultracentrifugation. </li><li> En utilisant une pipette Pasteur en verre et ampoule, transférer 1,2 M solution de tampon HEPES saccharose à 12 ml polyallomère tube d&#39;ultracentrifugation. Superposer soigneusement la solution de saccharose 1,0 M HEPES tamponné au-dessus de la solution de saccharose 1,2 M tamponné à l&#39;HEPES. Enfin, la couche de la solution de saccharose 0,8 M HEPES tamponné au-dessus de la solution de saccharose 1,0 M tamponné à l&#39;HEPES. Utilisez une pipette Pasteur frais pour chaque solution de saccharose et de prendre soin de ne pas perturber le gradient de saccharose. Vibrations d&#39;un tourbillon de banc ou centrifuger viendra troubler l&#39;intégrité du gradient. </li><li> En utilisant une pipette Pasteur, la couche de la fraction membranaire de synaptosomes remis en suspension (P3) sur le dessus du gradient discontinu de saccharose préparé.Veiller à ce que les gradients sont équilibrés en les pesant à l&#39;intérieur du seau du rotor à godets oscillants. Si les compartiments ne sont pas équilibrées, utiliser une pipette P200 à ajouter soigneusement saccharose 0,32 M tamponnée par HEPES à la partie supérieure du gradient et d&#39;équilibrer les godets de rotor. </li></ol> 9 Le fractionnement de la membrane plasmique synaptique (SPM) <ol><li> Ultracentrifugation dans un seau rotor oscillant à 150 000 g pendant 2 h à 4 ° C. Retirez délicatement les gradients de saccharose des seaux d&#39;ultracentrifugation. En utilisant une aiguille de 18 G et une seringue de 1 ml, la perforation du tube au fond de la solution de saccharose 1,0 M interphase / 1,2 M de tampon HEPES et retirer la couche de la membrane plasmique synaptique (SPM). </li><li> Prenez note du volume que chaque échantillon de SPM occupe dans la seringue de 1 ml. Passer la couche de SPM recueillies dans 3,5 ml d&#39;épaisseur de paroi des tubes d&#39;ultracentrifugation, ajouter exactement 2,5 volumes de 4 mM de HEPES à ajuster la concentration en saccharose de 1,2 M à 0,32 M. Une fois que chaque échantillon a été ajustered à 0,32 M de saccharose, équilibrer les tubes avec une solution de saccharose 0,32 M tampon HEPES. </li><li> Ultracentrifugation dans un rotor à angle fixe à 200 000 g pendant 30 min à 4 ° C. Retirer et jeter le surnageant. Remettre en suspension le culot de membranes synaptiques de plasma (SPM) dans 300 pi de solution de HEPES 50 mM / EDTA 2 mM. Conserver les échantillons à -80 ° C jusqu&#39;à ce que le transfert de Western. </li></ol> 10 Préparation de la fraction de la densité post-synaptique (PSD) <ol><li> Décongeler les échantillons SPM sur la glace. </li><li> Ajouter une solution de 0,54% de Triton X-100 dans une solution de HEPES 50 mM / EDTA 2 mM. </li><li> Dans un tube de 5 ml en polystyrène, mélanger 2,7 ml de Triton X-100 / solution HEPES / EDTA et 300 pi de la fraction de SPM et faire tourner l&#39;échantillon pendant 15 min à 4 ° C </li><li> échantillons de transfert à 3,5 ml tubes d&#39;ultracentrifugation à paroi épaisse et centrifuger les échantillons à 32 000 g pendant 20 min à 4 ° C. </li><li> Réservez le surnageant (Triton X-100 fraction soluble [TS]) et conserver à -80; ° C. NOTE: A ce stade, le culot représente le &quot;PSD-1T&quot; fraction. Un second traitement avec un détergent va produire un &quot;PSD-2T&quot; fraction. Si une fraction PSD-1T est souhaité, passez à l&#39;étape 10.6. <ol><li> Pour produire une fraction PSD-2T, remettre en suspension le culot dans 3 ml de 0,5% de Triton X-100 dans 50 mM de HEPES / 2 mM d&#39;EDTA solution. Tournez échantillons pendant 15 min à 4 ° C. Centrifuger les échantillons à 32 000 g pendant 20 min à 4 ° C et passez à l&#39;étape 10.6. </li></ol></li><li> Jeter le surnageant et remettre en suspension le post-synaptique (PSD) culot dans une solution à 50 mM HEPES / EDTA 2. REMARQUE: Le volume de remise en suspension peut dépendre de la taille de la pastille; un volume de 50-75 ul est recommandé. Étant donné que la fraction est une fraction PSD détergent insoluble, il est souvent nécessaire d&#39;ajouter de 1 à 2 pi de SDS à 0,5% pour permettre la remise en suspension de la protéine complète. Conserver les échantillons à -80 ° C jusqu&#39;à ce que le transfert de Western. </li></ol>

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E., Chammas, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+lysosomal-associated+membrane+protein+LAMP-1+is+a+novel+differentiation+marker+for+HC11+mouse+mammary+epithelial+cells.">The lysosomal-associated membrane protein LAMP-1 is a novel differentiation marker for HC11 mouse mammary epithelial cells.</a> Differentiation. 61, 113-120 (1996).</li><li>Otera, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peroxisomal+targeting+signal+receptor+Pex5p+interacts+with+cargoes+and+import+machinery+components+in+a+spatiotemporally+differentiated+manner:+conserved+Pex5p+WXXXF/Y+motifs+are+critical+for+matrix+protein+import.">Peroxisomal targeting signal receptor Pex5p interacts with cargoes and import machinery components in a spatiotemporally differentiated manner: conserved Pex5p WXXXF/Y motifs are critical for matrix protein import.</a> Mol Cell Biol. 22, 1639-1655 (2002).</li><li>Goubaeva, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+mitochondrial+connexin+43+regulates+apoptosis.">Cardiac mitochondrial connexin 43 regulates apoptosis.</a> Biochem Biophys Res Commun. 352, 97-103 (2007).</li><li>Lamers, K. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+S-100B,+neuron-specific+enolase+(NSE),+myelin+basic+protein+(MBP)+and+glial+fibrillary+acidic+protein+(GFAP)+in+cerebrospinal+fluid+(CSF)+and+blood+of+neurological+patients.">Protein S-100B, neuron-specific enolase (NSE), myelin basic protein (MBP) and glial fibrillary acidic protein (GFAP) in cerebrospinal fluid (CSF) and blood of neurological patients.</a> Brain Res Bull. 61, 261-264 (2003).</li><li>Arunachalam, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Munc18-1+is+critical+for+plasma+membrane+localization+of+syntaxin1+but+not+of+SNAP-25+in+PC12+cells.">Munc18-1 is critical for plasma membrane localization of syntaxin1 but not of SNAP-25 in PC12 cells.</a> Mol Biol Cell. 19, 722-734 (2008).</li><li>Brandstatter, J. H., Fletcher, E. L., Garner, C. C., Gundelfinger, E. D., Wassle, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+expression+of+the+presynaptic+cytomatrix+protein+bassoon+among+ribbon+synapses+in+the+mammalian+retina.">Differential expression of the presynaptic cytomatrix protein bassoon among ribbon synapses in the mammalian retina.</a> Eur J Neurosci. 11, 3683-3693 (1999).</li></ol>

The authors have no conflicts of interest to disclose.

Il ya plusieurs étapes de la procédure qui sont essentiels pour le succès. A l&#39;étape 3, il est important qu&#39;un certain degré de cohérence homogénéisation est obtenue pour chaque échantillon. Lorsque le tissu avec le moteur entraîné homogénéisateur homogénéisation, une vitesse constante est utilisée non seulement pour la rotation du pilon, mais également avec le nombre de coups. Le temps d&#39;incubation au cours du choc osmotique doit être précise, car homogénéisation prolongée ou incubatio...

Les neurones communiquent par des synapses, et la qualité de cette communication est régulé dans une large mesure par des modifications de la composition des protéines au niveau de la synapse. En particulier, les protéines qui se trouvent dans la densité post-synaptique des neurones participent à la communication par les récepteurs des neurotransmetteurs échafaudage intimement avec leurs systèmes de transduction de signaux 1. En outre, des modifications durables de la force de l&#39;efficacité synaptique sont contrôlés par l&#39;ajout ou le retrait de récepteurs à la densité post-synaptique 6.1. Par conséquent, l&#39;isolement et la quantification des protéines synaptiques est une technique nécessaire et utile pour mieux comprendre la façon dont les neurones répondent à des stimuli et modifier l&#39;efficacité synaptique 7. Cet article décrit une technique courante pour isoler les protéines synaptiques à partir de tissu cérébral des rongeurs par ultracentrifugation sur gradient de saccharose discontinu. La fraction de membrane synaptique de plasma peut être enrichie et isolées en se basant sursa densité en saccharose, qui a été déterminé de manière empirique pour être semblable à 1,2 M de saccharose. En fonction de la question biologique, des fractions subcellulaires peuvent être séparés par des gradients continus ou discontinus de Percoll ou saccharose soit. Gradients continus pour permettre la séparation de protéines en plusieurs fractions; cela peut être particulièrement utile pour démontrer la co-localisation des protéines dans une fraction donnée 8. Cependant, la préparation de gradients continus est plus laborieux et n&#39;est pas nécessaire pour de nombreuses applications. Gradients discontinus sont relativement plus faciles à préparer et peuvent être utilisées pour séparer les protéines en quelques fractions, généralement définies. Gradients discontinus qui sont composées de trois couches de saccharose d&#39;augmenter la molarité ont été largement utilisés pour isoler les protéines associées à la membrane plasmique synaptique (SPM). Cette fraction synaptique de la membrane plasmatique peut être encore traitée pour la FRACTIO densité post-synaptiquen (PSD) par traitement avec un détergent et de l&#39;isolement de la fraction insoluble dans le détergent. Lorsque ce procédé a été décrit pour la première dans les années 1960, 9,10, la microscopie électronique a été utilisé pour démontrer les organelles et de membranes qui définissent approximativement la membrane plasmique synaptique et les fractions de densité post-synaptiques 9-14. Ces études ont démontré l&#39;inclusion des membranes pré-et post-synaptiques et des vésicules synaptiques dans la fraction de SPM; après le traitement au détergent essentiellement la dense aux électrons, la densité post-synaptiques sont visibles. Dans la procédure, un choc hypotonique est utilisé pour pincer les processus synaptiques à partir du corps de la cellule 10. Cette étape prend avantage du fait que les mitochondries sont plus résistants aux chocs osmotiques et restent intacts, et ainsi ils sédimentent au fond du gradient de saccharose (figure 1). En utilisant la même technique d&#39;enrichissement, les SPM et PSD fractions ont d&#39;abord été biochimiquementdéfini par électrophorèse sur gel de polyacrylamide et la séquence des composantes majeures de protéines de 15 à 17. Par la suite l&#39;analyse par transfert de Western a été utilisée pour détecter et quantifier le niveau de protéines synaptiques et en outre définir ces fractions (Figure 2). Nous avons utilisé cette technique dans nos laboratoires pour quantifier les changements dans les niveaux synaptiques du transporteur de la dopamine qui se produisent lorsque le lieu SLC6A3 est dupliqué chez la souris 18. Nous avons également utilisé cette technique dans les souris déficientes en récepteurs de NMDA pour découvrir la réduction spécifique à la synapse en protéines qui font partie de la DISC1 interactome 19. Il est évident à partir de l&#39;analyse western blot que les fractions SPM contiennent synaptiques protéines de la membrane des vésicules, des marqueurs de endosome, les protéines mitochondriales, associées à la membrane enzymes synthétiques et des molécules de transduction de signaux, ainsi que des parties intégrantes de la densité post-synaptique et les membranes de plasma synaptiques 20-23. Efractions PSD VEN peuvent avoir une contamination par des protéines mitochondriales abondantes et il peut être nécessaire d&#39;effectuer une deuxième sédimentation de gradient ou des étapes supplémentaires de purification pour les 13 retirer. Récemment, la spectrométrie de masse quantitative a fourni une liste de plus de 100 protéines de la densité post-synaptique seul, ainsi qu&#39;une indication de l&#39;abondance relative de ces éléments 24,25.

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preparation of synaptic plasma membrane and postsynaptic density proteins using a discontinuous sucrose gradient

Techniques de fractionnement subcellulaire de neurones permettent la quantification de protéines que l&#39;on traite et à partir de la synapse. Comme décrit à l&#39;origine à la fin des années 1960, des protéines associées à la membrane plasmique synaptique peuvent être isolées par ultracentrifugation sur un gradient de densité de saccharose. Une fois les membranes synaptiques sont isolés, le complexe macromoléculaire appelé la densité post-synaptique peut être ensuite isolé en raison de son insolubilité de détergent. Les techniques utilisées pour isoler les membranes plasmiques synaptiques et les protéines de la densité post-synaptiques demeurent essentiellement les mêmes après 40 ans, et sont largement utilisés dans la recherche en neurosciences en cours. Cet article décrit le fractionnement de protéines associées à la membrane plasmique synaptique et la densité post-synaptique en utilisant un gradient discontinu de saccharose. Résultant des préparations de protéines sont adaptés pour western blot ou analyse 2D DIGE.

La préparation du gradient de densité de saccharose devrait se traduire par une séparation claire des trois solutions molaires de saccharose (0,8, 1,0, et 1,2 M de saccharose). Voir la figure 3A un exemple de la pente avant de l&#39;échantillon de protéine est ajoutée. Si le gradient est préparé trop à l&#39;avance, ou si elle est préparée sur une surface de banc avec la vibration d&#39;un autre appareil, le gradient sera compromise et la séparation adéquate ne sera pas atteint. Si une sép...

Watch this Scientific Journal Video about Preparation of Synaptic Plasma Membrane and Postsynaptic Density Proteins Using a Discontinuous Sucrose Gradient at JoVE.com

Preparation of Synaptic Plasma Membrane and Postsynaptic Density Proteins Using a Discontinuous Sucrose Gradient

Cet article décrit l&#39;enrichissement de protéines associées à la membrane plasmique synaptique par ultracentrifugation sur un gradient discontinu de saccharose. La préparation ultérieure des protéines de densités postsynaptiques est également décrite. préparations de protéines sont adaptés pour western blot ou analyse 2D DIGE.

Préparation de Synaptic membrane plasmique et de protéines de densité post-synaptique en utilisant un gradient discontinu de saccharose

Neuronal subcellular fractionation techniques allow the quantification of proteins that are trafficked to and from the synapse. As originally described in ...

preparation-synaptic-plasma-membrane-postsynaptic-density-proteins

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31.0K vues.  University of Toronto.  Cet article décrit l'enrichissement de protéines associées à la membrane plasmique synaptique par ultracentrifugation sur un gradient discontinu de saccharose. La préparation ultérieure des protéines de densités postsynaptiques est également décrite. préparations de protéines sont adaptés pour western blot ou analyse 2D DIGE. 

Vidéo: Preparation of Synaptic Plasma Membrane and Postsynaptic Density Proteins Using a Discontinuous Sucrose Gradient

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

Etudier Piscines vésicules synaptiques utilisant des colorants styryle photoconversion

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

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Membrane Potentials, Synaptic Responses, Neuronal Circuitry, Neuromodulation and Muscle Histology Using the Crayfish: Student Laboratory Exercises

The purpose of this report is to help develop an understanding of the effects caused by ion gradients across a biological membrane. Two aspects that influence a cell's membrane potential and which we address in these experiments are: (1) Ion concentration of K+ on the outside of the membrane, and (2) the permeability of the membrane to specific ions. The crayfish abdominal extensor muscles are in groupings with some being tonic (slow) and others phasic (fast) in their biochemical and physiological phenotypes, as well as in their structure; the motor neurons that innervate these muscles are correspondingly different in functional characteristics. We use these muscles as well as the superficial, tonic abdominal flexor muscle to demonstrate properties in synaptic transmission. In addition, we introduce a sensory-CNS-motor neuron-muscle circuit to demonstrate the effect of cuticular sensory stimulation as well as the influence of neuromodulators on certain aspects of the circuit. With the techniques obtained in this exercise, one can begin to answer many questions remaining in other experimental preparations as well as in physiological applications related to medicine and health. We have demonstrated the usefulness of model invertebrate preparations to address fundamental questions pertinent to all animals.

The experiments demonstrate an easy approach for students to gain experience in examining muscle structure, synaptic responses, the effects of ion gradients and permeability on membrane potentials. Also, a sensory-CNS-motor-muscle circuit is presented to show a means to test effects of compounds on a neuronal circuit.

Potentiels de membrane, les réponses synaptiques, circuits neuronaux, neuromodulation et histologie muscle en utilisant l&#39;écrevisse: des exercices de laboratoire étudiant

The purpose of this report is to help develop an understanding of the effects caused by ion gradients across a biological membrane. Two aspects that ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

Analyse quantitative des vésicules reconstitution Piscine synaptique dans des cultures de neurones granulaires du cervelet utilisant des colorants FM

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Preparation of Synaptoneurosomes from Mouse Cortex using a Discontinuous Percoll-Sucrose Density Gradient

Synaptoneurosomes (SNs) are obtained after homogenization and fractionation of mouse brain cortex. They are resealed vesicles or isolated terminals that break away from axon terminals when the cortical tissue is homogenized. The SNs retain pre- and postsynaptic characteristics, which makes them useful in the study of synaptic transmission. They retain the molecular machinery used in neuronal signaling and are capable of uptake, storage, and release of neurotransmitters.
The production and isolation of active SNs can be problematic using medias like Ficoll, which can be cytotoxic and require extended centrifugation due to high density, and filtration and centrifugation methods, which can result in low activity due to mechanical damage of the SNs. However, the use of discontinuous Percoll-sucrose density gradients to isolate SNs provides a rapid method to produce good yields of translationally active SNs. The Percoll-sucrose gradient method is quick and gentle as it employs isotonic conditions, has fewer and shorter centrifugation spins and avoids centrifugation steps that pellet SNs and cause mechanical damage.

A method to prepare translationally active, intact synaptoneurosomes (SNs) from mouse brain cortex is described. The method uses a discontinuous Percoll-sucrose density gradient allowing for the quick preparation of active SNs.

Préparation de Synaptoneurosomes de Mouse Cortex en utilisant un gradient de densité discontinu de Percoll-sucrose

Synaptoneurosomes (SNs) are obtained after homogenization and fractionation of mouse brain cortex. They are resealed vesicles or isolated terminals that ...

A General Method for Evaluating Incubation of Sucrose Craving in Rats

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and relapse to food taking 1. Food craving conceptualized in this way is akin to drug craving in response to drug-paired cues. A rich literature has been developed around understanding the behavioral and neurobiological determinants of drug craving; we and others have been focusing recently on translating techniques from basic addiction research to better understand addiction-like behaviors related to food 2-4.
As done in previous studies of drug craving, we examine sucrose craving behavior by utilizing a rat model of relapse. In this model, rats self-administer either drug or food in sessions over several days. In a session, lever responding delivers the reward along with a tone+light stimulus. Craving behavior is then operationally defined as responding in a subsequent session where the reward is not available. Rats will reliably respond for the tone+light stimulus, likely due to its acquired conditioned reinforcing properties 5. This behavior is sometimes referred to as sucrose seeking or cue reactivity. In the present discussion we will use the term "sucrose craving" to subsume both of these constructs.
In the past decade, we have focused on how the length of time following reward self-administration influences reward craving. Interestingly, rats increase responding for the reward-paired cue over the course of several weeks of a period of forced-abstinence. This "incubation of craving" is observed in rats that have self-administered either food or drugs of abuse 4,6. This time-dependent increase in craving we have identified in the animal model may have great potential relevance to human drug and food addiction behaviors.
Here we present a protocol for assessing incubation of sucrose craving in rats. Variants of the procedure will be indicated where craving is assessed as responding for a discrete sucrose-paired cue following extinction of lever pressing within the sucrose self-administration context (Extinction without cues) or as responding for sucrose-paired cues in a general extinction context (Extinction with cues).

Responding for food or drug-paired cues increases over the course of a period of abstinence and this may relate to an increased susceptibility to relapse behaviors. Here we detail a procedure for evaluating this "incubation of craving" in rats that have self-administered sucrose.

Une méthode générale pour l&#39;évaluation d&#39;incubation du désir saccharose chez les rats

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Diffusion latérale et l&#39;exocytose des protéines membranaires dans des neurones en culture évaluée à l&#39;aide de récupération de fluorescence et de fluorescence de perte de photoblanchiment

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Étiquetage immunologique post-enrobage des protéines synaptiques dans l&#39;hippocampe cultures Slice

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

Issu l&#39;évolution temporelle du glutamate Dégagement avec une analyse de déconvolution des courants de Transporteur ASTROCYTIQUES

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

The Neuromuscular Junction: Measuring Synapse Size, Fragmentation and Changes in Synaptic Protein Density Using Confocal Fluorescence Microscopy

The neuromuscular junction (NMJ) is the large, cholinergic relay synapse through which mammalian motor neurons control voluntary muscle contraction. Structural changes at the NMJ can result in neurotransmission failure, resulting in weakness, atrophy and even death of the muscle fiber. Many studies have investigated how genetic modifications or disease can alter the structure of the mouse NMJ. Unfortunately, it can be difficult to directly compare findings from these studies because they often employed different parameters and analytical methods. Three protocols are described here. The first uses maximum intensity projection confocal images to measure the area of acetylcholine receptor (AChR)-rich postsynaptic membrane domains at the endplate and the area of synaptic vesicle staining in the overlying presynaptic nerve terminal. The second protocol compares the relative intensities of immunostaining for synaptic proteins in the postsynaptic membrane. The third protocol uses Fluorescence Resonance Energy Transfer (FRET) to detect changes in the packing of postsynaptic AChRs at the endplate. The protocols have been developed and refined over a series of studies. Factors that influence the quality and consistency of results are discussed and normative data are provided for NMJs in healthy young adult mice.

The neuromuscular junction (NMJ) is altered in a variety of conditions that can sometimes culminate in synaptic failure. This report describes fluorescence microscope-based methods to quantify such structural changes.

La jonction neuromusculaire: Synapse mesurer la taille, fragmentation et les changements dans Synaptic Protein densité en utilisant microscopie de fluorescence confocale

The neuromuscular junction (NMJ) is the large, cholinergic relay synapse through which mammalian motor neurons control voluntary muscle contraction. ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.