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).

O protocolo a seguir está de acordo com as diretrizes do Conselho Canadense de Cuidado Animal e foi aprovado pela Faculdade de Medicina e do Comitê Animal Care Farmácia da Universidade de Toronto. 1 Prepare Obrigatório Reagentes conforme descrito na Tabela 1 <ol><li> Adicionar protease e da fosfatase (se necessário) os inibidores a todas as soluções de sacarose, tampões e ddH2O para concentrações descritas na Tabela 1. Importante: executar todos os passos a 4 ° C e pré-fresco todos os reagentes e equipamentos antes do início da experiência. Rotular todos os tubos, incluindo tubos de ultracentrífuga, com marcador permanente. </li></ol> 2 Dissecção da região do cérebro adequada <ol><li> Sacrificar o animal por deslocamento cervical ou decapitação. Casa e eutanásia de animais de acordo com as políticas institucionais e governamentais sobre cuidados com os animais. </li><li> Rapidamente remover ªe cérebro do crânio e colocar em um bloco de dissecção gelada. </li><li> Dissecar a região do cérebro apropriado de tecido fresco e vá para a Etapa 3. </li></ol> 3. Tissue homogeneização com Motor Impulsionada Glass-Teflon Homogeneizador <ol><li> Coloque 50-100 mg (de SPM), ou 100-200 mg (por PSD) do tecido dissecado em um tubo de polipropileno de 13 ml rotulados, contendo 4 ml de solução de sacarose 0,32 M tamponada com HEPES. Transfira a amostra para 15 ml de um tecido de vidro cónico-moinho Teflon / homogeneizador, definir o accionamento do motor para 900 rpm (posição 7) e homogeneizar a amostra com 12 pancadas ao longo de um período de 30 seg. Use um homogeneizador de vidro-Teflon diferente entre as amostras, ou lavar o homogeneizador com água fria destilada entre amostras e seque com um Kimwipe. NOTA: até 4 g de tecido podem ser homogeneizados em 4 ml de solução de sacarose 0,32 M tamponada com HEPES, no entanto, é aconselhável a utilização de menos tecido para assegurar o fraccionamento adequado. </li><li> Transferir a amostra homogeneizada para trás to o mesmo tubo de 13 ml de polipropileno. Reserva de 100 ul de homogeneizado e armazenar a -80 ° C para posterior quantificação de proteínas e análise de transferência de western da fracção de proteína total. </li></ol> 4. baixa velocidade de centrifugação para remover a Fração Nuclear (Yields sobrenadante S1) <ol><li> Centrifugar o homogenato num rotor de ângulo fixo a 900 xg durante 10 min a 4 ° C. Transferir o sobrenadante (S1) para um novo, rotulado 13 ml tubo e ressuspender o sedimento de fracção nuclear (P1) em 500 mL de sacarose 0,32 M tamponada com HEPES. NOTA: O (P1) fracção pode ser armazenada a -80 ° C e utilizada para a análise de transferência western da fracção nuclear. </li></ol> 5. enriquecimento da fracção bruta sinaptossomais (Yields Pellet P2) <ol><li> Centrifuga-se o sobrenadante (S1) a 10.000 xg durante 15 min a 4 ° C. Retirar o sobrenadante (S2) e 500 ul de reserva em um tubo de microcentrífuga para armazenar a -80 ° C para subsequente quant proteínasificação e western blot da fração citosólica da membrana / light. Ressuspender o sedimento fracção sinaptosomal em bruto remanescente (P2) em 1 ml de solução de sacarose 0,32 M tamponada com HEPES e, em seguida, adicionar uma outra solução de 3 ml de sacarose 0,32 M tamponada com HEPES. </li><li> Centrifuga-se a 10000 x g durante 15 min a 4 ° C. Retirar o sobrenadante (S2) e de reserva de 500 ul num tubo de microcentrífuga para armazenar a -80 ° C para posterior quantificação de proteínas e análise de Western blot da fracção citosólica membrana / luz. Salve o pellet sinaptosomal bruto lavado (P2 &#39;) no tubo de polipropileno. </li></ol> 6. Lise (hiposmótico Choque) da Fração Crude sinaptossomais <ol><li> Lise do sedimento sinaptosomal bruto (P2 &#39;) no tubo de polipropileno por ressuspensão lo em 1 ml de ddH 2 O. Adicionar mais 3 ml de DDH 2 O, transferir para um homogeneizador de tecido de vidro-Teflon, e homogeneizar à mão com três golpes. NOTA: É importantepara realizar esta etapa o mais rápido possível e rapidamente avançar para o passo 6.2. </li><li> Amostras de transferência do homogeneizador de volta na mesma 13 ml tubo de polipropileno e rapidamente ajustar a amostra de volta para HEPES 4 mM com 16 mL de uma solução M HEPES, inverter para misturar. </li><li> Rodar as amostras a 4 ° C durante 30 minutos para assegurar a lise completa. </li></ol> 7 Enriquecimento da Fração sinaptossomais Membrane (P3) <ol><li> Centrifuga-se a 25000 x g durante 20 min a 4 ° C. Retirar o sobrenadante fracção vesicular bruto (S3) e reservar com um tubo de 5 ml para armazenar a -80 ° C para posterior quantificação de proteínas e análise de transferência de western. Ressuspender a fracção de membrana sinaptosomal (P3) em 1 ml de solução de sacarose 0,32 M tamponada com HEPES. </li></ol> 8 Preparação do gradiente de sacarose descontínuo <ol><li> Pipetar exactamente 3,5 ml de solução de sacarose 1,2 M tamponada com HEPES e exactamente 3,0 ml cada de 1,0 M e 0,8 M HEPESolução de sacarose tamponada-S em separado 6 ml encaixar tubos de capitalização. NOTA: Isto é feito a pré-medir os volumes dos buffers que são usados ​​para fazer o gradiente de sacarose. Medição exacta dos volumes é importante para assegurar que os gradientes resultantes irá ser equilibrada por ultracentrifugação. </li><li> Usando uma pipeta Pasteur e bulbo de vidro, transferir 1,2 M HEPES solução de sacarose a 12 ml polialómero tubo ultracentrífuga. Colocar cuidadosamente a solução de sacarose 1,0 M HEPES no topo da solução de sacarose 1,2 M tamponado com HEPES. Por fim, a camada de solução de sacarose 0,8 M HEPES no topo da solução de sacarose 1,0 M tamponado com HEPES. Use uma pipeta Pasteur fresco para cada solução de sacarose e tomar cuidado para não perturbar o gradiente de sacarose. As vibrações de um vórtice banco ou microcentrifuge vai perturbar a integridade do gradiente. </li><li> Utilizando uma pipeta de Pasteur, a camada de fracção de membrana sinaptosomal ressuspenso (P3) na parte superior do gradiente de sacarose descontínuo preparado.Certifique-se de que os gradientes são equilibradas por pesá-las dentro do balde da caçamba móvel rotor. Se os recipientes não estão equilibrados, utilizar uma pipeta de P200 para adicionar cuidadosamente sacarose 0,32 M tamponada com HEPES para o topo do gradiente e equilibrar os baldes de rotor. </li></ol> 9 Fracionamento do Plasma Membrane Synaptic (SPM) <ol><li> Ultracentrífuga num rotor basculante a 150.000 xg durante 2 horas a 4 ° C. Remova cuidadosamente os gradientes de sacarose dos baldes de ultracentrífuga. Usando uma agulha de 18 G e seringa de 1 ml, perfurar o tubo no fundo da solução de sacarose 1,0 M interfase / 1,2 M de HEPES e retirar a camada de membrana de plasma sináptico (SPM). </li><li> Anote o volume que cada amostra SPM ocupa na seringa de 1 ml. Coloque a camada SPM recolhido em 3,5 ml de tubos de ultracentrífuga de parede espessa, adicionar exactamente 2,5 volumes de HEPES 4 mM para ajustar a concentração de sacarose de 1,2 M a 0,32 M. Depois de cada amostra foi ajustared para 0,32 M de sacarose, equilibrar os tubos com solução de sacarose 0,32 M HEPES. </li><li> Ultracentrífuga num rotor de ângulo fixo a 200.000 xg durante 30 min a 4 ° C. Remova e descarte o sobrenadante. Ressuspender o sedimento de membranas de plasma sináptico (SPM) em 300 ul de solução 50 mM de EDTA HEPES / 2 mM. Armazenar as amostras a -80 ° C até ser utilizado para transferência de western. </li></ol> 10 Preparação da Fração densidade pós-sináptico (PSD) <ol><li> Descongele amostras SPM no gelo. </li><li> Adicione uma solução de 0,54% de Triton X-100 em solução de EDTA a 50 mM de HEPES / 2 mM. </li><li> Em um tubo de poliestireno de 5 ml, 2,7 ml de combinar de Triton X-100 / HEPES / solução de EDTA e 300 ul fracção SPM e girar a amostra durante 15 min a 4 ° C </li><li> Amostras de transferência para 3,5 ml tubos de ultracentrífuga de parede espessa e amostras centrifugar a 32.000 xg por 20 min a 4 ° C. </li><li> Reservamo-nos o sobrenadante (Triton X-100 fração solúvel [TS]) e armazenar a -80; ° C. NOTA: Nesta fase, o pellet representa a fração &quot;PSD-1T&quot;. Um segundo tratamento com detergente vai produzir uma fracção de &quot;PSD-2T&quot;. Se uma fração PSD-1T é desejada, vá para o passo 10.6. <ol><li> Para produzir uma fracção PSD-2T, ressuspender o sedimento em 3 ml de 0,5% de Triton X-100 em 50 mM de HEPES / EDTA a 2 mM. Rodar as amostras durante 15 min a 4 ° C. Centrifugar as amostras a 32.000 xg por 20 min a 4 ° C e vá para a etapa 10.6. </li></ol></li><li> Descartar o sobrenadante e ressuspender o pós-sináptica (PSD) sedimento em solução de 50 HEPES / EDTA 2 mM. NOTA: O volume de ressuspensão pode depender do tamanho do pelete; Recomenda-se um volume de 50-75 mL. Porque a fracção PSD é uma fracção insolúvel em detergente, é muitas vezes necessário adicionar 1-2 mL de 0,5% de SDS, para permitir a ressuspensão completa de proteína. Armazenar as amostras a -80 ° C até ser utilizado para transferência 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.

Existem várias etapas do procedimento que são fundamentais para um bom resultado. No passo 3, é importante que um grau consistente de homogeneização é obtido para cada amostra. Quando homogeneizar o tecido com o homogeneizador accionado por motor, uma velocidade constante é utilizado não só para a rotação do pilão, mas também com o número de acidentes vasculares cerebrais. O tempo de incubação durante o choque osmótico deve ser mais preciso, uma vez que a homogeneização prolongada ou de incubação na...

Os neurônios se comunicam através de sinapses, ea qualidade desta comunicação é regulado, em grande medida por alterações na composição das proteínas na sinapse. Em particular, as proteínas localizadas na densidade pós-sináptica participar na comunicação neuronal por receptores de neurotransmissores intimamente andaime com os seus sistemas de transdução de sinal 1. Além disso, alterações na duração da força de eficácia sináptica são controladas por meio da adição ou remoção de receptores na densidade pós-sináptica 1-6. Portanto, o isolamento e quantificação de proteínas sinápticas é uma técnica útil e necessário para obter insights sobre os caminhos que os neurônios respondem a estímulos e alterar eficácia sináptica 7. Este artigo descreve uma técnica comum para isolar proteínas sinápticas de tecido cerebral dos roedores por ultracentrifugação em gradiente de sacarose descontínuo. A fracção de membrana de plasma sináptico podem ser enriquecidas e isoladas com base ema sua densidade em sacarose, que tenha sido determinada empiricamente para ser semelhante a 1,2 M de sacarose. Dependendo da causa biológica, fracções subcelulares podem ser separados por gradientes contínuos ou descontínuos de Percoll ou sacarose ou. Gradientes contínuos permitem a separação de proteínas em várias fracções; isto pode ser particularmente útil para demonstrar a co-localização de proteínas no interior de uma dada fracção 8. No entanto, a preparação de gradientes contínuos é mais trabalhoso e não é necessária para muitas aplicações. Gradientes descontínuos são comparativamente mais fáceis de preparar e pode ser usada para separar proteínas em algumas fracções, geralmente definidos. Gradientes descontínuos que são compostos por três camadas de aumento de sacarose molaridade têm sido amplamente utilizados para isolar as proteínas associadas com a membrana de plasma sináptico (SPM). Esta fracção de membrana de plasma sináptico pode ser ainda processado para a fractio densidade pós-sináptican (PSD) por tratamento com detergente e isolamento da fracção insolúvel em detergente. Quando este processo foi descrito pela primeira vez na década de 1960 9,10, microscopia eletrônica foi utilizada para demonstrar as organelas e membranas que cerca definem a membrana plasmática sináptica e frações de densidade pós-sináptica 9-14. Estes estudos demonstraram a inclusão de membranas pré-e pós-sinápticos e vesículas sinápticas em fracção do SPM; após tratamento com detergente principalmente o elétron-denso, densidade pós-sináptica eram visíveis. No processo, um choque hipotónico é usado para comprimir fora os processos sinápticas do corpo da célula 10. Esta etapa leva vantagem de o facto de que as mitocôndrias são mais resistentes a choque osmótico e permanecem intactos, de modo que eles e sedimentam na parte inferior do gradiente de sacarose (Figura 1). Usando a mesma técnica de enriquecimento, as frações de SPM e PSD foram os primeiros bioquimicamentedefinido por electroforese em gel de poliacrilamida e sequenciação dos principais componentes proteicos 15-17. Análise de mancha ocidental Subsequentemente foi usado para detectar e quantificar os níveis de proteínas sinápticas e ainda definir estas fracções (Figura 2). Nós temos usado essa técnica em nossos laboratórios para quantificar mudanças nos níveis sinápticos do transportador de dopamina que ocorrem quando o locus SLC6A3 é duplicada em camundongos 18. Também tenho usado essa técnica em receptores NMDA camundongos deficientes para descobrir reduções específicas de sinapse em proteínas que fazem parte da DISC1 interactome 19. É evidente a partir da análise de Western blot que fracções SPM contêm proteínas de membrana de vesícula sináptica, marcadores endossoma, proteínas mitocondriais, enzimas associadas à membrana sintéticas e moléculas de transdução de sinal, bem como componentes integrais da densidade pós-sináptica e as membranas de plasma sinápticas 20-23. Efracções PSD ven pode ter contaminação com proteínas mitocondriais abundantes e pode ser necessário realizar um segundo gradiente de sedimentação ou de passos de purificação adicionais para removê-los 13. Recentemente, a espectrometria de massa quantitativo tem fornecido uma lista de mais de 100 proteínas na densidade pós-sináptica por si só, bem como uma indicação da abundância relativa destes componentes 24,25.

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

Técnicas de fraccionamento subcelular neuronais permitir a quantificação de proteínas, que são objecto de tráfico de e para a sinapse. Tal como foi inicialmente descrito, no final dos anos 1960, as proteínas associadas à membrana plasmática sináptica pode ser isolado por ultracentrifugação em gradiente de densidade de sacarose. Uma vez que as membranas sinápticas são isolados, o complexo macromolecular conhecido como a densidade pós-sináptica pode ser subsequentemente isolado devido à sua insolubilidade em detergente. As técnicas utilizadas para isolar membrana plasmática sináptica e proteínas de densidade pós-sináptica permanecer essencialmente o mesmo depois de 40 anos, e são amplamente utilizados em pesquisas de neurociência atual. Este artigo fornece detalhes sobre o fraccionamento de proteínas associadas com a membrana plasmática sináptica e pós-sináptica de densidade utilizando um gradiente de sacarose descontínuo. Resultantes preparações de proteínas são adequados para western blot ou análise 2D DIGE.

A preparação do gradiente de densidade de sacarose deve resultar em uma separação clara entre as três soluções molares de sacarose (0,8, 1,0 e 1,2 M de sacarose). Ver Figura 3A para um exemplo do gradiente é adicionado antes da amostra de proteína. Se o gradiente é preparado com muita antecedência, ou se ele é preparado em uma superfície de bancada com a vibração de outro equipamento, o gradiente será comprometida e separação adequada não será alcançado. Se uma separação clara ent...

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

Este artigo fornece detalhes sobre o enriquecimento de proteínas associadas com a membrana de plasma sináptico por ultracentrifugação num gradiente de sacarose descontínuo. A elaboração definitiva de proteínas de densidade pós-sináptica é também descrito. Preparações de proteínas são adequados para western blot ou análise 2D DIGE.

Preparação do Synaptic Plasma Membrane e Proteínas de densidade pós-sinápticos Usando um gradiente de sacarose descontínuo

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

Research

JoVE Journal

Neuroscience

31.0K visualizações.  University of Toronto.  Este artigo fornece detalhes sobre o enriquecimento de proteínas associadas com a membrana de plasma sináptico por ultracentrifugação num gradiente de sacarose descontínuo. A elaboração definitiva de proteínas de densidade pós-sináptica é também descrito. Preparações de proteínas são adequados para western blot ou análise 2D DIGE. 

Vídeo: 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.

Estudando Pools vesículas Synaptic usando fotoconversão de Corantes Styryl

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

Neurociência

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.

Potenciais de membrana, Responses Synaptic, circuitos neuronais, neuromodulação e Histologia Muscle Usando a lagosta: Exercícios Laboratório Student

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.

Análise quantitativa de Reposição da vesícula sináptica Piscina em Cultured neurônios granulares do cerebelo usando Dyes 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.

Preparação de Synaptoneurosomes de rato Cortex usando um gradiente de densidade Percoll descontínuo Sacarose-

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.

Um método geral para a avaliação de Incubação de Craving Sacarose em Ratos

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.

Difusão lateral e Exocitose de proteínas de membrana em neurônios cultivados avaliada através da recuperação de fluorescência e fluorescência perda de fotobranqueamento

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.

Pós-incorporação de Etiquetagem immunogold de proteínas sinápticas em culturas de hipocampo

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.

Derivando o Curso momento do desembaraço glutamato com uma Análise de Deconvolução das correntes Transporter astrocitárias

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.

A Junção Neuromuscular: Synapse Medir o tamanho, Mudanças de fragmentação e em Synaptic Density proteína utilizando microscopia de fluorescência confocal

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.