Name: measuring synaptic vesicle endocytosis in cultured hippocampal neurons
Uploaded: 2017-09-04T12:30:00.000+00:00
Duration: 7 min 30 s
Description: Watch this Scientific Journal Video about Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons at JoVE.com

Agradecemos o Dr. Yong-Ling Zhu por prevê construção Sinaptofisina-pHluorin2x e Dr. James E. Rothman fornecendo VAMP2-phluorin. Agradecemos o Dr. Susan Cheng e Virginia Crocker da facilidade de microscopia eletrônica de NINDS pelo apoio técnico e ajuda. Este trabalho foi financiado pelo Instituto Nacional de Disorders Neurological e curso Intramural Research Program nos EUA e uma bolsa da KRIBB iniciativa programa de pesquisa (Korean biomédica cientista Fellowship Program), Instituto de pesquisa da Coreia do Biociências e biotecnologia, República da Coreia.

Nota: O seguinte protocolo descreve os métodos de imagem pHluorin e métodos EM neurônios hippocampal culta. pHluorin monitores vesícula sináptica proteína absorção em células vivas e EM detecta a absorção da vesícula sináptica e as alterações ultra-estruturais.
 cuidados com animais e procedimento seguiu as diretrizes do NIH e foram aprovados pelo Comitê de uso e cuidados do Animal NIH.
 1. pHluorin Imaging <ol><li>cultura neurônio Hippocampal <ol><li>preparar hipocampo Buffer (HB) combinando NaHCO 3 de 4 mM e 5 mM HEPES e ajustar o pH 7.3 com 5 M de NaOH. Fazer o meio de cultura misturando neurobasal médio, 2% B27, 0.5 mM L-glutamina e 1% penicilina-estreptomicina. Além disso, prepare uma mistura do buffer de HB com 20% de soro Fetal bovino (HB/20% FBS). 
		Nota: Este protocolo cultivo baseia-se na Serra, et al 29 e Wu et al 24</li> <li>decapitar filhotes de rato entre pós-Natal dia 0 ao dia 2 em meio de cultura e extrair o cérebro em 4 ° C HB/20% FBS. Retire o tronco cerebral e tálamo para expor o hipocampo. Dissecar para fora o hipocampo, após expor-lo pela remoção do tronco cerebral e tálamo e transferir para a 4 ° C HB/20% FBS. 
		Nota: Normalmente, o rendimento é em torno de 4 x 10 5 células/mL para um filhote de cachorro (dois hipocampo). Limpar as membranas aderente, usando uma pinça ou tesoura, então a transferência para a 4 ° C HB/20% FBS. Desenrole o giro denteado e isolar o subiculum cortando com uma tesoura e, em seguida, transferir a 4 ° C HB/20% FBS.</li>
		<li>Dividir cada hipocampo, por um corte de ponta a ponta, em cerca de 10 fatias e transfira para um tubo cónico polipropileno de 15 mL.</li>
		<li>Depois permitindo que o tecido resolver, lavar com 10 mL de HB/20% FBS e depois lavar três vezes com 10ml de HB.</li>
		<li>Preparar a solução de digestão de 137 mM de NaCl, 5 mM KCl, 7mm Na2HPO 4 e 25mm HEPES e ajustar o pH 7.2 com 5 M de NaOH. Retire o sobrenadante do hipocampo. Adicionar 2 mL de solução de digestão 10 mg de tripsina e 1 mg de DNase e filtrar através de uma membrana esterilizada 0,22 µm diretamente sobre a pelota amostra.</li>
		<li>Incubar o hipocampo por 5 min a 37 ° C, em seguida, lave duas vezes com 10 mL de HB/20% FBS e então lave uma vez com 10 mL de HB.</li>
		<li>Dissociar as células com 6 mg de MgSO 4 &#8729; 7 H 2 O e 1 mg de DNase 2ml HB e filtro estéril sobre o hipocampo de pelotas através de um filtro de 0,22 µm estéril. Delicadamente, dissocia as células por moer cuidado, enquanto tendo o cuidado de evitar a introdução de bolhas de ar. Permitir que as partículas de tecido repousar por 2 min e em seguida lentamente transfira o sobrenadante para outro tubo.</li>
		<li>, Adicionar 3 mL HB/20% FBS para a suspensão de células e centrífuga ou 10 min a 4 ° C e 1.000 rpm. Desprezar o sobrenadante e ressuspender em meio de cultura.</li>
		<li>Placa de 60.000 células suspendidas em meio de cultura de 150 µ l sobre uma lamela de diâmetro de 25 mm de poli-D-lisina revestido, sem derramamento da lamela. Adicione 2 mL de meio de cultura previamente aquecido 2 h após chapeamento. As lamelas são mantidas em placas de cultivo de 6 boas ou pratos de Petri estéril.</li>
		<li>Células mantêm a 37 ° C em um 5% CO 2 humedecidas incubadora em meio de cultura por 14-21 dias antes da gravação. Durante o crescimento da cultura, mudar a metade superior do meio duas vezes por semana.</li>
	</ol></li> <li>Transfection <ol><li>6-7 dias após o chapeamento, transfect Sinaptofisina-pHluorin 2 X (SpH) ou VAMP2-pHluorin em neurônios hippocampal. Para SpH, use um promotor de citomegalovírus (CMV), inserido em um vetor de pcDNA3 30. Para VAMP2-pHluorin, use um promotor CMV inserido em um vetor de pCI 31.</li>
		<li>Transfect qualquer vector por transportadora de lipídios em células-alvo, usando 1 µ g de plasmídeo. Use o meio de cultura de passo protocolo 1.1.1, que carece de soro, para o transfeccao. Alterar o meio 2 h após a transfeccao para reduzir a toxicidade. 
		Nota: Em caso de baixa expressão da SpH nos boutons, aumente a concentração de DNA de 2 µ g ou o tempo de incubação com a transportadora de lipídios, a menos que as células são insalubres. Normalmente, as células 4-10 (0,006-0,008%) de neurônios foram transfectadas.</li>
	</ol></li> <li>Microscopia de luz <ol><li>preparar solução salina normal é composto por 119 mM NaCl, 2 mM CaCl 2, 2,5 mM KCl, 25mm HEPES (pH 7,4), glicose de 30 mM, 2 mM MgCl 2, 6-cyano-7-nitroquinoxaline-2,3-dione (0.01 mM CNQX) e 0,05 mM ácido DL-2-amino-5-phosphonovaleric (AP5). Leve uma lamela da placa de meio de cultura e coloque em uma câmara de imagem que permite a estimulação do campo, usando lubrificante e selante para evitar vazamentos. Evite deixar o vidro seco durante a transferência de uma lamela da placa para a câmara, adicionando imediatamente a solução salina normal de 750 µ l. 
		Nota: CNQX AP5 utilizaram e bloquear a atividade pós-sináptica, que tem o potencial para atividade recorrente. Antes de colocar uma câmara em um microscópio de fluorescência invertido, use lenço de limpeza para confirmar que a câmara não vaza. Vazamento lento provoca alterações de foco durante a gravação, pela mistura de óleo normal saline e imersão, que resulta em mudanças no índice de refração.</li>
		<li>Estimulação e gravação de imersão de óleo <ol><li>imagem em um microscópio invertido widefield com um 60 X (1.4 abertura numérica) lente com uma lâmpada de iodetos metálicos. Visualizar pHluorin com um filtro definido para um pico de excitação de 480 nm, um espelho de passe longo 490 nm, um filtro de emissão 500-550 nm e um obturador manual de aleta. Capturar imagens cada 100 ms, com binning 2x2, usando uma câmera de elétron multiplicando carga acoplado dispositivo (EMCCD). 
			Nota: Vários recursos devem ser considerados para atingir as condições apropriadas para evitar fotobranqueamento, incluindo o intervalo de captura de imagem e filtrar e binning, que são dependentes de equipamentos. No caso da imagem latente confocal, tempo de alimentação e exposição do laser deve ser considerado para evitar fotobranqueamento. A instalação foi determinada após a gravação de pelo menos 3 min sem estímulo e a gravação foi realizada pelo menos 10 s sem estímulo para verificar se há fotobranqueamento.</li>
			<li>Escolher uma área com alta densidade de boutons para facilidade de análise em cada experimento. Identificar células transfectadas por seu sinal de fluorescência verde, com ênfase em padrões de expressão redondas ou ovais no bouton e expressão contínua entre boutons.</li>
			<li>Induzir endocitose por estimulação de campo usando um 1 pulso de ms, 20 mA ação potencial (AP) fornecida a partir de um estimulador de pulso e entregues através de um eletrodo de platina dentro da unidade de isolamento de estímulo. Atividade de fluorescência da imagem no decorrer da estimulação e durante a recuperação celular. 
			Nota: No caso de células mortas, expressão foi mais forte do que viver boutons e mostrou um padrão desigual entre boutons.</li>
		</ol></li></ol></li> <li>Análise de imagem <ol><li>Para analisar a intensidade de fluorescência em um único bouton, definido por uma região de interesse (ROI) como uma de 1,5 x 1,5 µm quadrados; o tamanho de um bouton é dentro traços cerca 1,5 µm. uso antes da estimulação para verificar fotobranqueamento e subtrair como pano de fundo.</li>
		<li>Normalize fluorescência muda (ΔF) com a equação: <img alt="Equation" height="11" src="/files/ftp_upload/55862/55862eq1.jpg" width="172" /> onde F máx e F 0 referem-se a máximo aumento após estimulação e fluorescência de base, respectivamente. Medir as endocitose taxas como a taxa de decaimento durante os primeiros 4-10 s após o ponto máximo da fluorescência de pHluorin. Obter a constante de tempo (τ) de endocitose por encaixe o decaimento de fluorescência pHluorin do pico aumento da linha de base com uma mono-exponencial função.</li>
	</ol></li></ol> 2. Microscopia eletrônica de <ol><li>Prepare uma poli-D-lisina revestido 6 placa aplicando 1,5 mL de solução de poli-D-lisina 0,01% estéril filtrada a cada poço por 1h à temperatura ambiente e, em seguida, lavar três vezes com água esterilizada. Dissecar, cultura e manter os neurônios hippocampal como em etapas 1.1.1, 1.1.2 e 1.1.3, respectivamente.</li>
	<li>Prepare uma solução de estimulação K + alta com HRP como 31,5 mM NaCl, 2 mM CaCl 2, 90 mM KCl, 25mm HEPES (pH 7,4), glicose de 30 mM, 2 mM MgCl 2, 0,01 mM CNQX, 0,05 mM AP5 e 5 mg/mL HRP, então ajustam o pH 7,4 com 5 M NaOH.
	Solução de estimulação de K + <ol><li>estimular a cultura de neurônio hippocampal com 1,5 mL de alta à temperatura ambiente (referida como K +) por adição de 1,5 mL para cada poço para 90 s. Na condição de repouso (referida como R), aplicar a mesma concentração de 5mg/mL HRP para 90 s, mas com solução salina normal. Para o exemplo de recuperação, aplicar solução de estimulação alta K + como com a amostra de K +, em seguida, rapidamente lavar substituir com solução salina normal e incubar durante 10 min.</li>
	</ol></li> <li>Fixação e coloração <ol><li>preparar o tampão de cacodylate at 0,1 M usando at cacodylate de 21,4 g/L pH 7,4. Consertar as células com 4% de glutaraldeído no buffer de cacodylate at 0.1 M pelo menos 1 h à temperatura ambiente. Lavar três vezes com 0,1 M at cacodylate o buffer para 7 min cada.</li>
		<li>Solução de preparar diaminobenzidina (DAB), composto de 0,5 mg/mL de DAB com 0,3% H 2 O 2 em ddH 2 O e filtrar com um filtro de 0,22 µm. Aplicar 1,5 mL de solução DAB por 30 min a 37 ° C. lavar três vezes com 0,1 M at cacodylate o buffer para 7 min cada. 
		Cuidado: DAB é um carcinógeno tóxico e suspeito. Por favor, use luvas e jalecos. 
		Nota: Etiquetando com DAB ocorre devido à sua oxidação por H 2 O 2, como catalisada por HRP. Pequenos aumentos nos componentes resultam em um sinal de aumento na amostra. Aumentando a concentração de HRP acelera o efeito do catalisador. Suficientemente grandes concentrações de H 2 O 2 permitem debilitante reações colaterais com HRP, inibindo o efeito da rotulagem 32. Neste trabalho, as concentrações deste sistema de rotulagem foram escolhidas com base na pesquisa atualmente disponíveis 33 , 34.</li>
		<li>Incubar os neurônios com 1,5 mL de 1% OsO 4 no buffer de cacodylate at 0,1 M por 1 h a 4 ° C, como fixação de postar. Lavar três vezes com 1,5 mL de tampão de cacodylate de nd de 0,1 M por 7 min cada. 
		Atenção: Devido à toxicidade e reatividade de OsO 4, mantendo a amostra no gelo em uma capa de química é preferível em muitos casos usar uma geladeira para a incubação.</li>
		<li>Preparar 0,1 M de tampão de acetato de sódio com 13,61 g/L de sódio acetato e 11,43 mL/L de ácido acético glacial em pH 5,0. Lavar três vezes com tampão de acetato 1,5 mL 0,1 M em pH 5.0 para 7 min cada e incubar com acetato de uranilo 1,5 mL 1% em tampão de acetato 0,1 M em pH 5.0 para 1 h a 4 ° C. Wash três vezes com tampão de acetato 1,5 mL 0,1 M por 7 min cada.</li>
	</ol></li> <li>Epóxi incorporação <ol><li>desidratar a cultura neurônio com lavagens único 1,5 mL de etanol a 50%, 70% e 90%, para 7 min em cada um e em seguida 3 lavagens de 1,5 mL de etanol a 100% por 7 min em uma coifa.</li>
		<li>Mix 485 mL/L bisphenol-A-(epichlorhydrin) epóxi resina, anidrido de 160 mL/L dodecenilo succínico (DDSA), 340 mL/L metil-5-Norborne-2,3-dicarboxílico fenol 2.4.6-tris (dimetilaminometil) anidrido (NMA) e 15 mL/L (DMP-30) para criar o epóxi resina. Mistura cuidadosamente, em seguida, armazenar sob vácuo para remover as bolhas de ar. 
		Nota: É fundamental para remover as bolhas de ar na resina, especialmente aqueles de menor do que é visível a olho nu, porque eles podem causar cáries na resina durante o corte.</li>
		<li>Infiltrar a amostra, substituindo o etanol com epóxi de 50% de resina em etanol por 30 min à temperatura ambiente em um shaker, então 70% epóxi resina em etanol por 30 min à temperatura ambiente num agitador.</li>
		<li>Interruptor o epóxi resina solução com resina epóxi de 100% e incubar durante 10 min a 50 bolsas de executar 2 ° C. de resina epóxi de 100% fresco com incubação por 1h em 50 ° C. Adicionar fresco 100% epóxi resina e permite a endurecer a 50 ° C durante a noite e depois no 60 & #17 6. o; C por mais de 36 h endurecer.</li>
	</ol></li> <li>Retire a placa de muti cada amostra com um joalheiro &#39; Serra de s. Selecione as regiões de interesse, densas concentrações de células, usando um microscópio invertido e depois cortar blocos de 70 a 80 nm para secionar pelo micrótomo. Montar a região de corte no mandril de micrótomo e carregar o micrótomo. Posicione o chuck em micrótomo e montar uma faca de diamante com a borda paralela à superfície do bloco. Recolher seções de espessura de 70 a 80 nm diretamente sobre grades individuais.</li>
	<li>Dissolver acetato de uranilo em água para uma solução de 1%, em peso e separadamente dissolver o citrato de chumbo na água para uma solução de 3% em peso. Counterstain as seções por submersão com acetato de uranilo aquosa de 1% por 15 min e, em seguida, citrato de chumbo aquoso 3% por 5 min aumentar o contraste das amostras.</li>
	<li>Imagem EM <ol><li>examinar as seções com um microscópio eletrônico de transmissão e gravar imagens com uma câmera digital CCD em uma ampliação primária de 10.000-20, 000 X 3.</li>
	</ol></li> <li>Estatísticas <ol><li>executar um t-teste para identificar diferenças significativas comparando-se a média e o desvio-padrão de medida (no MEV mostrou) entre o controle e amostras experimentais.</li>
	</ol>
	</li>
</ol>

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Aqui demonstramos dois métodos para monitorar a endocitose da vesícula sináptica. O primeiro método, temos monitorado pHluorin fundiu-se com uma proteína de vesícula sináptica nos neurônios transfectadas e posteriormente eletricamente estimulado. Em segundo lugar, usamos EM imagem latente de captação HRP como induzida por KCl. Usamos diferentes estímulos por duas razões. Primeiro, aplicação elevada de potássio induz a despolarização dos neurônios todos na cultura. Isso facilita o exame EM, dado que a no...

Os neurotransmissores são armazenados em vesículas sinápticas e liberados por exocitose. A membrana da vesícula sináptica e a proteína são então internalizadas por endocitose e reutilizadas na próxima rodada de exocitose. Endocitose das vesículas sinápticas é importante para a manutenção de piscinas de vesícula sináptica e remove salientes vesículas de membrana plasmática. O pHluorin sensíveis ao pH proteína verde fluorescente, que é saciada em circunstâncias ácidas e dequenched em pH neutro, tem sido usado para medir tempo de endocitose cursos em vivem células1,2,3. A proteína de pHluorin normalmente é unida ao lado lumenal das proteínas de vesículas sinápticas, tais como Sinaptofisina ou VAMP2/synaptobrevin da. No resto, pHluorin é extinguido no lúmen 5.5 pH das vesículas sinápticas. Fusão de vesículas para a membrana plasmática apresenta o lúmen vesicular à solução extracelular onde o pH é ~ 7.3, resultando em um aumento na fluorescência do pHluorin. Após a exocitose, decompõe-se o aumento da fluorescência, devido a endocitose das proteínas de vesículas sinápticas seguido por re-acidificação de vesículas dentro dessas vesículas recuperadas. Embora a decadência reflete tanto a endocitose e a re-acidificação vesicular, principalmente reflete endocitose, porque re-acidificação é mais rápido que endocitose na maioria das condições1,4. A constante de tempo de re-acidificação é 3-4 s ou menos5,6, que é geralmente mais rápido que o 10 s ou mais necessários para vesículas endocitose4,5. Se experimentos são necessários para distinguir a endocitose de re-acidificação, ácidos resfriamento experimentos usando a solução de (MES) ácido 4-Morpholineethanesulfonic (25 mM) com um pH de 5,5 podem ser usados para determinar se as proteínas de vesículas sinápticas são recuperadas da membrana plasmática através de endocitose1,3,4. Assim, o aumento de intensidade de fluorescência pHluorin reflete um equilíbrio de exo e endocitose, e a diminuição após a estimulação do nervo especificamente reflete endocitose.
pHluorin imagem pode ser usada não apenas para medir o tempo de curso de endocitose, mas também o tamanho da vesícula sináptica piscinas7,8, e9de lançamento a probabilidade de lançamento evocado e espontânea. Muitos fatores e proteínas envolvidas na regulação da endocitose, tais como cálcio, solúvel NSF-acessório da proteína do receptor (SNARE) proteínas, neurotrófico derivado do cérebro factor(BDNF) e calcineurina foram identificadas usando pHluorin de imagens1 , 2 , 10 , 11 , 12 , 13 , 14 , 15 , 16. Além disso, a liberação do neurotransmissor pode ser detectada nos neurônios não apenas primários, mas em células de neuroblastoma com TIRFM17. Recentemente, pHTomato, dsRed, mOrange e variantes de pHluorin foram desenvolvidos para o monitoramento de gravações simultâneas de vários fatores em uma única sinapse18,19. Por exemplo, pHTomato tem sido fundido com Sinaptofisina e usado com um indicador de cálcio geneticamente codificado (GCaMP5K) para monitorar fusão de vesícula pré-sináptica e Ca2 + influxo no compartimento pós-sináptica20. Portanto, pHluorin ligado a proteínas sinápticas fornece um método útil para analisar a relação entre endocitose e exocitose.
É outro método comumente usado para estudar a endocitose, devido a alta resolução espacial que mostra as alterações ultra-estruturais durante a endocitose. Duas áreas gerais são a capacidade de visualizar as alterações patológicas dentro de células neuronais21 e faixa de proteínas de vesícula22. Em particular, a observação da absorção da vesícula sináptica, curvatura de membrana revestida por Clatrina na zona de periactive, e CDDP estruturas são possíveis com EM3,23,24,25 ,26,,27,28. Enquanto EM envolve artefatos potenciais, tais como malformações induzidas pelo fixador, que pode afetar a endocitose e análise de dados é de trabalho intensivo, que a resolução prevê uma oportunidade atraente para visualizar a estrutura celular. Potenciais problemas de fixador e a limitação em resolução temporal EM podem ser superados por alta pressão, congelamento, fornecendo um método rápido e não-químicas de estabilizar as delicadas estruturas presentes durante a endocitose27.

<table><tbody><tr><td>Lipofectamine LTX with Plus</td><td>Thermo Fisher</td><td>15338-100</td><td>Transfection of plasmid DNA including synaptophysin or VAMP2-pHluorin</td></tr><tr><td>neurobasal medium</td><td>Thermo Fisher</td><td>21103-049</td><td>Growth medium for neuron, Warm up to 37&amp;deg;C before use</td></tr><tr><td>B27</td><td>Thermo Fisher</td><td>17504-044</td><td>Gradient for neuronal differentiation</td></tr><tr><td>Glutamax</td><td>Thermo Fisher</td><td>35050-061</td><td>Gradient for neuronal culture</td></tr><tr><td>Poly-D-Lysine coated coverslip</td><td>Neuvitro</td><td>GG-25-pdl</td><td>Substrate for neuronal growth and imaging of pHluorin</td></tr><tr><td>Trypsin XI from bovine pancrease</td><td>Sigma</td><td>T1005</td><td>Neuronal culture-digest hippocampal tissues</td></tr><tr><td>Deoxyribonuclease I from bovine pancreas</td><td>Sigma</td><td>D5025</td><td>Neuronal culture-inhibits viscous cell suspension</td></tr><tr><td>pulse stimulator</td><td>A-M systems</td><td>model 2100</td><td>Apply electrical stimulation</td></tr><tr><td>Slotted bath with field stimulation</td><td>Warner Instruments</td><td>RC-21BRFS</td><td>Apply electrical stimulation</td></tr><tr><td>stimulus isolation unit</td><td>Warner Instruments</td><td>SIU102</td><td>Apply electrical stimulation</td></tr><tr><td>lubricant</td><td>Dow corning</td><td>111</td><td>pHluorin imaging-seal with coverslip and imaging chamber, avoid leak from chamber</td></tr><tr><td>AP5</td><td>Tocris</td><td>3693</td><td>Gradient for normal saline, selective NMDA receptor antagonist, inhibit postsynaptic activity which have potential for recurrent activity</td></tr><tr><td>CNQX</td><td>Tocris</td><td>190</td><td>Gradient for normal saline, competitive AMPA/kainate receptor antagonist, inhibit postsynaptic activity which have potential for recurrent activity</td></tr><tr><td>Illuminator</td><td>Nikon</td><td>C-HGFI</td><td>Metal halide light source for pHluorin</td></tr><tr><td>EMCCD camera</td><td>Andor</td><td>iXon3</td><td>pHluorin imaging, detect pHluorin fluorescence intensity</td></tr><tr><td>Inverted microscopy</td><td>Nikon</td><td>Ti-E</td><td>Imaging for synaptophysin or VAMP2 pHluorin transfected cells</td></tr><tr><td>NIS-Elements AR</td><td>Nikon</td><td>NIS-Elements Advanced Research</td><td>Software for imaging acquisition and analysis</td></tr><tr><td>Igor Pro</td><td>WaveMetrics</td><td>Igor pro</td><td>Software for imaging analysis and data presentation</td></tr><tr><td>imaging chamber</td><td>Warner Instruments</td><td>RC21B</td><td>pHluorin imaging, apply field stimulation on living cells</td></tr><tr><td>poly-l-lysine</td><td>Sigma</td><td>P4832</td><td>Electron microscopy, substrate for neuronal growth, apply on multiwell plate for 1 h at room temperature then wash with sterilized water 3 times</td></tr><tr><td>Horseradish peroxidase(HRP)</td><td>Sigma</td><td>P6782</td><td>Electron microscopy, labeling of endocytosed synaptic vesicles by catalyzing DAB in presence hydrogen peroxide, final concentration is 5 mg/mL in normal saline, make fresh before use</td></tr><tr><td>Na cacodylate</td><td>Electron Microscopy Sciences</td><td>12300</td><td>Electron microscopy, buffer for fixatives and washing, final concentration is 0.1 N</td></tr><tr><td>3,3&amp;prime;-Diaminobenzidine(DAB)</td><td>Sigma</td><td>D8001</td><td>Electron microscopy, labeling of endocytosed synaptic vesicles, substrate for HRP, final concentration is 0.5 mg/mL in DDW and filtered, make fresh before use</td></tr><tr><td>Hydrogen peroxide solution</td><td>Sigma</td><td>H1009</td><td>Electron microscopy, labeling of endocytosed synaptic vesicles by inducing HRP-DAB reaction, final concentration is 0.3% in DDW, make fresh before use</td></tr><tr><td>glutaraldehyde</td><td>Electron Microscopy Sciences</td><td>16365</td><td>Electron microscopy, fixatives, final concentration is 4% in Na-cacodylate buffer, make fresh before use, shake well before to use</td></tr><tr><td>TEM</td><td>JEOL</td><td>200CX</td><td>Electron microscopy, imaging of endocytosed vesicles and ultrastructural changes</td></tr><tr><td>CCD digital camera</td><td>AMT</td><td>XR-100</td><td>Electron microscopy, capturing images</td></tr><tr><td>Lead citrate</td><td>Leica microsystems</td><td>16707235</td><td>Electron microscopy, grid staining</td></tr></tbody></table>

measuring synaptic vesicle endocytosis in cultured hippocampal neurons

Durante a endocitose, fundidas vesículas sinápticas são recuperadas em terminais nervosos, permitindo a reciclagem de vesículas e, portanto, a manutenção da transmissão sináptica durante o disparo repetitivo do nervo. Endocitose prejudicada em condições patológicas leva a diminuições em funções sinápticas de força e cérebro. Aqui, descrevemos os métodos utilizados para medir a endocitose da vesícula sináptica na sinapse hippocampal mamífera na cultura neuronal. Temos monitorado endocitose de proteínas de vesícula sináptica pela fusão de uma proteína de membrana vesicular sináptica, incluindo Sinaptofisina e VAMP2/synaptobrevin da, o lado lumenal vesicular, com pHluorin, uma proteína fluorescente verde sensíveis ao pH que aumenta sua intensidade de fluorescência como o pH aumenta. Durante a exocitose, pH do lúmen vesicular aumenta, Considerando que durante o lúmen vesicular endocitose pH é re-acidificado. Assim, um aumento da intensidade de fluorescência pHluorin indica fusão, enquanto uma diminuição indica endocitose da proteína rotulados vesícula sináptica. Além de usar o pHluorin método de imagem para gravar endocitose, temos monitorado endocitose da membrana vesicular por microscopia de elétron (EM) medições de absorção de Horseradish peroxidase (HRP) por vesículas. Finalmente, temos monitorado a formação dos poços de membrana terminal do nervo em vários momentos após a despolarização induzida por potássio alto. O tempo de curso de formação de poço de captação e membrana HRP indica o tempo de curso de endocitose.

Usando o método de transportadora de lipídios, SpH foi expressa em neurônios hippocampal, permitindo a identificação de boutons (Figura 1a). Estimulação elétrica das células induzidas por exocitose e um correspondente aumento na intensidade de fluorescência. O aumento na fluorescência (ΔF) foi interrompido por encerrar o estímulo (Figura 1b). O aumento da fluorescência foi seguido por uma diminuição lenta, devido ...

Watch this Scientific Journal Video about Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons at JoVE.com

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

Endocitose da vesícula sináptica é detectada pela microscopia de luz de pHluorin fundido com proteínas de vesículas sinápticas e por microscopia eletrônica de absorção da vesícula.

Medindo a endocitose da vesícula sináptica nos neurônios Hippocampal culturas

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic ...

measuring-synaptic-vesicle-endocytosis-in-cultured-hippocampal-neurons

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Neuroscience

9.8K visualizações.  National Institute of Neurological Disorders and Stroke. Endocitose da vesícula sináptica é detectada pela microscopia de luz de pHluorin fundido com proteínas de vesículas sinápticas e por microscopia eletrônica de absorção da vesícula.

Vídeo: Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

Measuring Exocytosis in Neurons Using FM Labeling

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time imaging of vesicles labeled with FM dye to monitor the rate of presynaptic vesicle release.  FM4-64 is a red fluorescent amphiphilic styryl dye that embeds into the membranes of synaptic vesicles as endocytosis is stimulated.  Lipophilic interactions cause the dye to greatly increase in fluorescence, thus emitting a bright signal when associated with vesicles and a nominal one when in the extracellular fluid. After a wash step is used to help remove external dye within the plasma membrane, the remaining FM is concentrated within the vesicles and is then expelled when exocytosis is induced by another round of electrical stimulation. The rate of vesicles release is measured from the resulting decrease in fluorescence.  Since FM dye can be applied external and transiently, it is a useful tool for determining rates of exocytosis in neuronal cultures, especially when comparing the rates between transfected synapses and neighboring control boutons.



The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission. Here we used real-time imaging of vesicles labeled with the red fluorescent dye FM 4-64 to measure the rate of presynaptic vesicle release in hippocampal neuronal cultures.

Medição Exocitose em Neurônios Usando Labeling FM

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time ...

Biologia

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Vibrodissociation de neurônios de fatias do cérebro de roedores para estudar a transmissão sináptica e Imagem Terminais pré-sináptica

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Neurociência

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 ...

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 ...

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Imagens ao vivo de vesículas com núcleo denso na Primária Cultivadas neurônios do hipocampo

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

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 ...

Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using styryl FM dyes, which reveal membrane turnover. Conventional protocols for the use of FM dyes were designed for analyzing neurons following stimulated (evoked) synaptic activity. Recently, protocols have become available for analyzing the FM signals that accompany weaker synaptic activities, such as spontaneous or miniature synaptic events. Analysis of these small changes in FM signals requires that the imaging system is sufficiently sensitive to detect small changes in intensity, yet that artifactual changes of large amplitude are suppressed. Here we describe a protocol that can be applied to evoked, spontaneous, and miniature synaptic activities, and use cultured hippocampal neurons as an example. This protocol also incorporates a means of assessing the rate of photobleaching of FM dyes, as this is a significant source of artifacts when imaging small changes in intensity.

We describe the use of styryl FM dyes to image synaptic vesicle recycling in functional nerve terminals. This protocol can be applied not only to evoked, but also spontaneous and miniature synaptic activities. The protocol expands the variety of synaptic events that can be effectively evaluated.

Exame do Synaptic Vesicle Reciclagem Usando FM Corantes Durante Evocado, espontânea, e as atividades de Synaptic miniatura

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

Quantificação de Endosome e Moosidades de Lisosoma em Neurônios Cultivados Usando Sondas Fluorescentes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane proteins become exposed at the cell surface. One of these proteins is Synaptotagmin-1 (Syt1). An antibody directed against the luminal domain of Syt1, once added to the culture medium, is taken up during the exo-endocytotic cycle. This uptake is proportional to the amount of SV recycling and can be quantified through immunofluorescence. Here, we combine Syt1 antibody uptake with double transfection of cultured hippocampal neurons. This allows us to (1) localize presynaptic sites based on expression of recombinant presynaptic marker Synaptophysin, (2) determine their functionality using Syt1 uptake, and (3) characterize the targeting and effects of a protein of interest, GFP-Rogdi.

We describe an optical assay for synaptic vesicle (SV) recycling in cultured neurons. Combining this protocol with double transfection to express a presynaptic marker and protein of interest allows us to locate presynaptic sites, their synaptic vesicle recycling capacity, and determine the role of the protein of interest.

Um ensaio óptico para vesícula sináptica reciclagem nos neurônios cultivados superexpressão de proteínas pré-sináptica

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane ...

Transmission Electron Microscopic Imaging of Synaptic Vesicle Endocytosis in Mouse Hippocampal Neurons 

<a href='https://app.jove.com/v/55862/measuring-synaptic-vesicle-endocytosis-in-cultured-hippocampal-neurons'>Source: Villarreal, S., et. al., Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons. J. Vis. Exp. (2017)</a>This video demonstrates the analysis of synaptic vesicle endocytosis in cultured hippocampal neurons through horseradish peroxidase labeling and electron microscopy.

Source: Villarreal, S., et. al., Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons. J. Vis. Exp. (2017)This video demonstrates the ...