Este trabalho foi financiado pelo National Institutes of Health (K22NS050137 a JCB).

1. Cultura de células <ol><li> Neurônios corticais estão isolados e cultivados com técnicas previamente descritas e semeadas em placas de cultura com um fundo de vidro (Mattek Corporation, Ashland, MA) revestidas com poli-D-lisina e laminina 1. </li></ol> 2. Preparação das soluções estoque para o fluorescente sondas TMRM e 2 H DCF-DA <ol><li> Prepare uma solução estoque de 10 mM de TMRM pela dissolução de 5,0 mg de TMRM em 1 ml de dimetilsulfóxido anidro. Vortex por 1 min. Então, faça alíquotas e armazená-los a -20 ° C, proteger da luz, e usar dentro de um mês. </li><li> Em seguida, preparar uma solução estoque de 10 mM de H 2 DCF-DA dissolvendo 4,87 mg de H 2 DCF-DA em 1 ml de DMSO anidro. Da mesma forma vortex, por 1 min. Então, faça alíquotas e armazená-los a -20 ° C, proteger da luz, e usar dentro de uma semana. </li></ol> 3. Carregando neurônios de rato cortical com TMRM e 2 H DCF-DA TMRM é um potenciométrica, indicador de célula-permeável fluorescentes que se acumula no interior altamente carregadas negativamente das mitocôndrias. É importante usar a baixas concentrações (10-50 nm) de TMRM para evitar a auto-extinção da TMRM mitocondrial. Então, o sinal de fluorescência de TMRM pode ser diretamente co-relacionados com a ΔΨm através da membrana mitocondrial interna. A perda de ΔΨm causas TMRM a fuga de mitocôndrias, resultando em uma perda de intensidade de fluorescência. H 2 DCF-DA é a célula-permeável sonda convertido em DCF-DA por esterases intracelulares, e os resultados a sua oxidação no DCF fluorescente. A concentração final de H 2 DCF-DA varia entre 2-10 mM e deve ser testado empiricamente em neurônios derivados de diferentes regiões do cérebro vez que a concentração de carga elevada pode resultar na saturação de fluorescência DCF, mesmo na ausência de H 2 O 2. A presença de qualquer oxidante endógeno ou exógeno (por exemplo, óxido nítrico, peróxido de hidrogênio) aumentará DFC intensidade de fluorescência. Abaixo, fornecemos um protocolo para o carregamento com neurônios de rato cortical TMRM e 2 H DCF-DA. <ol><li> Para carregar os neurônios de rato cortical com TMRM, primeiro, lave os neurônios cultivados 3 vezes com tampão Tyrode do (Overlay Texto: TB: 145 mM NaCl, 5 mM KCl, 10 mM de glucose, 1,5 mM CaCl 2, 1 mM MgCl 2, e 10 HEPES mM; ajustar o pH para 7,4 com NaOH). Então, prepare 20 nM de TMRM diluindo a 10 mM de ações TMRM 1 / 1000 vezes em TB e depois adicionar 2 l da TMRM diluído por 1 ml de TB. Incubar os neurônios com TMRM por 45 minutos no escuro à temperatura ambiente. Após 45 min, monte o prato de cultura no palco do microscópio e começar de imagem. </li><li> Para carregar os neurônios de rato cortical com H 2 DCF-DA, lave os neurônios cultivados 3 vezes com TB. Em seguida, prepare 2 mM de H 2 DCF-DA diluindo a 10 mM H 2 DCF-DA estoque 10/01 vezes em TB e depois adicionar 2 l da diluição H 2 DCF-DA por 1 ml de TB. Então, incubar os neurônios com H 2 DCF-DA por 45 minutos no escuro à temperatura ambiente. Após 45 min, lave os neurônios 4 vezes com TB para remover indicador fluorescente excesso antes de obter imagens. </li></ol> 4. Ao vivo imagens de neurônios incubadas com TMRM para determinar ΔΨm <ol><li> Para realizar imagens ao vivo de neurônios incubadas com TMRM, microscopia confocal de varredura a laser (Overlay Texto: LSM 510, Carl Zeiss Inc.), com a aplicação do programa de séries temporais ao vivo, é usado. Aplicar de baixa resolução e atenuada a laser de potência (Overlay Texto: baixa resolução: 256 x 256; potência do laser: 1%) para minimizar o tempo necessário para obter imagens e evitar fotodegradação. </li><li> . Em seguida, ajustar o foco dos neurônios montado carregado com TMRM usando luz refletida. Examine a fluorescência TMRM pela iluminação a 514 nm e detecção a 570 nm. Definir o ganho de detecção de uma câmera, logo abaixo do nível de saturação. </li><li> Uma vez que todos os parâmetros que incluem resolução, potência do laser, o ganho de detecção de uma câmera, e lapso de tempo de intervalo para obter imagens são definidas, não alterar essas configurações entre experimentos. Em seguida, altere o campo. Começar a recolher imagens. </li><li> Para testar as alterações no ΔΨm, estímulos como uma mM de FCCP ou 2 mg / ml de oligomicina, pode ser aplicado, de forma significativa ou despolarizar hiperpolarizar o potencial de membrana mitocondrial, respectivamente. Estas alterações serão refletidas por uma diminuição na intensidade de fluorescência TMRM comparada com a intensidade de fluorescência de base, no caso de FCCP, ou um aumento na intensidade de fluorescência TMRM no caso de oligomicina. </li></ol> 5. Imagens ao vivo de neurônios incubadas com H 2 DCF-DA para determinar ROS <ol><li> Para realizar imagens ao vivo de neurônios incubadas com H 2 DCF-DA, primeiro, montar a placa de cultura no palco de um microscópio. Ajustar o foco das células-nosing luz refletida. Examine DCF fluorescência por excitação a 488 nm e emissão a 515 nm. </li><li> Em seguida, ajuste a potência do laser para 5-7%, o ganho do detector, e resolução de 256 x 256. Não altere estas configurações entre experimentos. Em seguida, defina a freqüência para a obtenção de imagens ao vivo usando o programa de séries temporais. </li><li> Selecione um campo novo e começar a adquirir as imagens. Para detectar mudanças nos níveis de ROS, tratar células com 100-200 mM de H 2 O 2. Isso será refletido por um aumento na intensidade de fluorescência DCF em comparação com o nível basal. </li></ol> 6. Análise de dados <ol><li> Use região de interesse (Overlay Texto: ROI) ferramenta do programa de LSM para selecionar as áreas. Então, medir o TMRM ou ROS intensidades de fluorescência. Selecione ROIs de regiões mitocondriais ou ROIs do corpo celular em células de toda fotografada para medir a intensidade de fluorescência de TMRM ou ROS, respectivamente. </li><li> Calcular a intensidade média de fluorescência de todos os ROIs de cada célula para TMRM ou de corpos de células inteiras para todas as células com imagens de ROS para cada ponto de tempo. Selecione regiões próximas às células para calcular a intensidade de fluorescência de fundo. Tomar várias medidas e calcular a intensidade média de fundo. </li><li> Subtrair a intensidade de fluorescência média de fundo de média intensidades de fluorescência das ROIs em cada célula para cada ponto de tempo usando o Microsoft Excel. Depois de subtrair a intensidade de fundo, normalizar a intensidade TMRM ou DCF fluorescência para a fluorescência de base usando esta fórmula (Overlay Texto: ΔF = FF o / F o x 100, onde F = intensidade de fluorescência em qualquer ponto do tempo, Para = linha de base de fluorescência). Então, use o programa Sigma Plot para gerar o gráfico mostrando as mudanças na intensidade de fluorescência ao longo do tempo. </li></ol> 7. Resultados representante Figura 1A mostra uma imagem de fluorescência de neurônios corticais de rato incubadas com TMRM. Adição de FCCP, um desacoplador mitocondrial, leva a despolarização mitocondrial e uma perda de TMRM intensidade de fluorescência (Figura 1B). A linha de base nível de fluorescência TMRM permanece estável antes da adição de FCCP (o primeiro 350 seg;. Fig 1C). Análise quantitativa de fluorescência TMRM mudanças ao longo do tempo mostra uma diminuição significativa na fluorescência TMRM após a adição de FCCP (Fig. 1C). Figura 1D mostra a imagem de fluorescência de neurônios corticais de rato carregado com DCF. Adição de H 2 O 2 resulta em aumento da intensidade de fluorescência DCF nos corpos celulares (Fig. 1E). A linha de base DCF nível de fluorescência mantém-se inalterado (os primeiros 120 segundos) antes da aplicação de H 2 O 2. Lapso de tempo de medições de fluorescência DCF mostram seus níveis estáveis, o que aumenta depois de H 2 O 2 de tratamento (Fig. 1F). <img src="/files/ftp_upload/2704/2704fig1.jpg" alt="Figura 1"/> Figura 1. Avaliação do potencial de membrana mitocondrial e os níveis de ROS em neurônios corticais rato vivo. (A) Representante imagem de fluorescência de neurônios corticais carregado com TMRM. Após a digitalização da linha de base TMRM de fluorescência, os neurônios foram tratados com o FCCP protonophore (1 mM). Para a direita é um bar pseudo intensidade de fluorescência TMRM com o máximo de amarelo e preto brilhante representação e intensidade mínima, respectivamente. A perda de fluorescência TMRM das regiões mitocondrial indica o colapso do ΔΨm sobre FCCP tratamento (painel B). A representação quantitativa da mudança na intensidade de fluorescência TMRM em momentos diferentes, antes e após o tratamento FCCP é mostrado no painel C. (D) a imagem de fluorescência de neurônios corticais de rato carregado com H2DCF-DA. Após a determinação da linha de base DCF fluorescência, as células foram tratadas com 200 mM H 2 O 2, ea mudança no DCF fluorescência foi avaliada. Um aumento na fluorescência do DCF reflete o aumento nos níveis de ROS em H 2 O 2 tratamento (E). Análise quantitativa das mudanças no DCF fluorescência, antes e depois do H 2 O 2 tratamento, é mostrado no painel da barra de escala F. = 10 mM Video.7.1 - labmedia 2704_Joshi.avi Ao vivo imagens de células de TMRM em neurônios corticais antes e depois disso FCCP utilizando objetiva de 40X. A intensidade mostra um pseudo (amarelo brilhante, antes da adição FCCP) e diminuição máxima (cor vermelha, após a adição FCCP) TMRM intensidade de fluorescência após a adição FCCP. <a href="http://www.jove.com/files/ftp_upload/2704/supplemental_video1.avi">Clique aqui para ver o vídeo</a> Vídeo. 7.5 - labmedia 2704_Joshi.avi Ao vivo imagens de células de DCF nos neurônios corticais antes e depois do H 2 O 2 disso utilizando objetiva de 40X. A linha de base DCF fluorescência tem cor verde-claro nos corpos celulares e H2O2 disso aumenta o DCF intensidade de fluorescência de cor verde brilhante. <a href="http://www.jove.com/files/ftp_upload/2704/supplemental_video2.avi">Clique aqui para ver o vídeo</a>

<ol>
	<li>Hilgenberg, L. G., Smith, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+dissociated+mouse+cortical+neuron+cultures.">Preparation of dissociated mouse cortical neuron cultures.</a> J. Vis. Exp. , (2007).</li><li>Scaduto, R. C., Grotyohann, L. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+mitochondrial+membrane+potential+using+fluorescent+rhodamine+derivatives.">Measurement of mitochondrial membrane potential using fluorescent rhodamine derivatives.</a> Biophys. J. 76, 469-477 (1999).</li><li>Ward, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+amyloid+precursor+protein+intracellular+domain+(AICD)+disrupts+actin+dynamics+and+mitochondrial+bioenergetics.">The amyloid precursor protein intracellular domain (AICD) disrupts actin dynamics and mitochondrial bioenergetics.</a> J. Neurochem. 113, 275-284 (2010).</li><li>Gunasekar, P. G., Kanthasamy, A. G., Borowitz, J. L., Isom, G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NMDA+receptor+activation+produces+concurrent+generation+of+nitric+oxide+and+reactive+oxygen+species:+implication+for+cell+death.">NMDA receptor activation produces concurrent generation of nitric oxide and reactive oxygen species: implication for cell death.</a> J. Neurochem. 65, 2016-2021 (1995).</li></ol>

Não há conflitos de interesse declarados.

Nós apresentamos um procedimento passo-a-passo descreve como determinar ΔΨm e ROS em neurônios corticais de rato usando o fluorescente indicadores TMRM e H 2 DCF-DA, respectivamente. Para outros tipos de células, é importante determinar empiricamente a concentração final e tempo de carregamento para TMRM ou 2 H DCF-DA. Em geral, as concentrações de gama TMRM 2-20 nM, eo tempo de incubação das células com TMRM varia de 20 a 60 min. A concentração final de H 2 DCF-DA faixas de 2-10 mM, e incubação de células em uma solução de carga contendo este indicador varia de 30-45 min. É importante para otimizar a potência do laser e velocidade de digitalização de levar as imagens para evitar a foto-toxicidade para as células e as mudanças na intensidade de fluorescência (por exemplo cintilação de TMRM fluorescência), na ausência de qualquer estímulo. As configurações otimizadas óptico deve resultar em um sinal de fluorescência que não está acima ou abaixo saturadas (threshold) na ausência de estímulo. As condições ideais para coletar as imagens de um campo selecionado em uma potência do laser em particular e uma velocidade de digitalização são obtidas quando não há mudanças na intensidade de fluorescência da sonda na ausência de qualquer estímulo por 10-15 min de imagens ao vivo. Outras sondas de fluorescência para determinar ΔΨm incluem rodamina 123 e tetra metil etil éster rodamina (TMRE). No entanto, eles foram encontrados para inibir os processos respiratórios em dois mitocôndrias isoladas. Importante, TMRM não tem nenhum efeito sobre a respiração mitocondrial em baixas concentrações 2 e tem fototoxicidade baixa e fotobranqueamento 3, em comparação com outras sondas. H 2 DCF-DA é um bom indicador para ROS como é bem retida nas células e reconhece as espécies oxidantes diversos, tais como peróxidos, óxidos de super, e óxido nítrico 4.

<table border="1"><tbody><tr><th> Nome do reagente </th><th> Companhia </th><th> Número de catálogo </th></tr><tr><td> De vidro prato de cultura de fundo </td><td> Mattek Corporação </td><td> P35G-1.5-14-C </td></tr><tr><td> NbActive4 </td><td> BrainBits </td><td> NbActive4 </td></tr><tr><td> TMRM </td><td> Invitrogen </td><td> T668 </td></tr><tr><td> H 2 DCF-DA </td><td> Invitrogen </td><td> C400 </td></tr><tr><td> NaCl </td><td> Sigma </td><td> S6191 </td></tr><tr><td> KCl </td><td> Sigma </td><td> P3911 </td></tr><tr><td> • CaCl 2 2H 2 O </td><td> Sigma </td><td> C3306 </td></tr><tr><td> MgCl 2 • 6H 2 O </td><td> Sigma </td><td> M2670 </td></tr><tr><td> D-Glicose </td><td> Sigma </td><td> G6152 </td></tr><tr><td> HEPES </td><td> Invitrogen </td><td> 15630 </td></tr></tbody></table>

determination of mitochondrial membrane potential and reactive oxygen species in live rat cortical neurons

Potencial de membrana mitocondrial (ΔΨm) é fundamental para manter a função fisiológica da cadeia respiratória para gerar ATP. A perda significativa de células ΔΨm torna esgotado de energia, com posterior morte. Espécies reativas de oxigênio (ROS) são importantes moléculas sinalizadoras, mas a sua acumulação em condições patológicas leva ao estresse oxidativo. As duas principais fontes de ROS nas células são toxinas ambientais e do processo de fosforilação oxidativa. Disfunção mitocondrial eo estresse oxidativo têm sido implicados na fisiopatologia de muitas doenças e, portanto, a capacidade de determinar ΔΨm e ROS pode fornecer pistas importantes sobre o estado fisiológico da célula ea função das mitocôndrias. Várias sondas fluorescentes (Rodamina 123, TMRM, TMRE, JC-1) pode ser usado para determinar Δψm em uma variedade de tipos celulares, e muitos indicadores de fluorescência (Dihydroethidium, Dihydrorhodamine 123, H 2 DCF-DA) pode ser usado para determinar ROS . Quase todas as sondas de fluorescência disponíveis utilizados para avaliar ΔΨm ou ROS são um único comprimento de onda de indicadores, que aumentar ou diminuir sua intensidade de fluorescência proporcional a um estímulo que aumenta ou diminui os níveis de ΔΨm ou ROS. Assim, é imperativo para medir a intensidade de fluorescência destas sondas ao nível basal e após a aplicação de um estímulo específico. Isto permite uma para determinar a porcentagem de mudança na intensidade de fluorescência entre o nível de referência e um estímulo. Esta mudança na intensidade da fluorescência reflete a mudança nos níveis relativos de ΔΨm ou ROS. Neste vídeo, demonstramos como aplicar o indicador de fluorescência, TMRM, em neurônios corticais de ratos para determinar a variação percentual na intensidade de fluorescência TMRM entre o nível de base e após a aplicação de FCCP, um desacoplador mitocondrial. Os níveis mais baixos de TMRM fluorescência resultante do tratamento FCCP refletem a despolarização do potencial de membrana mitocondrial. Também mostramos como aplicar a fluorescência da sonda H 2-DA DCF para avaliar o nível de ROS em neurônios corticais, primeiro na linha de base e, em seguida, após a aplicação de H 2 O 2. Este protocolo (com pequenas modificações) também podem ser usados ​​para determinar mudanças no ΔΨm e ROS em diferentes tipos celulares e em neurônios isolados de outras regiões do cérebro.

Watch this Scientific Journal Video about Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons at JoVE.com

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Nós demonstramos a aplicação do indicador de fluorescência, TMRM, em neurônios corticais para determinar a alterações relativas na intensidade de fluorescência TMRM antes e após a aplicação de um estímulo específico. Mostramos também a aplicação da sonda de fluorescência H<sub&gt; 2</sub&gt; DCF-DA para avaliar o nível relativo de espécies reativas de oxigênio nos neurônios corticais.

Determinação de Membrana Mitocondrial Espécies de Oxigênio Reativas Potencial e em neurônios de rato vivo Cortical

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

determination-mitochondrial-membrane-potential-reactive-oxygen

Research

JoVE Journal

Neuroscience

64.7K visualizações.  Loyola University Chicago. Nós demonstramos a aplicação do indicador de fluorescência, TMRM, em neurônios corticais para determinar a alterações relativas na intensidade de fluorescência TMRM antes e após a aplicação de um estímulo específico. Mostramos também a aplicação da sonda de fluorescência H 2 DCF-DA para avaliar o nível relativo de espécies reativas de oxigênio nos neurônios corticais.

Vídeo: Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

Neuromodulação e Transportes mitocondrial: Imagens ao vivo em neurônios do hipocampo durante muito tempo

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

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 ...

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or co-culture model. This system is suitable for a variety of experimental needs requiring either a glass or plastic growth substrate and can also be used for culture of other types of neurons.
Rat cortical neurons obtained from the late embryonic stage (E17) are plated on glass coverslips or tissue culture dishes facing a feeder layer of glia grown on dishes or plastic coverslips (known as Thermanox), respectively. The choice between the two configurations depends on the specific experimental technique used, which may require, or not, that neurons are grown on glass (e.g. calcium imaging versus Western blot). The glial feeder layer, an astroglia-enriched secondary culture of mixed glia, is separately prepared from the cortices of newborn rat pups (P2-4) prior to the neuronal dissection.
A major advantage of this culture system as compared to a culture of neurons only is the support of neuronal growth, survival, and differentiation provided by trophic factors secreted from the glial feeder layer, which more accurately resembles the brain environment in vivo. Furthermore, the co-culture can be used to study neuronal-glial interactions1.
At the same time, glia contamination in the neuronal layer is prevented by different means (low density culture, addition of mitotic inhibitors, lack of serum and use of optimized culture medium) leading to a virtually pure neuronal layer, comparable to other established methods1-3. Neurons can be easily separated from the glial layer at any time during culture and used for different experimental applications ranging from electrophysiology4, cellular and molecular biology5-8, biochemistry5, imaging and microscopy4,6,7,9,10. The primary neurons extend axons and dendrites to form functional synapses11, a process which is not observed in neuronal cell lines, although some cell lines do extend processes.
A detailed protocol of culturing rat hippocampal neurons using this co-culture system has been described previously4,12,13. Here we detail a modified protocol suited for cortical neurons. As approximately 20x106 cells are recovered from each rat embryo, this method is particularly useful for experiments requiring large numbers of neurons (but not concerned about a highly homogenous neuronal population). The preparation of neurons and glia needs to be planned in a time-specific manner. We will provide the step-by-step protocol for culturing rat cortical neurons as well as culturing glial cells to support the neurons.

Here we provide a protocol for culturing rat cortical neurons in the presence of a glial feeder layer. The cultured neurons establish polarity and create synapses, and can be separated from the glia for use in various applications, such as electrophysiology, calcium imaging, cell survival assays, immunocytochemistry, and RNA/DNA/protein isolation.

Bilaminar Co-cultura de neurônios de rato Primária Cortical e Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

Vivo em dois fótons de imagem de experiência dependentes de alterações moleculares em neurônios corticais

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Membrana Potencial Dye imagem de Ventromedial Hipotálamo Neurônios de camundongos adultos para estudar Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Quantificação de filamentosas actina (F-actina) puncta no Rat neurônios corticais

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Na Vivo Imagem de dois fotões de neurônios Cortical em camundongos Neonatais

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using 128-channel high-density electroencephalography (EEG). The specific aim of the described stimuli and analyses is to examine whether it is feasible to replicate previously reported CVEP morphological patterns elicited by an apparent motion stimulus, designed to simultaneously stimulate both ventral and dorsal central visual networks, using object and motion stimuli designed to separately stimulate ventral and dorsal visual cortical networks.&#160; Four visual paradigms are presented: 1. Randomized visual objects with consistent temporal presentation. 2. Randomized visual objects with inconsistent temporal presentation (or jitter).&#160; 3. Visual motion via a radial field of coherent central dot motion without jitter.&#160; 4. Visual motion via a radial field of coherent central dot motion with jitter.&#160; These four paradigms are presented in a pseudo-randomized order for each participant.&#160; Jitter is introduced in order to view how possible anticipatory-related effects may affect the morphology of the object-onset and motion-onset CVEP response.&#160; EEG data analyses are described in detail, including steps of data exportation from and importation to signal processing platforms, bad channel identification&#160;and removal, artifact rejection, averaging, and categorization of average CVEP morphological pattern type based upon latency ranges of component peaks. Representative data show that the methodological approach is indeed sensitive in eliciting differential object-onset and motion-onset CVEP morphological patterns and may, therefore, be useful in addressing the larger research aim. Given the high temporal resolution of EEG and the possible application of high-density EEG in source localization analyses, this protocol is ideal for the investigation of distinct CVEP morphological patterns and the underlying neural mechanisms which generate these differential responses.&#160; &#160; &#160;&#160;

In this paper, we present a protocol to investigate differential cortical visual evoked potential morphological patterns through stimulation of ventral and dorsal networks using high-density EEG. Visual object and motion stimulus paradigms, with and without temporal jitter, are described. Visual evoked potential morphological analyses are also outlined.&#160;&#160;

Padrões morfológicos do potencial evocado cortical Visual específico do estímulo

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using ...

Generation of Neurospheres from Mixed Primary Hippocampal and Cortical Neurons Isolated from E14-E16 Sprague Dawley Rat Embryo

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have a robust in vitro model that can be exploited for various neurobiology studies. Though primary neuron cultures (i.e., long-term hippocampal cultures) have provided scientists with models, it does not yet represent the complexity of brain network completely. In the wake of these limitations, a new model has emerged using neurospheres, which bears a closer resemblance to the brain tissue. The present protocol describes the plating of high and low densities of mixed cortical and hippocampal neurons isolated from the embryo of embryonic day 14-16 Sprague Dawley rats. This allows for the generation of neurospheres and long-term primary neuron culture as two independent platforms to conduct further studies. This process is extremely simple and cost-effective, as it minimizes several steps and reagents previously deemed essential for neuron culture. This is a robust protocol with minimal requirements that can be performed with achievable results and further used for a diversity of studies related to neuroscience.

Presented here is a protocol for the spontaneous generation of neurospheres enriched in neural progenitor cells from high density plated neurons. During the same experiment, when neurons are plated at a lower density, the protocol also results in prolonged primary rat neuron cultures.

Geração de neuroesferas do hipocampal preliminar misturado e dos neurônios corticais isolados de E14-E16 embrião do rato de Sprague Dawley

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have ...

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. In addition, photoreceptors also exhibit high aerobic glycolysis like cancer cells. Precise measurements of these metabolic activities can provide valuable insights into cellular homeostasis under physiological conditions and in disease states. High throughput microplate-based assays have been developed to measure mitochondrial respiration and various metabolic activities in live cells. However, a vast majority of these are developed for cultured cells and have not been optimized for intact tissue samples and for application ex vivo. Described here is a detailed step-by-step protocol, using microplate-based fluorescence technology, to directly measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration, as well as extracellular acidification rate (ECAR) as an indicator of glycolysis, in intact ex vivo retinal tissue. This method has been used to successfully assess metabolic activities in adult mouse retina and demonstrate its application in investigating cellular mechanisms of aging and disease.

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Determinação da respiração mitocondrial e glicolise em amostras de tecido retinal ex vivo

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. ...