Este trabalho foi financiado pela Agência de Pesquisa esloveno com o projeto Z2-9229 e um programa de P2-0249. Este vídeo representa o material suplementar para o &quot;Eletroporação baseado em Tecnologias e Tratamentos&quot; workshop científico e pós-graduação, organizado a cada dois anos pela Faculdade de Engenharia Elétrica da Universidade de Ljubljana, na Eslovénia.

Parte I: Etapas preliminares <ol><li> Neste experimento de hamster chinês linhagem de células de ovário (CHO-K1) é usado. Células são banhados em câmaras II Lab-Tek (2 poços, 4 cm 2 cada) (Nalge Nunc, Alemanha) em ~ 0.7x10 5 células / ml no HAM-F12 meio de cultura suplementado com 8% de soro fetal bovino, 0,15 mg / ml L-glutamina, 16 mg / ml de gentamicina (todos da Sigma-Aldrich, Steinheim, Alemanha), e 200 unidades / ml Crystacillin (Pliva, Zagreb, Croácia), e incubados em 5% de CO 2 a 37 ° C. Alternativamente, as células também podem ser semeadas em # 1 entra tampa de vidro (0,13-0,16 mm de espessura), revestida com um adesivo celulares como polilisina. </li><li> Incubar as células no seu meio de cultura. Incubação durando 2-4 rendimentos horas células que ainda estão mais ou menos esférica, mas firmemente ligada à superfície com uma pequena parte de sua membrana. Alternativamente, após 16-20 horas de incubação, as células são totalmente ligada à superfície e têm formas mais complexas, mas a maioria deles ainda não estão se dividindo. </li><li> Prepare uma solução estoque 10 mM de di-8-ANEPPS (Invitrogen, Eugene, Oregon, EUA), adicionando 843 mL de DMSO (Sigma-Aldrich, Steinheim, Alemanha) a 5 mg do corante no frasco Invitrogen original. A solução de reserva podem ser armazenadas no frigorífico a 4 ° C durante vários meses. Antes de iniciar os experimentos, aquecer a solução até que os cristais de DMSO dissolver. </li><li> Algumas linhas de celular podem exigir o uso de PLURONIC para facilitar a incorporação de corante na membrana celular. PLURONIC pode ser comprado em solução estoque de 20% em DMSO (F-127, Invitrogen, Eugene, Oregon, EUA), ou uma solução-mãe da mesma concentração pode ser preparada dissolvendo PLURONIC em DMSO. Solução estoque de PLURONIC pode ser armazenado em temperatura ambiente. </li></ol> Parte II: Carregando as células com di-8-ANEPPS <ol><li> Misture 3 mL de 10 mM di-8-ANEPPS e 2,5 mL de PLURONIC 20% em 1 ml da modificação Spinner (cálcio depleção version) do smem Médio Mínimo Essencial (médio M8167 ou M4767, Sigma-Aldrich, Steinheim, Alemanha ) em um tubo Eppendorf 1,5 ml. Isso gera uma &quot;solução de carga&quot;, contendo aproximadamente 30 mM di-8-ANEPPS e 0,05% de PLURONIC. Para outros tipos celulares, seus meios de comunicação cultura nativa pode ser usado em vez de smem. </li><li> Substituir o meio de cultura na câmara Lab-Tek com a solução de carga. Transferência da câmara para a geladeira por 10 min a 4 ° C. A esta temperatura, a internalização do corante através da membrana plasmática é em grande parte inibida. </li><li> Após a coloração, lavar cuidadosamente o excesso de corante 2-3 vezes com smem puro. </li><li> Deixe 1,5 ml de smem na câmara. </li></ol> Parte III: Experimento e de aquisição de imagem <ol><li> As células são observados usando um microscópio de fluorescência (no nosso caso Zeiss Axiovert 200, Zeiss, Alemanha) equipado com um objetivo de imersão em óleo (X63, NA 1.4), uma VisiCam monocromador (Polychrome IV, Visitron, Alemanha) e uma câmera CCD refrigerado ( 1280, Visitron, Alemanha). As imagens são adquiridas com MetaFluor 7.1.1 e processadas em Metamorph 7.1.1 (ambos dispositivos moleculares, Downingtown, PA, EUA), mas o software de aquisição semelhante também pode ser usado. Definir o comprimento de onda de excitação no software de aquisição de 490 nm e escolha um filtro de emissão adequada para ANEPPS, por exemplo, um filtro passa-banda centrado em nm 605 (605/55m, dicróicas 565 DCXR, Chroma, Rockingham, VT, EUA). Se um monocromador não estiver disponível, um filtro de excitação centrado em 490 nm pode ser usado em seu lugar. </li><li> Coloque a câmara com as células para o palco microscópio, a posição dos eletrodos na parte inferior da câmara e conectá-los ao gerador de pulsos. O meio deve cobrir os eletrodos. </li><li> Para manter a viabilidade das células e reduzir o aquecimento, o pulso deve ser de amplitude suficientemente baixa e de curta duração. Neste experimento um pulso quadrado com 35 V de amplitude e 50 ms de duração (baixa o suficiente para evitar a eletroporação) é gerada utilizando uma fonte de tensão DC e uma custom-made controlado por microprocessador dispositivo switcher. Alternativamente, um gerador de pulso comercial, tais como 33210A (Agilent, Santa Clara, CA, EUA) conectado a um amplificador ou um estimulador de comerciais, como S88 (Grass, West Warwick, RI, EUA), pode ser usado. O pulso é entregue a duas paralelas eletrodos Pt / Ir fio com 0,8 mm de diâmetro e 4 mm de distância entre eles, criando um campo elétrico de aproximadamente 88 V / cm entre os eletrodos e ΔΦ indução. A distribuição exata de campo entre os eletrodos utilizados neste experimento é descrito em outros lugares [1]. </li><li> Encontrar as células de interesse. Aplicar um único pulso elétrico ou uma seqüência de pulsos. Para cada pulso, adquirir duas imagens de fluorescência: uma imediatamente antes do pulso (a imagem de controle), e uma durante o pulso (a imagem de pulso). Devido à baixa resposta de di-8-ANEPPS, as mudanças na fluorescência são difíceis de discernir a olho nu e se tornar aparente only após o processamento. A entrega de pulso deve ser sincronizada com a aquisição de imagem, que é uma característica do software de aquisição. Para evitar a fotodegradação e possível aquecimento, a iluminação pode ser limitado à duração do pulso. </li></ol> Parte IV: Processamento de imagem e análise <ol><li> Abra as imagens no software Metamorph. Para cada pulso, subtrair o fundo em ambos o controle e a imagem de pulso. </li><li> Escolha uma célula e definir a região de interesse para que ele corresponde à membrana. Medir a intensidade de fluorescência ao longo desta região na imagem e controle de pulso e transferir os valores para uma planilha. </li><li> Para cada pulso, subtrair os dados de controle a partir dos dados de pulso, e dividir o resultado por dados de controle para obter as alterações em relação fluorescência. Se uma seqüência de pulsos é aplicada, os valores de fluorescência em relação as mudanças determinadas para cada pulso pode ser a média para obter uma medição mais confiável. </li><li> Transformar as mudanças em relação fluorescência em ΔΦ usando uma curva de calibração. Uma estimativa grosseira da curva pode ser obtida a partir da literatura, mas para maior precisão, tem que ser medidos para cada configuração específica. Para as células e da configuração utilizados neste experimento uma diminuição de 6% na fluorescência corresponde a um aumento de 100 mV em ΔΦ [2]. </li><li> Finalmente, plotar a tensão em função do comprimento do arco relativo em um pacote de software de gráficos como o Excel (Microsoft Corp, Redmond, WA, EUA), Sigma Plot (Systat Software Inc., San Jose, CA, EUA), ou Origem (Origin Corp, Northampton, MA, EUA). A curva também pode ser suavizada com um filtro adequado, tais como o filtro de média móvel. </li></ol>

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Eng. 34, 642-652 (2006).</li><li>Huang, C. J., Harootunian, A., Maher, M. P., Quan, C., Raj, C. D., McCormack, K., Numann, R., Negulescu, P. A., Gonzalez, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+voltage-gated+sodium-channel+blockers+by+electrical+stimulation+and+fluorescence+detection+of+membrane+potential.">Characterization of voltage-gated sodium-channel blockers by electrical stimulation and fluorescence detection of membrane potential.</a> Nat. Biotechnol. 24, 439-446 (2006).</li><li>Bedlack, R. S., Wei, M., Fox, S. H., Gross, E., Loew, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+electric+potentials+in+soma+and+neurite+membranes.">Distinct electric potentials in soma and neurite membranes.</a> Neuron. 13, 1187-1193 (1994).</li><li>Cheng, D. K. 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Medições da membrana induzidas (transmembrana) de tensão, ΔΦ, pode ser importante em várias situações experimentais, tais como estudos de canais dependentes da voltagem da membrana, a propagação do potencial de ação, a estimulação de células cardíacas, ou eletroporação da membrana da célula [3, 4, 5, 6 , 7]. Com formas simples célula, ΔΦ pode ser calculado analiticamente. Por exemplo, para uma célula esférica, ΔΦ é dada pela equação a Schwan s, que afirma que a tensão é proporcional à intensidade do campo e do tamanho das células e segue a função cosseno ao longo da membrana [8, 9]. Para mais formas de células complicado, ΔΦ pode se desviar consideravelmente do cosseno e deve ser determinado numericamente, usando um computador [2, 10, 11], ou experimentalmente, utilizando um corante potenciométrica [12, 13, 14, 15]. Um dos corantes potenciométrica amplamente utilizado para esta finalidade é di-8-ANEPPS (di-8-butil-amino-naftil-etileno-piridínio-propil-sulfonato), um corante rápido com espectros de excitação e emissão dependente da tensão de membrana, não-invasivo que permite observações das variações do ΔΦ na membrana celular e medir o seu valor. Neste vídeo, vamos mostrar uma abordagem experimental para a determinação de ΔΦ usando di-8-ANEPPS. O corante foi desenvolvido pelo Professor Leslie Loew e colegas [13, 14] na Universidade de Connecticut e pertence à classe dos corantes de resposta rápida. di-8-ANEPPS é nonfluorescent em água e torna-se fortemente fluorescente quando se incorpora na bicamada lipídica da membrana celular. Uma mudança no resultado ΔΦ em uma mudança da distribuição de carga intramolecular e alterações correspondentes no perfil espectral e intensidade de fluorescência do corante. A intensidade de fluorescência de di-8-ANEPPS varia proporcionalmente à mudança de ΔΦ; a resposta do corante é linear para tensões variando de -280 mV a +250 mV [4, 16]. Mudanças relativamente pequenas na fluorescência do corante coloração da membrana, desigual, e internalização corante fazer di-8-ANEPPS menos adequado para medições absolutas de tensão de membrana, por exemplo, a sua componente de repouso, embora esses esforços também foram relatados [17]. É, no entanto, adequado para medir as mudanças maiores na tensão de membrana, tais como o início da tensão da membrana induzidas em células não excitáveis ​​expostos a campos elétricos externos [12, 13], ou potenciais de ação em células excitáveis ​​[4, 5]. Embora não seja aplicado aqui, di-8-ANEPPS também permite a determinação do ΔΦ por medições raciométrica de excitação de fluorescência [18] ou de emissão [19], o que aumenta a sensibilidade da resposta e reduz os efeitos acima referidos. Como di-8-ANEPPS manchas da membrana, ela também pode ser usado simplesmente como um marcador de membrana [2]. Um dos inconvenientes do corante é que ele está propenso a fotodegradação, de modo que a exposição prolongada à luz forte devem ser evitados. Calibração do corante é feita com um valinomicina ionóforo (i) de potássio e um conjunto de diferentes concentrações de potássio no meio externo [2,18], ou (ii) patch-clamp clamp no modo de tensão [17]. Finalmente, com as medidas de ΔΦ em células esféricas e células de formas mais complexas, o vídeo demonstra a influência da forma da célula sobre a distribuição espacial da amplitude e ΔΦ. Assim, para células esféricas ΔΦ está perto de uma co-seno, de acordo com a equação Schwan, enquanto para formas mais complicadas de células a distribuição espacial da ΔΦ é mais complexa [20] ...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>di-8-ANEPPS</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>D-3167</td>
<td>potentiometric fluorescent dye</td>
</tr>
<tr>
<td>pluronic</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>P3000MP</td>
<td>potassium ionophore</td>
</tr>
<tr>
<td>DMSO</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>D2650</td>
<td>&nbsp;</td>
<tr>
<td>SMEM</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>M8167 or M4767</td>
<td>Spinner modification of the Minimum Essential Medium</td>
</tr>
<tr>
<td>Ham-F12</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>N4888</td>
<td>culture medium</td>
</tr>
<tr>
<td>fetal calf serum</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>F4135</td>
<td>&nbsp;</td>
<tr>
<td>L-glutamine</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>G7513</td>
<td>&nbsp;</td>
<tr>
<td>crystacillin</td>
<td>&nbsp;</td>
<td>Pliva</td>
<td>625110</td>
<td>antibiotic</td>
</tr>
<tr>
<td>gentamicin</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>G1397</td>
<td>antibiotic</td>
</tr>
<tr>
<td>Lab-Tek II</td>
<td>&nbsp;</td>
<td>Nalge Nunc</td>
<td>155379</td>
<td>chamber</td>
</tr>
<tr>
<td>DC voltage supply</td>
<td>&nbsp;</td>
<td>Elektro-Automatik</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>microprocessor-controlled switcher</td>
<td>&nbsp;</td>
<td>Custom made</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>electrodes</td>
<td>&nbsp;</td>
<td>Custom made</td>
<td>&nbsp;</td>
<td>Pt/Ir</td>
</tr>		</tbody>
		</table>
		</div>

measuring the induced membrane voltage with di-8-anepps

Colocação de uma célula em um campo elétrico externo causa uma redistribuição no interior locais de carga e fora da célula nas proximidades da membrana celular, resultando em uma tensão através da membrana. Esta tensão, denominado tensão de membrana induzida (tensão induzida transmembrana também, ou diferença de potencial transmembrana induzido) e denotado por ΔΦ, existe apenas enquanto o campo externo está presente. Se a tensão de repouso está presente na membrana, a tensão induzida sobrepõe (adiciona) para ele. Usando um dos corantes fluorescentes potenciométrica, como di-8-ANEPPS, é possível observar as variações de ΔΦ na membrana celular e medir o seu valor de forma não invasiva. di-8-ANEPPS torna-se fortemente fluorescente quando vinculado à bicamada lipídica da membrana celular, com a mudança da intensidade de fluorescência proporcional à mudança de ΔΦ. Este vídeo mostra o protocolo para a medição ΔΦ utilizando di-8-ANEPPS e também demonstra a influência da forma da célula sobre a amplitude ea distribuição espacial dos ΔΦ.

Watch this Scientific Journal Video about Measuring the Induced Membrane Voltage with Di-8-ANEPPS at JoVE.com

Measuring the Induced Membrane Voltage with Di-8-ANEPPS

Campo elétrico externo induz uma tensão na membrana de uma célula, chamado de tensão de membrana induzida (ΔΦ). Usando o corante potenciométrica di-8-ANEPPS, é possível medir a ΔΦ não invasiva. Este vídeo mostra o protocolo para a medição ΔΦ utilizando di-8-ANEPPS.

Medir a tensão de membrana induzida com Di-8-ANEPPS

Placement of a cell into an external electric field causes a local charge redistribution inside and outside of the cell in the vicinity of the cell ...

measuring-the-induced-membrane-voltage-with-di-8-anepps

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JoVE Journal

Biology

17.2K visualizações.  University of Ljubljana. Campo elétrico externo induz uma tensão na membrana de uma célula, chamado de tensão de membrana induzida (ΔΦ). Usando o corante potenciométrica di-8-ANEPPS, é possível medir a ΔΦ não invasiva. Este vídeo mostra o protocolo para a medição ΔΦ utilizando di-8-ANEPPS.

Vídeo: Measuring the Induced Membrane Voltage with Di-8-ANEPPS

Measuring Near Plasma Membrane and Global Intracellular Calcium Dynamics in Astrocytes

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, increasing evidence suggests that astrocytes may also actively participate in brain function through functional interactions with neurons.  However, many fundamental aspects of astrocyte biology remain controversial, unclear and/or experimentally unexplored. One important issue is the dynamics of intracellular calcium transients in astrocytes. This is relevant because calcium is well established as an important second messenger and because it has been proposed that astrocyte calcium elevations can trigger the release of transmitters from astrocytes. However, there has not been any detailed or satisfying description of near plasma membrane calcium signaling in astrocytes. Total internal reflection fluorescence (TIRF) microscopy is a powerful tool to analyze physiologically relevant signaling events within about 100 nm of the plasma membrane of live cells. Here, we use TIRF microscopy and describe how to monitor near plasma membrane and global intracellular calcium dynamics almost simultaneously. The further refinement and systematic application of this approach has the potential to inform about the precise details of astrocyte calcium signaling. A detailed understanding of astrocyte calcium dynamics may provide a basis to understand if, how, when and why astrocytes and neurons undergo calcium-dependent functional interactions.

We describe how to measure near membrane and global intracellular calcium dynamics in cultured astrocytes using total internal reflection and epifluorescence microscopy.

Medição Perto membrana plasmática e global Dynamics cálcio intracelular em astrócitos

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, ...

Biologia

Measuring Plasma Membrane Protein Endocytic Rates by Reversible Biotinylation

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins is responsible for a variety of key cellular events, including nutrient delivery, cellular excitability, and chemical signaling.  Many plasma membrane proteins are dynamically regulated by endocytic trafficking, which modulates protein function by altering protein surface expression.  The mechanisms that facilitate protein endocytosis are complex and are not fully understood for many membrane proteins.  In order to fully understand the mechanisms that control the endocytic trafficking of a given protein, it is critical that the protein s endocytic rate be precisely measured.  For many receptors, direct endocytic rate measurements are frequently achieved utilizing labeled receptor ligands.  However, for many classes of membrane proteins, such as transporters, pumps and ion channels, there is no convenient ligand that can be used to measure the endocytic rate.  In the present report, we describe a reversible biotinylation method that we employ to measure the dopamine transporter (DAT) endocytic rate.  This method provides a straightforward approach to measuring internalization rates, and can be easily employed for trafficking studies of most membrane proteins.

Regulated endocytosis governs the cell surface expression levels of the majority of membrane proteins. Here we utilize reducible, membrane impermeant biotinylation reagents to measure the endocytic rate of the dopamine transporter (DAT), a polytopic membrane protein. The method facilitates a straightforward approach to measuring the endocytic rate of most plasma membrane proteins.

Taxas de medição de plasma Proteína de membrana endocítica por Biotinilação reversível

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins ...

Measuring Caenorhabditis elegans Life Span in 96 Well Microtiter Plates

Lifespan is a biological process regulated by several genetic pathways. One strategy to investigate the biology of aging is to study animals that harbor mutations in components of age-regulatory pathways. If these mutations perturb the function of the age-regulatory pathway and therefore alter the lifespan of the entire organism, they provide important mechanistic insights1-3.
Another strategy to investigate the regulation of lifespan is to use small molecules to perturb age-regulatory pathways. To date, a number of molecules are known to extend lifespan in various model organisms and are used as tools to study the biology of aging4-16. The number of molecules identified thus far is small compared to the genetic "toolset" that is available to study the biology of aging.
Caenorhabditis elegans is one of the principle models used to study aging because of its excellent genetics and short lifespan of three weeks. More recently, C.elegans has emerged as a model organism for phenotype based drug screens5,7,16-20 because of its small size and its ability to grow in microtiter plates.
Here we present an assay to measure C.elegans lifespan in 96 well microtiter plates. The assay was developed and successfully used to screen large libraries for molecules that extend C.elegans lifespan7. The reliability of the assay was evaluated in multiple tests: first, by measuring the lifespan of wild type animals grown at different temperatures; second, by measuring the lifespan of mutants with altered lifespans; third, by measuring changes in lifespan in response to different concentrations of the antidepressant Mirtazepine. Mirtazepine has previously been shown to extend lifespan in C.elegans7. The results of these tests show that the assay is able to replicate previous findings from other assays and is quantitative. The microtiter format also makes this lifespan assay compatible with automated liquid handling systems and allows integration into automated platforms.

Measuring Caenorhabditis elegans Life Span in 96 Well Microtiter Plates

In this protocol we present a method to measure Caenorhabditis elegans lifespan in 96 well microtiter plates.

Medição Caenorhabditis elegans Expectativa de Vida em 96 placas de microtitulação Bem

Lifespan is a biological process regulated by several genetic pathways. One strategy to investigate the biology of aging is to study animals that harbor ...

Crystallization of Membrane Proteins in Lipidic Mesophases

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are implicated in a multitude of malfunctions and diseases. However, a structural and functional understanding of membrane proteins is strongly lagging behind that of their soluble partners, mainly, due to difficulties associated with their solubilization and generation of diffraction quality crystals. Crystallization in lipidic mesophases (also known as in meso or LCP crystallization) is a promising technique which was successfully applied to obtain high resolution structures of microbial rhodopsins, photosynthetic proteins, outer membrane beta barrels and G protein-coupled receptors. In meso crystallization takes advantage of a native-like membrane environment and typically produces crystals with lower solvent content and better ordering as compared to traditional crystallization from detergent solutions. The method is not difficult, but requires an understanding of lipid phase behavior and practice in handling viscous mesophase materials. Here we demonstrate a simple and efficient way of making LCP and reconstituting a membrane protein in the lipid bilayer of LCP using a syringe mixer, followed by dispensing nanoliter portions of LCP into an assay or crystallization plate, conducting pre-crystallization assays and harvesting crystals from the LCP matrix. These protocols provide a basic guide for approaching in meso crystallization trials; however, as with any crystallization experiment, extensive screening and optimization are required, and a successful outcome is not necessarily guaranteed.

The protocols describe the essential steps for obtaining diffraction quality crystals of a membrane protein starting from reconstitution of the protein in a lipidic cubic phase (LCP), finding initial conditions with LCP-FRAP pre-crystallization assays, setting up LCP crystallization trials and harvesting crystals.

Cristalização de proteínas da membrana lipídica em mesofases

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Examinando o Dynamics Conformacional de Proteínas de Membrana In situ Com Labeling-Site dirigido Fluorescência

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion channels in oocytes. Recordings from the cut-open setup are particularly useful for resolving low magnitude gating currents, rapid ionic current activation, and deactivation. The main benefits over the two-electrode voltage clamp (TEVC) technique include increased clamp speed, improved signal-to-noise ratio, and the ability to modulate the intracellular and extracellular milieu.
Here, we employ the human cardiac sodium channel (hNaV1.5), expressed in Xenopus oocytes, to demonstrate the cut-open setup and protocol as well as modifications that are required to add voltage clamp fluorometry capability.
The properties of fast activating ion channels, such as hNaV1.5, cannot be fully resolved near room temperature using TEVC, in which the entirety of the oocyte membrane is clamped, making voltage control difficult. However, in the cut-open technique, isolation of only a small portion of the cell membrane allows for the rapid clamping required to accurately record fast kinetics while preventing channel run-down associated with patch clamp techniques.
In conjunction with the COVG technique, ion channel kinetics and electrophysiological properties can be further assayed by using voltage clamp fluorometry, where protein motion is tracked via cysteine conjugation of extracellularly applied fluorophores, insertion of genetically encoded fluorescent proteins, or the incorporation of unnatural amino acids into the region of interest1. This additional data yields kinetic information about voltage-dependent conformational rearrangements of the protein via changes in the microenvironment surrounding the fluorescent molecule.

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open Vaseline gap approach is used to obtain low noise recordings of ionic and gating currents from voltage-dependent ion channels expressed in Xenopus oocytes with high resolution of fast channel kinetics. With minor modification, voltage clamp fluorometry can be coupled to the cut-open oocyte protocol.

O<em&gt; Xenopus</em&gt; Oocyte Cut-aberto vaselina Gap Técnica Voltage-clamp Com Fluorometria

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion ...

Imaging Plasma Membrane Deformations With pTIRFM

To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during the fusion of vesicle and plasma membranes. These rapid and localized changes are achieved by dynamic interactions between lipids and specialized proteins that control membrane curvature. The absence of such interactions would not only have devastating consequences for vesicle fusion, but a host of other cellular functions that involve control of membrane shape. In recent years, the identity of a number of proteins with membrane-shaping properties has been determined. What remains missing is a roadmap of when, where, and how they act as fusion and content release progress.
Our understanding of the molecular events that enable membrane remodeling has historically been limited by a lack of analytical methods that are sensitive to membrane curvature or have the temporal resolution to track rapid changes. PTIRFM satisfies both of these criteria. We discuss how pTIRFM is implemented to visualize and interpret rapid, submicron changes in the orientation of chromaffin cell membranes during dense core vesicle (DCV) fusion. The chromaffin cells we use are isolated from bovine adrenal glands. The membrane is stained with a lipophilic carbocyanine dye,1,1&#39;-dioctadecyl-3,3,3&#39;,3&#39;-tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate, or&#160;diD. DiD intercalates in the membrane plane with a &#34;fixed&#34; orientation and is therefore sensitive to the polarization of the evanescent field. The diD-stained cell membrane is sequentially excited with orthogonal polarizations of a 561&#160;nm laser (p-pol, s-pol). A 488&#160;nm laser is used to visualize vesicle constituents and time the moment of fusion. Exocytosis is triggered by locally perfusing cells with a depolarizing KCl solution. Analysis is performed offline using custom-written software to understand how diD emission intensity changes relate to fusion pore dilation.

Polarization-based Total Internal Reflection Fluorescence Microscopy (pTIRFM) enables real-time detection of cell membrane dynamics. This article describes the implementation of pTIRFM for the study of membrane remodeling during regulated exocytosis. The technique is generalizable to other processes in cell biology that directly or indirectly involve changes in membrane shape.

Imagem Plasma Membrane Deformações Com pTIRFM

To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during ...

Measuring Glutathione-induced Feeding Response in Hydra

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. The movement of prey organisms causes mechanosensory discharge of the stinging cells called nematocysts from hydra, which are inserted into the prey. The feeding response in hydra, which includes curling of the tentacles to bring the prey towards the mouth, opening of the mouth and consequent engulfing of the prey, is triggered by GSH present in the fluid released from the injured prey. To be able to identify the molecular mechanism of the feeding response in hydra which is unknown to date, it is necessary to establish an assay to measure the feeding response. Here, we describe a simple method for the quantitation of the feeding response in which the distance between the apical end of the tentacle and mouth of hydra is measured and the ratio of such distance before and after the addition of GSH is determined. The ratio, called the relative tentacle spread, was found to give a measure of the feeding response. This assay was validated using a starvation model in which starved hydra show an enhanced feeding response in comparison with daily fed hydra.

Here we describe a simple assay for the quantification of the feeding response in hydra induced by the reduced form of glutathione. This assay relies on measuring the distance between the apical end of the tentacle and mouth of hydra.

Medindo alimentação resposta induzida por glutationa em Hydra

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

Microperf Técnica para Investigar Regulamento de microvasos Permeabilidade de Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

Estado fundamental depleção Super-resolução Imaging em células de mamíferos

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...