KB é suportado pelo NIH conceder GM54544.

Parte 1: Equipamentos e Suprimentos <ol><li> Um microscópio de dissecção bom é necessário para dissecção embrião; aumento de 40 vezes é sugerido, com 25X de olhos pedaços ao máximo aumentar a ampliação. </li><li> Forceps multa (número 5) são necessários para a seleção manual de embriões e devitellinization de embriões. </li><li> Equipamentos para fazer e modificar agulhas finas de vidro é necessário. Puxado vidro agulhas são necessários para a dissecção. Nós preferimos vidro sólido para dissecção (duram mais), mas tubos de vidro padrão de paredes espessas (diâmetro externo 1-1,5 mm) funciona tão bem. Algumas pessoas usam agulhas de tungstênio fino, eletrolítica para a nitidez desejada em um banho de 1M NaOH utilizando uma bateria de 10 + autotransformador Amp. Agulhas dissecção afiada de tungstênio têm o benefício que eles são relativamente resistentes e de longa duração. Agulhas de vidro oco (formada semelhante a correção eletrodos grampo) estão ligados a tubos de plástico e usado durante a dissecção para sucção e expulsão de solução salina, bem como a liberação controlada de cola. Uma variedade de puxadores de vidro estão disponíveis para a fabricação de agulhas de vidro. Computador programado extratores podem ser predefinidos para puxar uma variedade de formas e tamanhos. A Brown-Flaming modelos (Sutter Instrument Co.) são amplamente favorecidos. Um microscópio composto (20-40X objetivo) deve estar disponível para inspecionar e modificar eletrodos antes do uso. </li><li> Dois tipos de soluções são comumente usados ​​para o banho de tecidos embrionários durante a dissecção: 1) &quot;standard&quot; (Jan e Jan, 1976a) ou &quot;padrão modificado&quot; (Broadie, 2000) salinas, com base em soluções comumente utilizadas para a gravação em outros sistemas de invertebrados e 2) &quot;hemolinfa-like&quot; (HL) salinas (Stewart et al 1994), um compromisso entre a solução salina padrão e as concentrações iónicas medidas na hemolinfa Drosophila. Note-se que nenhuma dessas salinas imitar com precisão a composição química da hemolinfa in vivo (Broadie, 2000). Padrão de solução salina contém (em mM): 135 NaCl, KCl 5, 4 MgCl 2, CaCl 2 1,8, 72 de sacarose, 5 TES, pH 7.2. Experimentadores também pode querer considerar salina inseto comercialmente preparado (Media Insect por exemplo Schneider), que é inadequado para a gravação eletrofisiológicas mas muito provavelmente para proteger a saúde dos tecidos expostos. </li></ol> Parte 2: Estadiamento Embrião e Dissection <ol><li> Para coleta de ovos ideal, manter jovens (&lt;7 dias), do sexo masculino saudáveis ​​e moscas fêmeas (20-40) em potes de doce, que por 2-3 dias. Deitado potes comumente consistem em copos de 100 ml de plástico (Tri-Pour) perfurada para proporcionar a circulação de ar adequada, cobrindo uma placa de agar 60 mm. Placas de ágar contém maçã ou suco de uva temperado com agar. Existem várias receitas, mas um exemplo é a seguinte: 700 ml de água, 25-30 agar gramas, 300 ml de suco concentrado (uva ou maçã), 0,5 g metil paraben (p-hydroxymethylbenzoate), e 30g de açúcar. Autoclave da água e ágar, e, separadamente, ferver a p-hydroxymethylbenzoate com o açúcar e suco. Misture o suco de agar e soluções e rapidamente despeje em placas de 60 mm, removendo as bolhas. </li><li> Antes da coleta de ovos, as moscas são alimentados com pasta de levedura (7 gramas de fermento de padeiro em 9 ml de água armazenada a 4 ° C) em placas fresca, trocada pelo menos duas vezes por dia durante pelo menos dois dias. De dia 3, um bom vaso deve produzir 100-200 ovos por hora. Postura melhor ovo é observado em um pote mantido em seu lado com o agar na placa riscada. Muitos embriões serão colocados dentro ou ao lado da arranhões. </li><li> Para coletar um grande número de embriões, delicadamente a transferência dos embriões a partir da placa de ágar em uma cesta usando um pincel. A cesta pode ser feita com um tubo de centrífuga de 15 ou 50 ml. Cortar o fundo deixando área de superfície suficiente para prender uma tela (70 mesh mm) entre a tampa ea parte superior do tubo. Enxágüe os embriões com dH 2 0 e colocar em uma placa de Petri de lixívia 50% a 100% fresca para remover o córion exterior. Alternativamente, os ovos podem ser recolhidos individualmente usando uma pinça fina, e dechorionated, colocando-os manualmente em uma gota de água sanitária. De-chorionation pode levar de 30 segundos a dois minutos (lixívia fresca é muito mais rápido), e devem ser monitorizados sob o microscópio de dissecação. Remoção do córion expõe a brilhante, membrana vitelina transparente. Dechorionated ovos deverão ser lavados brevemente, ou colocados em solução salina, como alvejantes destrói rapidamente os tecidos devitellinized. </li><li> É fundamental cuidadosamente estágio embriões para a idade desejada de desenvolvimento. Os estágios definidos Drosophila embrionárias (1-17; Campos-Ortega e Hartenstein, 1985) não são úteis, como todo o desenvolvimento funcional neuromuscular ocorre em 16 fase tardia ou estágio 17. Assim, os embriões são encenadas por hora de desenvolvimento a 25 ° C em uma incubadora umidificada, em horas após a fecundação ou, mais comumente, após a postura de ovos (AEL). Sob essas condições, a embriogênese dura 21 + / - 1 hr a 25 ° C. </li><li> Ovos de uma cronometrado ovo-lay (1-2 hrs) que estão devidamente idosos são vistas próximas de critérios morfológicos. Após a lavagem extensiva em dH 2 O, os ovos dechorionated são colocados em uma placa de cultura de plástico (sob dH 2 O) para visualização. Preparação correta pode ser confirmada com a critérios morfológicos usando luz refletida com um microscópio de dissecção (Campos-Ortega e Hartenstein, 1985). Madura fase final de embriões (22-24h AEL) à beira de incubação são geralmente reconhecidos pela segmentação da cutícula e traquéia inflado. Embriões também podem ser genotipados utilizando GFP balanceadores e um microscópio de fluorescência dissecção com filtros adequados. </li><li> Para dissecção, embriões e dechorionated developmentally encenado estão ligados (lado dorsal para cima) em uma lamela em uma gota de solução salina (veja o passo 1.5). Embriões são retirados da membrana vitelina por punção da membrana perto da extremidade anterior ou posterior com uma micropipeta de vidro ou a agulha de tungstênio, e, em seguida, suavemente libertando o embrião da membrana. Em preparação para a dissecação, o embrião deve ser posicionado sobre a lamela com a superfície até dorsal (face dorsal até dissecção irá expor o sistema nervoso e ventral neuromusculature para experimentos. No entanto, o embrião pode, obviamente, ser posicionados em outras maneiras de facilitar o acesso a outros tecidos.) </li><li> Embriões em estágio precoce (&lt;16 hrs AEL) atribuem directamente para limpar o vidro ou o vidro revestido com polilisina (poli-L-lisina hydrobromide; Broadie e Bate, 1993a, b). Embriões mais velhos (&gt; 16 hrs AEL), após a formação da cutícula, deve ser colado, de preferência para Sylgard revestido (Dow Corning) lamínulas. A água-polimerização cola de cianoacrilato cirúrgicos ou veterinários é usado (Broadie e Bate, 1993c, d). A cola é entregue através de uma pipeta de vidro pequeno eletrodo (10-20 micrômetros ponta diâmetro interno) anexado a um tubo de borracha com o fluxo de cola controlado por pressão na boca. Cuidados devem ser tomados durante a entrega de cola, como a cola polimeriza rapidamente, ao contatar a solução salina. Note também que a colagem não pode ser realizada em solução salina sem ou com baixas concentrações de íons bivalentes, como a cola exige íons divalentes para polimerizar. Pequenas quantidades de cola são utilizados pela primeira vez para anexar firmemente a cabeça ea cauda. </li><li> Uma vez que as extremidades do embrião são colados à lamela, uma incisão é feita no dorso com um eletrodo de vidro ou a agulha de tungstênio afiadas. Isso é facilitado pelo cuidado perfurantes da linha de incisão antes de fazer o corte final. Os lados da incisão são, então, ligado à lamela com mais cola, para delicadamente espalhar o apartamento embrião. Os órgãos internos, incluindo o intestino, órgãos de gordura e, opcionalmente, traquéia são então removidas por sucção usando uma pipeta de vidro ligado a tubos de borracha (Broadie eo Bate 1993a-c). O embrião agora deve ser anexado plana para as lamelas, epiderme para baixo com o sistema central nervoso ventral, nervos periféricos, e somáticas musculatura expostos para a experimentação. </li></ol>

Encenação precisa de embriões é crítica devido à maturação rápida de propriedades funcionais ao longo do tempo de apenas algumas horas. Várias questões complicar este teste. Primeiro, a maioria dos pesquisadores usa ovo cronometrado estabelece a fase de embriões, mas o tempo de postura pode variar enormemente de animal para animal em diferentes condições (Broadie et al. 1992). Em particular, as fêmeas em uma dieta limitada tendem a reter ovos fertilizados por períodos prolongados antes do assentamento. P...

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		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
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		<th>Catalogue Number</th>
		<th>Comment</th>
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<td>Embryo Collection Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific (www.flystuff.com)</td>
<td>59-100 (for 60 mm dish; other sizes available)</td>
<td>Cages can also be home made using punctured tri-pour beakers, as shown in video</td>
</tr>
<tr>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>Available from various companies</td>
<td>Surgical glue adheres better to sylgard-coated coverslips</td>
</tr>
<tr>
<td>Fine glass tubing, outer diameter 1.0-1.5 mm</td>
<td>&nbsp;</td>
<td>various</td>
<td>&nbsp;</td>
<td>For pulling into fine glass needles for dissection and tubes for glue delivery</td>
</tr>
<tr>
<td>Plastic tubing, to attach to glass pipette for mouth suction and glue application</td>
<td>&nbsp;</td>
<td>Tygon</td>
<td>&nbsp;</td>
<td>Tubing inner diameter needs to match glass outer diameter.</td>
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harvesting and preparing drosophila embryos for electrophysiological recording and other procedures

Drosophila é um modelo premier genética para o estudo de ambos o desenvolvimento embrionário e neurociência funcional. Tradicionalmente, esses campos são muito afastadas umas das outras, com histórias em grande parte independentes e comunidades científicas. No entanto, a interface entre esses campos normalmente díspares são os programas de desenvolvimento subjacente aquisição de propriedades funcionais elétricos de sinalização e diferenciação das sinapses químicas funcionais durante as fases finais da formação de circuitos neurais. Esta interface é uma área extremamente importante para a investigação. Em Drosophila, estas fases de desenvolvimento funcional ocorrem durante um período de &lt;8 horas (a 25 ° C) durante o último terço da embriogênese. Este período final de desenvolvimento foi considerado por muito tempo intratáveis ​​devido a investigação para a deposição de um resistente, impermeável cutícula epidérmica. Um avanço de avanço foi a aplicação de cola de água polimerização cirúrgica que pode ser aplicado localmente para a cutícula para que a dissecção controlada de embriões em estágio avançado. Com uma incisão dorsal longitudinal, o embrião pode ser deitadas, expondo o cordão nervoso ventral e musculatura da parede do corpo à investigação experimental. Este sistema tem sido fortemente usada para isolar e caracterizar mutantes genéticos que impedem a formação da sinapse embrionárias e, assim, revelar os mecanismos moleculares que regem a especificação e diferenciação das conexões sinápticas funcionais sinapse e propriedades de sinalização.

Watch this Scientific Journal Video about Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures at JoVE.com

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Esta técnica expõe a Drosophila neuromusculature embrionárias para imuno-histoquímica ou gravação eletrofisiológico. É útil para o estudo de eventos no início do desenvolvimento neuromuscular ou a realização de eletrofisiologia em mutantes que não podem eclodir.

Colheita e Preparação Drosophila Os embriões para Gravação eletrofisiológicos e outros procedimentos

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

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Biology

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

13.3K visualizações.  University of Illinois. Esta técnica expõe a Drosophila neuromusculature embrionárias para imuno-histoquímica ou gravação eletrofisiológico. É útil para o estudo de eventos no início do desenvolvimento neuromuscular ou a realização de eletrofisiologia em mutantes que não podem eclodir.

Vídeo: Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

Preparação de culturas de neurônios a partir de embriões Midgastrula Stage Drosophila

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

Biologia

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

Métodos eletrofisiológicos de gravação Potenciais Synaptic do MNJ de larvas de Drosophila

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Gravação eletrofisiológicas na Drosophila Embrião

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

Preparação de embriões de Microscopia Eletrônica do Drosophila Tubo cardíaco embrionário

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

In vivo Centrifugação de Embriões Drosophila

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Imagem vertical de Drosophila Os embriões

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

Estratégia de Transfection otimizado para gravação de Expressão e eletrofisiológicos de Recombinante dos canais de íon em HEK-293T Células

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Isolamento e Purificação de cinesina partir Drosophila Os embriões

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Preparação Individual Drosophila Ovos Câmaras de imagens ao vivo

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

Estudar checkpoint mitótico por Ilustrando comportamento dinâmico cinetocoro Proteína e Movimento de cromossomos em Viver<em&gt; Drosophila</em&gt; Os embriões Sincicial