Receita encolher solução é a partir do laboratório de RW Aldrich. Agradecemos aos seguintes agências de fomento e fundações de apoio: National Institutes of Health, National Science Foundation, a American Heart Association, Associação de Distrofia Muscular, o B. Donald e Delia E. Baxter Foundation, o Fundo Klingenstein e da Fundação McKnight for Neuroscience.

Parte 1: Remover a membrana vitelina <ol><li> Prepare dois conjuntos de fórceps. Nós preferimos No. 5 fórceps, onde um conjunto foi ligeiramente afiada com uma lima. Também preparar uma pipeta de vidro por corte e polimento-fogo de 7 &quot;Pasteur pipeta. </li><li> Recomendações: Para evitar que os oócitos de escorregar, corte um círculo de grade (1 praças mm) e cola na parte inferior de uma placa de Petri 60mm. Este prato pode ser reutilizado indefinidamente se mantidos limpos entre os usos. </li><li> Encha o prato de Petri até a metade com solução hiperosmótico (ver receita) e colocar sob um microscópio de dissecação. </li><li> Remover oócitos 03/01 Xenopus da sua solução de incubação com uma pipeta Pasteur polido e coloque no prato. </li><li> Aguarde 15s - 2 min para que as células começam a encolher. Vezes mais fazer as células mais fáceis de descascar, tempos mais curtos para as células saudáveis ​​para remendar. </li><li> Com a redução da célula, a membrana vitelina começará a tornar-se visível apenas como uma camada transparente sobre a célula. Selecione uma célula saudável (sem voltas ou defeitos) para peeling. </li><li> Com um par de fórceps, gentilmente compreender a membrana vitelina sem danificar a membrana plasmática. Com a outra, segure perto do mesmo local e gentilmente rasgar a membrana transparente e livre para além da célula. </li><li> Com a pipeta de Pasteur, cuidadosamente mover a célula para a câmara de gravação. Note-se que a célula será frágil após a remoção vitelínico. </li></ol> Parte 2: geração Gigaseal <ol><li> Preencha uma pipeta de gravação com solução salina. Use o volume mínimo de solução que faz um bom contato elétrico com o fio de eletrodo de modo a minimizar a capacitância pipeta. </li><li> Flick os tempos de pipeta para permitir que várias bolhas a flutuar e deslizar no fio do eletrodo. Aplicar pressão para a pipeta cheia. Isto pode ser feito por via oral ou com uma bomba de aquário de uma loja de animais. Esta pressão é fundamental para manter a pipeta limpa, uma vez que atravessa a interface líquido e se move para a célula. </li><li> Encontrar o óvulo na câmara e focar fortemente na borda da célula. </li><li> Com a pipeta de sair do banho, o centro da ponta (fora de foco) acima da célula. Desta vez, minimiza gasto no banho antes da geração do selo. </li><li> Queda da pipeta para baixo em foco ao lado do celular. Trazê-lo em estreita proximidade (~ 50 microns). Você deverá ver uma pequena distorção de solução empurrado para fora da pipeta pela pressão de volta. </li><li> Usando o software patch clamp, verifique a resistência da pipeta (nossa meta é de aproximadamente 3-4 mohms) e zerar o atual usando a tensão de offset. </li><li> Com uma pressão positiva sobre a pipeta, mova a ponta lentamente em direção ao celular enquanto monitoramento da resistência visualmente ou utilizando um monitor de áudio que gera um tom cuja freqüência é proporcional à resistência. Como a ponta começa a tocar o celular, a resistência vai aumentar. </li><li> De repente, mas com cuidado mudar de positivo para a pressão negativa de aproximadamente o mesmo valor absoluto. Resistência deve aumentar para selar formação. </li><li> A maior parte do tempo, a formação de vedação ocorre dentro de alguns segundos, ocasionalmente 30-60 segundos são necessários. </li><li> Uma vez que a resistência está em 1GΩ, a pressão pode ser liberado para neutro. Você agora tem um patch on-celular. Para um patch de dentro para fora, afastar a célula rapidamente. Para fora-out, veja abaixo. </li></ol> Parte 5: Fora-out topologia (opcional) <ol><li> Imediatamente após atingir uma ruptura, gigaseal da membrana. Isto pode ser feito com sucção forte, mas nós preferimos a 1 V de pulso para 1 ms. A resistência deve cair para pouco mais do que a resistência inicial da pipeta. </li><li> Mover a pipeta fora da célula lenta e suavemente. Você deverá ver uma parte da célula com o puxar da pipeta. De repente, e dentro de alguns segundos, o celular vai pular de volta ea resistência deve imediatamente e, simultaneamente, voltar ao GΩs. </li><li> Você agora tem um patch na configuração fora-out. Ectodomínios de qualquer canais iônicos incluído é exposta ao banho. </li></ol> Parte 6: Resultados Previstos Geralmente, os focas maior resistência são preferíveis e mais vivido. 10 GÊ é uma boa orientação de alta qualidade selos. Em nossa experiência, as chances de selos desta qualidade variam de acordo com muitos fatores, especialmente celular e pipeta de qualidade. Taxas de aquisição patch pode ser mais de 95% com as células saudáveis, pipetas limpas, e um investigador experiente. Essas manchas podem durar muitos minutos e são adequados para qualquer estudo eletrofisiológico de canais iônicos expressos em oócitos Xenopus, incluindo um único canal de gravações.

<ol>
	<li>Neher, E., Sakmann, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-channel+currents+recorded+from+membrane+of+denervated+frog+muscle+fibres.">Single-channel currents recorded from membrane of denervated frog muscle fibres.</a> Nature. 260, 799-802 (1976).</li><li>Sackmann, B., Neher, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Single-channel Recording. , (1983).</li></ol>

Há muitos parâmetros de gravação eletrofisiológico não serão discutidos aqui. Configuração de equipamento, sistema de gestão de ruído, expressão de canais e protocolos de gravação são também essenciais para bons resultados experimentais. Em nossa experiência, estes são os parâmetros mais críticos para a formação de selos de confiança: oócitos de alta qualidade, a remoção da membrana vitelina rapidamente (aka, &quot;peeling&quot;), use pipetas recém puxado protegido de poeira, pipetas suave, a pressão posi...

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		<div>
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		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Patch Clamp Amplifier & Software</td>
<td>Instrument</td>
<td>HEKA or Axon</td>
<td>EPC-10</td>
<td>Or other similar device (e.g. HEKA EPC-9 or Axopatch 200B)</td>
</tr>
<tr>
<td>No. 5 forceps (two pairs)</td>
<td>Tool</td>
<td>Fine Science Tools</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Oocyte shrinking solution</td>
<td>Reagent</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>Ingredient In mM
N-methyl-D-glucamine 220
HEPES 10
MgCl2 1
EGTA 10
Aspartic Acid 220
KCl 2
pH to 7.4 w/ N-methyl-D-glucamine</td>
</tr>
<tr>
<td>woven mesh</td>
<td>800 um</td>
<td>Spectrum Laboratories INC</td>
<td>146481</td>
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patch clamp recording of ion channels expressed in  xenopus oocytes

Desde o seu desenvolvimento por Sakmann e Neher 1, 2, o patch clamp se estabeleceu como uma técnica extremamente úteis para a aferição eletrofisiológica de canais de iões simples ou múltiplas nas células. Esta técnica pode ser aplicada a canais iônicos tanto no ambiente de sua nativa e expresso em células heterólogas, como oócitos colhidos o sapo com garras Africano, Xenopus laevis. Aqui, descrevemos a técnica bem estabelecida de gravação de patch-clamp de oócitos Xenopus. Esta técnica é utilizada para medir as propriedades de canais iônicos, expressos quer em populações (macropatch) ou individualmente (single-channel de gravação). Nós nos concentramos em técnicas para maximizar a qualidade da preparação de oócitos e geração de selo. Com todos os fatores otimizados, esta técnica dá uma probabilidade de geração de selo de sucesso mais de 90 por cento. O processo pode ser otimizado de forma diferente por cada pesquisador com base nos fatores que ele ou ela acha mais importante, e nós apresentamos a abordagem que levaram à maior sucesso em nossas mãos.

Watch this Scientific Journal Video about Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes at JoVE.com

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Este destina-se como uma introdução para o patch clamp de gravação a partir de oócitos Xenopus laevis. Abrange vitelínico remoção da membrana, a formação de um selo gigaohm (gigaseal), ea conversão opcional do patch para a topologia de fora-out.

Gravação de patch-clamp de canais iônicos Expresso em Oócitos Xenopus

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

patch-clamp-recording-of-ion-channels-expressed-in-xenopus-oocytes

Research

JoVE Journal

Biology

26.0K visualizações.  Stanford University. Este destina-se como uma introdução para o patch clamp de gravação a partir de oócitos Xenopus laevis. Abrange vitelínico remoção da membrana, a formação de um selo gigaohm (gigaseal), ea conversão opcional do patch para a topologia de fora-out.

Vídeo: Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Methods for Patch Clamp Capacitance Recordings from the Calyx

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. Electrical recordings from the presynaptic terminal allow the measurement of action potentials, calcium channel currents, vesicle fusion (exocytosis) and subsequent membrane uptake (endocytosis). The fusion of vesicles containing neurotransmitter causes the vesicle membrane to be added to the cell membrane of the calyx. This increase in the amount of cell membrane is measured as an increase in capacitance. The subsequent reduction in capacitance indicates endocytosis, the process of membrane uptake or removal from the calyx membrane. Endocytosis, is necessary to maintain the structure of the calyx and it is also necessary to form vesicles that will be filled with neurotransmitter for future exocytosis events. Capacitance recordings at the calyx of Held have made it possible to directly and rapidly measure vesicular release and subsequent endocytosis in a mammalian CNS nerve terminal. In addition, the corresponding postsynaptic activity can be simultaneously measured by using paired recordings. Thus a complete picture of the presynaptic and postsynaptic electrical activity at a central nervous system synapse is achievable using this preparation. Here, the methods for slice preparation, morphological features for identification of calyces of Held, basic patch clamping techniques, and examples of capacitance recordings to measure exocytosis and endocytosis are presented.

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal.

Métodos para Recordings patch-clamp Capacitância da Calyx

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. ...

Biologia

Pressure-polishing Pipettes for Improved Patch-clamp Recording

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording. We first developed this method for fabricating pipettes suitable for recording from small (&lt;3 m) neuronal cell bodies. The basic principal is similar to glass-blowing and combines air pressure and heat to modify the shape of patch pipettes prepared by a conventional micropipette puller. It can be applied to so-called soft (soda lime) and hard (borosilicate) glasses. Generally speaking, pressure polishing can reduce pipette resistance by 25% without decreasing the diameter of the tip opening (Goodman and Lockery, 2000). It can be applied to virtually any type of glass and requires only the addition of a high-pressure valve and fitting to a microforge. This technique is essential for recording from ultrasmall cells (&lt;5 m) and can also improve single-channel recording by minimizing pipette resistance. The blunt shape is also useful for perforated-patch clamp recording since this tip shape results in a larger membrane bleb available for perforation.

This is a guide to modifying the shape of glass micropipettes. Specifically, by using heat and air pressure the taper is widened without increasing the tip opening, leading to lower pipette resistance. This is critical to obtain low noise recordings of small cells but is useful in many applications.

Pressão polimento Pipetas para Gravação de patch-clamp melhorada

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording.  We first developed this method for fabricating pipettes suitable for ...

Microinjection of Xenopus Laevis Oocytes

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic transport. Because of its large nucleus and high density of nuclear pore complexes (NPCs), nuclear transport can be easily visualized in the Xenopus oocyte. Much insight into the mechanisms of nuclear import and export has been gained through use of this system (reviewed by Pant&eacute;, 2006). In addition, we have used microinjection of Xenopus oocytes to dissect the nuclear import pathways of several viruses that replicate in the host nucleus. 
Here we demonstrate the cytoplasmic microinjection of Xenopus oocytes with a nuclear import substrate. We also show preparation of the injected oocytes for visualization by thin-sectioning EM, including dissection, dehydration, and embedding of the oocytes into an epoxy embedding resin. Finally, we provide representative results for oocytes that have been microinjected with the capsid of the baculovirus Autographa californica nucleopolyhedrovirus (AcMNPV) or the parvovirus Minute Virus of Mice (MVM), and discuss potential applications of the technique.

Here we demonstrate cytoplasmic microinjection of Xenopus laevis oocytes with a nuclear import substrate, as well as preparation of the injected oocytes for visualization by thin-sectioning electron microscopy.

Microinjeção de Oócitos Xenopus laevis

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic ...

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

Visualizando Localização RNA em Xenopus Oócitos

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

Fertilization of Xenopus oocytes using the Host Transfer Method

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to generate maternal-effect mutant animals. Additionally, many of the techniques to overexpress or inhibit gene function in organisms such as Xenopus and zebrafish fail to sufficiently target critical maternal signaling pathways, such as Wnt signaling. In Xenopus, manipulating gene function in cultured oocytes and subsequently fertilizing them can ameliorate these problems to some extent. Oocytes are manually defolliculated from donor ovary tissue, injected or treated in culture as desired, and then stimulated with progesterone to induce maturation. Next, the oocytes are introduced into the body cavity of an ovulating host female frog, whereupon they will be translocated through the host's oviduct and acquire modifications and jelly coats necessary for fertilization. The resulting embryos can then be raised to the desired stage and analyzed for the effects of any experimental perturbations. This host-transfer method has been highly effective in uncovering basic mechanisms of early development and allows a wide range of experimental possibilities not available in any other vertebrate model organism.

Fertilization of Xenopus oocytes using the Host Transfer Method

Procedure for fertilizing Xenopus oocytes by the host transfer method.

Fertilização de Oócitos Xenopus Utilizando o método de transferência do host

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to ...

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

Mutagênese e Análise Funcional de canais iônicos Heterologously Expresso em células de mamíferos

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

Grampo patch e Técnicas de Perfusão para estudar canais iônicos Expresso em Xenopus Oócitos

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

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) ...

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

Preparação de lipossomas gigantes por Imagem e patch-clamp Eletrofisiologia

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

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 ...