Ce travail a été soutenu par l&#39;Agence slovène pour la recherche avec le projet Z2-9229 et le programme P2-0249. Cette vidéo représente le matériau supplémentaire pour les &quot;électroporation basée sur les technologies et traitements&quot; atelier scientifique et cours de troisième cycle, organisé tous les deux ans par la Faculté de génie électrique à l&#39;Université de Ljubljana, en Slovénie.

Partie I: Les étapes préliminaires <ol><li> Dans cette expérience chinoise en ligne de hamster cellules ovariennes (CHO-K1) est utilisé. Les cellules sont étalées dans le Lab-Tek II chambres (2 puits, 4 cm 2 chacun) (Nalge Nunc, Allemagne) à ~ 0.7x10 5 cellules / ml dans HAM-F12 milieu de culture supplémenté avec 8% sérum de veau foetal, 0,15 mg / ml L-glutamine, 16 mg / ml de gentamicine (tous de Sigma-Aldrich, Steinheim, Allemagne), et 200 unités / ml Crystacillin (Pliva, Zagreb, Croatie), et incubés dans 5% de CO 2 à 37 ° C. Alternativement, les cellules peuvent également être plaqués sur le n ° 1 des lamelles de verre (0,13 à 0,16 mm d&#39;épaisseur), enduit d&#39;un adhésif cellulaires tels que la polylysine. </li><li> Incuber les cellules dans leur milieu de culture. L&#39;incubation dure de 2 à 4 heures rendements des cellules qui sont encore à peu près sphérique, mais fermement attaché à la surface avec une petite partie de leur membrane. Sinon, après 16 à 20 heures d&#39;incubation, les cellules sont totalement attachés à la surface et ont des formes plus complexes, mais la plupart d&#39;entre eux ne sont toujours pas se diviser. </li><li> Préparer une solution stock de 10 mM de di-8-ANEPPS (Invitrogen, Eugene, Oregon, USA) en ajoutant 843 ul de DMSO (Sigma-Aldrich, Steinheim, Allemagne) à 5 mg de colorant dans le flacon d&#39;origine Invitrogen. La solution mère peut être stockée dans un réfrigérateur à 4 ° C pendant plusieurs mois. Avant de commencer les expériences, réchauffer la solution jusqu&#39;à ce que les cristaux du DMSO de se dissoudre. </li><li> Certaines lignées cellulaires peuvent nécessiter l&#39;utilisation de pluronic pour faciliter l&#39;incorporation du colorant dans la membrane cellulaire. Pluronic peut être acheté dans une solution stock de 20% dans le DMSO (F-127, Invitrogen, Eugene, Oregon, USA), ou une solution stock de la même concentration peut être préparée en dissolvant pluronique dans le DMSO. Solution mère de Pluronic peut être conservé à température ambiante. </li></ol> Partie II: Chargement des cellules avec des di-8-ANEPPS <ol><li> Mélanger 3 ul de 10 mM di-8-ANEPPS et 2,5 l de pluronic 20% dans 1 ml de la modification Spinner (calcium appauvri version) de l&#39;SMEM milieu essentiel minimum (moyenne M8167 ou M4767, Sigma-Aldrich, Steinheim, Allemagne ) dans un tube Eppendorf de 1,5 ml. Cela donne une «solution de chargement&quot;, contenant environ 30 uM di-8-ANEPPS et 0,05% de pluronic. Pour d&#39;autres types cellulaires, leurs milieux de culture indigène peut être utilisé au lieu du SMEM. </li><li> Remplacez le milieu de culture dans la chambre de Lab-Tek avec la solution de chargement. Transfert de la chambre au réfrigérateur pendant 10 min à 4 ° C. A cette température, l&#39;internalisation du colorant à travers la membrane plasmique est largement inhibée. </li><li> Après coloration, laver délicatement l&#39;excès de colorant deux à trois fois avec des SMEM pur. </li><li> Laisser 1,5 ml de SMEM dans la chambre. </li></ol> Partie III: Expérimentation et d&#39;acquisition d&#39;image <ol><li> Les cellules sont observées en utilisant un microscope à fluorescence (dans notre cas Zeiss Axiovert 200, Zeiss, Allemagne) équipé d&#39;un objectif à immersion (x63, NA 1,4), un VisiCam monochromateur (Polychrome IV, Visitron, Allemagne) et une caméra CCD refroidie ( 1280, Visitron, Allemagne). Les images sont acquises avec MetaFluor 7.1.1 et traitées dans MetaMorph 7.1.1 (deux Molecular Devices, Downingtown, PA, USA), mais d&#39;autres logiciels d&#39;acquisition similaire peut également être utilisé. Régler la longueur d&#39;onde d&#39;excitation dans le logiciel d&#39;acquisition à 490 nm et de choisir un filtre d&#39;émission approprié pour ANEPPS, par exemple, un filtre passe-bande centré à 605 nm (605/55m, dichroïque 565 DCXR, Chroma, Rockingham, Vermont, USA). Si un monochromateur n&#39;est pas disponible, un filtre d&#39;excitation centrée à 490 nm peut être utilisée à la place. </li><li> Placez la chambre avec des cellules sur la scène microscope, la position des électrodes dans le bas de la chambre et les connecter au générateur d&#39;impulsions. Le milieu doit recouvrir les électrodes. </li><li> Afin de maintenir la viabilité cellulaire et de réduire le chauffage, l&#39;impulsion doit être d&#39;amplitude suffisamment faible et de courte durée. Dans cette expérience, une impulsion carrée avec 35 V d&#39;amplitude et de 50 ms (assez bas pour éviter l&#39;électroporation) est généré en utilisant une alimentation en tension DC et une sur-mesure à microprocesseur dispositif de commutation. Alternativement, un générateur d&#39;impulsions commerciales, telles que 33210A (Agilent, Santa Clara, CA, USA) relié à un amplificateur ou un stimulateur commerciaux, tels que S88 (Grass, West Warwick, Rhode Island, USA), peuvent être utilisés. L&#39;impulsion est délivrée à deux parallèles électrodes en fil de Pt / Ir de 0,8 mm de diamètre et 4 mm de distance entre eux, créant un champ électrique d&#39;environ 88 V / cm entre les électrodes et ΔΦ inducteur. La distribution du champ exact entre les électrodes utilisées dans cette expérience est décrite par ailleurs [1]. </li><li> Trouver les cellules d&#39;intérêt. Appliquer une seule impulsion électrique ou une séquence d&#39;impulsions. Pour chaque impulsion, l&#39;acquisition de deux images de fluorescence: un immédiatement avant que l&#39;impulsion (l&#39;image de contrôle), et un cours de l&#39;impulsion (l&#39;image d&#39;impulsion). En raison de la faible réponse du di-8-ANEPPS, les changements dans la fluorescence sont difficiles à discerner à l&#39;œil nu et deviennent apparents ONLY après traitement. La livraison d&#39;impulsion doit être synchronisé avec l&#39;acquisition de l&#39;image, qui est une caractéristique du logiciel d&#39;acquisition. Pour éviter de photoblanchiment et possible de chauffage, l&#39;éclairage peut être limitée à la durée de l&#39;impulsion. </li></ol> Partie IV: traitement et analyse d&#39;images <ol><li> Ouvrez les images dans le logiciel MetaMorph. Pour chaque impulsion, il faut soustraire le fond à la fois dans le contrôle et l&#39;image de pouls. </li><li> Choisissez une cellule et définir la région d&#39;intérêt afin qu&#39;elle corresponde à la membrane. Mesurer les intensités de fluorescence le long de cette région dans l&#39;image de contrôle et d&#39;impulsion et de transférer les valeurs vers un tableur. </li><li> Pour chaque impulsion, il faut soustraire les données de contrôle à partir des données d&#39;impulsion, et divisez le résultat par des données de contrôle pour obtenir les changements relatifs à fluorescence. Si une séquence d&#39;impulsions est appliqué, les valeurs de relative changements de fluorescence déterminés pour chaque impulsion peut être calculée pour obtenir une mesure plus fiable. </li><li> Transformer les changements relatifs à fluorescence en utilisant un ΔΦ courbe de calibration. Une estimation approximative de cette courbe peut être obtenu à partir de la littérature, mais pour une plus grande précision, elle doit être mesurée pour chaque installation particulière. Pour les cellules et la configuration utilisés dans cette expérience une baisse de 6% de la fluorescence correspond à une augmentation de 100 mV dans ΔΦ [2]. </li><li> Enfin, l&#39;intrigue de la tension en fonction de la longueur d&#39;arc relative dans un logiciel graphique comme Excel (Microsoft Corp, Redmond, WA, USA), Terrain de Sigma (Systat Software Inc, San Jose, CA, USA), ou Origine (OriginLab Corp, Northampton, MA, USA). La courbe peut aussi être lissé à l&#39;aide d&#39;un filtre approprié, tel que le filtre moyenne mobile. </li></ol>

<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. L., Tung, L., Sobie, E. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonuniform+responses+of+transmembrane+potential+during+electric+field+stimulation+of+single+cardiac+cells.">Nonuniform responses of transmembrane potential during electric field stimulation of single cardiac cells.</a> Am. J. Physiol. 277, H351-H362 (1999).</li><li>Teissie, J., Eynard, N., Gabriel, B., Rols, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electropermeabilization+of+cell+membranes.">Electropermeabilization of cell membranes.</a> Adv. Drug. Del. 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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ratiometry+of+transmembrane+voltage-sensitive+fluorescent+dye+emission+in+hearts.">Ratiometry of transmembrane voltage-sensitive fluorescent dye emission in hearts.</a> Am. J. Physiol. Heart Circ. Physiol. 279, H1421-H1433 (2000).</li><li>Pucihar, G., Miklav&#269;i&#269;, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+time-dependent+numerical+model+of+transmembrane+voltage+inducement+and+electroporation+of+irregularly+shaped+cells.">A time-dependent numerical model of transmembrane voltage inducement and electroporation of irregularly shaped cells.</a> IEEE T. Biomed. Eng. 56, 1491-1501 (2009).</li></ol>

Mesures de la membrane induite (transmembranaire) de tension, ΔΦ, peut être important dans divers contextes expérimentaux, tels que les études des canaux membranaires voltage-dépendants, la propagation du potentiel d&#39;action, la stimulation des cellules cardiaques, ou électroporation membrane cellulaire [3, 4, 5, 6 , 7]. Avec des formes simples cellules, ΔΦ peut être calculé analytiquement. Par exemple, pour une cellule sphérique, ΔΦ est donné par l&#39;équation de Schwan s, ce qui indique que la tension est proportionnelle à l&#39;intensité du champ et de la taille des cellules et suit la fonction cosinus long de la membrane [8, 9]. Pour plus de formes cellulaires complexes, ΔΦ peuvent s&#39;écarter considérablement de la cosinus et doit être déterminée soit numériquement, en utilisant un ordinateur [2, 10, 11], ou expérimentalement, en utilisant un colorant potentiométrique [12, 13, 14, 15]. Un des colorants potentiométrique largement utilisés à cette fin est di-8-ANEPPS (di-8-butyl-amino-naphtyl-éthylène-pyridinium-propyl-sulfonate), un colorant rapide avec excitation et d&#39;émission des spectres dépendant de la tension de membrane, qui permet des observations non invasif des variations de l&#39;ΔΦ sur la membrane cellulaire et de mesurer sa valeur. Dans cette vidéo, nous montrons une approche expérimentale pour la détermination de ΔΦ en utilisant di-8-ANEPPS. Le colorant a été développé par le professeur Leslie Loew et ses collègues [13, 14] à l&#39;Université du Connecticut et appartient à la classe des colorants à réponse rapide. di-8-ANEPPS est non fluorescent dans l&#39;eau et devient fortement fluorescent lorsque l&#39;intègre dans la bicouche lipidique de la membrane cellulaire. Un changement dans les résultats ΔΦ à un changement de la distribution de charge intramoléculaire et les changements correspondants dans le profil spectral et l&#39;intensité de la fluorescence du colorant. L&#39;intensité de fluorescence des di-8-ANEPPS varie proportionnellement à la variation des ΔΦ, la réponse de la teinture est linéaire pour des tensions allant de -280 mV à 250 mV [4, 16]. Des changements relativement petits de la fluorescence du colorant, coloration de la membrane inégale, et l&#39;internalisation de teinture font di-8-ANEPPS moins approprié pour des mesures absolues de la tension de la membrane, par exemple, sa composante de repos, même si ces efforts ont également été rapportés [17]. Il est, cependant, adapté à la mesure de plus grands changements dans la tension de membrane, tels que l&#39;apparition de la tension de la membrane induite dans les cellules exposées nonexcitable aux champs électriques externes [12, 13], ou de potentiels d&#39;action dans les cellules excitables [4, 5]. Même s&#39;il n&#39;est pas appliqué ici, di-8-ANEPPS permet également la détermination de ΔΦ par des mesures d&#39;excitation de fluorescence ratiométrique [18] ou de l&#39;émission [19], ce qui augmente la sensibilité de la réponse et réduit les effets mentionnés ci-dessus. Comme di-8-ANEPPS taches de la membrane, il peut également être utilisé simplement comme un marqueur de membrane [2]. Un des inconvénients de la teinture, c&#39;est qu&#39;il est enclin à photoblanchiment, de sorte que les expositions prolongées à la lumière forte devrait être évitée. Calibrage de la teinture est réalisée soit avec valinomycine (i) ionophore de potassium et d&#39;un ensemble de différentes concentrations de potassium en milieu externe [2,18], ou (ii) de patch-clamp en mode voltage [17]. Enfin, avec les mesures de ΔΦ sur les cellules sphériques et des cellules de formes plus complexes, la vidéo montre l&#39;influence de la forme de la cellule sur la distribution spatiale de l&#39;amplitude et ΔΦ. Ainsi, pour les cellules sphériques ΔΦ est proche d&#39;un cosinus, en accord avec l&#39;équation de Schwan, tandis que pour des formes plus compliquées de cellules de la distribution spatiale des ΔΦ est plus complexe [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

Placement d&#39;une cellule dans un champ électrique externe provoque un intérieur frais locaux de redistribution et à l&#39;extérieur de la cellule dans le voisinage de la membrane cellulaire, résultant en une tension à travers la membrane. Cette tension, appelée tension de la membrane induite (tension transmembranaire a également induit, ou induite différence de potentiel transmembranaire) et désigné par ΔΦ, n&#39;existe que tant que le champ extérieur est présent. Si la tension au repos est présent sur la membrane, la tension induite superpose (ADDS) sur elle. En utilisant l&#39;un des colorants fluorescents potentiométrique, tels que le di-8-ANEPPS, il est possible d&#39;observer les variations de ΔΦ sur la membrane cellulaire et de mesurer sa valeur non invasive. di-8-ANEPPS devient fortement fluorescent lorsqu&#39;il est lié à la bicouche lipidique de la membrane cellulaire, avec le changement de l&#39;intensité de fluorescence proportionnelle à la variation des ΔΦ. Cette vidéo montre le protocole de mesure de ΔΦ utilisant di-8-ANEPPS et démontre aussi l&#39;influence de la forme des cellules sur l&#39;amplitude et la répartition spatiale des 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

Champ électrique externe induit une tension sur la membrane d&#39;une cellule, appelée tension de membrane induite (ΔΦ). En utilisant le colorant potentiométrique di-8-ANEPPS, il est possible de mesurer la ΔΦ non invasive. Cette vidéo montre le protocole de mesure de ΔΦ utilisant di-8-ANEPPS.

Mesurer la tension de membrane induite avec 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 ...

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JoVE Journal

Biology

17.3K vues.  University of Ljubljana. Champ électrique externe induit une tension sur la membrane d'une cellule, appelée tension de membrane induite (ΔΦ). En utilisant le colorant potentiométrique di-8-ANEPPS, il est possible de mesurer la ΔΦ non invasive. Cette vidéo montre le protocole de mesure de ΔΦ utilisant di-8-ANEPPS.

Vidéo: 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.

Mesure Proche membrane plasmique et mondiale dynamique du calcium intracellulaire dans les 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, ...

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Biologie

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.

Mesure du taux plasmatique des protéines membranaires par endocytique Biotinylation réversible

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

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

Cristallisant protéines membranaires pour déterminer la structure à l&#39;aide mésophases lipidique

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

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.

La cristallisation des protéines membranaires dans des mésophases lipidique

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.

L&#39;examen de la dynamique conformationnelle des protéines membranaires In situ Avec site-dirigé marquage fluorescent

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.

La<em&gt; Xenopus</em&gt; Ovocytes Cut-ouvert vaseline Gap voltage-clamp Technique Avec Fluorimétrie

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.

Imaging Plasma Membrane déformations Avec 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.

Mesure induite glutathion-réponse d&#39;alimentation à 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.

Microperfusion Technique chargé d&#39;enquêter sur le règlement des microvaisseaux perméabilité au 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.

État fondamental l’appauvrissement Super-résolution imagerie dans les cellules de mammifères

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

Methods Article

Mesurer la tension de membrane induite avec Di-8-ANEPPS