Diese Arbeit wurde von der slowenischen Research Agency mit Projekt-Z2-9229 und das Programm P2-0249 unterstützt. Dieses Video stellt die ergänzende Materialien für den &quot;Elektroporation basierenden Technologien und Behandlungsmethoden&quot; wissenschaftlichen Workshop und Aufbaustudium, halbjährlich von der Fakultät für Elektrotechnik an der University of Ljubljana, Slowenien organisiert.

Teil I: Vorbereitende Schritte <ol><li> In diesem Experiment chinesischen Hamsters (CHO-K1) verwendet wird. Die Zellen werden in Lab-Tek II Kammern (2 Brunnen, 4 cm je 2) (Nalge Nunc, Deutschland) bei ~ 0.7x10 5 Zellen / ml in HAM-F12 Kulturmedium mit 8% fötalem Kälberserum, 0,15 mg / ml beschichtet L-Glutamin, 16 mg / ml Gentamycin (alle von Sigma-Aldrich, Steinheim, Deutschland) und 200 Einheiten / ml Crystacillin (Pliva, Zagreb, Kroatien) und inkubiert in 5% CO 2 bei 37 ° C. Alternativ können die Zellen auch auf # 1 Glasdeckgläschen (0,13 bis 0,16 mm dick), mit einer zellulären Klebstoffs wie Polylysin beschichtet werden. </li><li> Inkubieren Sie die Zellen in ihrem Kulturmedium. Inkubation dauert 2 bis 4 Stunden liefert Zellen, die noch annähernd kugelförmige, sondern fest auf der Oberfläche mit einem kleinen Teil ihrer Membran angebracht. Alternativ kann nach 16 bis 20 Stunden Inkubation werden die Zellen vollständig auf der Oberfläche angebracht und haben komplexere Formen, aber die meisten von ihnen sind noch nicht geteilt. </li><li> Bereiten Sie eine 10 mM Stammlösung von di-8-ANEPPS (Invitrogen, Eugene, Oregon, USA) durch Zugabe von 843 l DMSO (Sigma-Aldrich, Steinheim, Deutschland) bis 5 mg des Farbstoffs in der ursprünglichen Invitrogen Durchstechflasche. Die Stammlösung kann in einem Kühlschrank bei 4 ° C für mehrere Monate gelagert. Vor Beginn der Experimente, erwärmen die die Lösung, bis die Kristalle von DMSO auflösen. </li><li> Einige Zelllinien können die Verwendung von Pluronic, um den Farbstoff Einbau in die Zellmembran erleichtern. Pluronic kann in 20% Stammlösung in DMSO (F-127, Invitrogen, Eugene, Oregon, USA), oder eine Stammlösung der gleichen Konzentration kann durch Lösen von Pluronic in DMSO vorbereitet gekauft werden. Stammlösung von Pluronic kann bei Raumtemperatur gelagert werden. </li></ol> Teil II: Das Laden der Zellen mit di-8-ANEPPS <ol><li> Mix 3 ul 10 mM di-8-ANEPPS und 2,5 ul 20% Pluronic in 1 ml der Spinner Modifikation (Calcium-abgereicherten Version) des Minimum Essential Medium SMEM (medium M8167 oder M4767, Sigma-Aldrich, Steinheim, Deutschland ) in einem 1,5 ml Eppendorf-Röhrchen. Dies ergibt eine &quot;loading solution&quot; mit etwa 30 uM di-8-ANEPPS und 0,05% Pluronic. Für andere Zelltypen, können ihre eigene Kultur Medien statt SMEM verwendet werden. </li><li> Ersetzen Sie das Kulturmedium in den Lab-Tek Kammer mit dem Be-Lösung. Übertragen Sie die Kammer wieder in den Kühlschrank für 10 min bei 4 ° C. Bei dieser Temperatur ist die Internalisierung der Farbstoff durch die Plasmamembran weitgehend gehemmt. </li><li> Nach der Färbung, vorsichtig waschen die überschüssige Farbe zwei vor drei Mal mit reinem SMEM. </li><li> Lassen Sie 1,5 ml SMEM in der Kammer. </li></ol> Teil III: Experiment und Bildaufnahme <ol><li> Die Zellen werden beobachtet mit einem Fluoreszenzmikroskop (in unserem Fall Zeiss Axiovert 200, Zeiss, Deutschland) mit einem Ölimmersionsobjektiv (x63, NA 1,4), einem Monochromator (Polychrome IV, Visitron, Deutschland) und eine gekühlte CCD-Kamera (VisiCam ausgestattet 1280, Visitron, Deutschland). Die Bilder werden mit MetaFluor 7.1.1 erfasst und verarbeitet in MetaMorph 7.1.1 (beide Molecular Devices, Downingtown, PA, USA), aber auch andere ähnliche Software zur Erfassung kann ebenfalls verwendet werden. Stellen Sie die Anregungswellenlänge in der Software zur Erfassung bis 490 nm, und wählen Sie einen geeigneten Emissions-Filter für ANEPPS, zum Beispiel ein Band-Pass-Filter bei 605 nm zentriert (605/55m, dichroitische 565 DCXR, Chroma, Rockingham, VT, USA). Wenn ein Monochromator nicht verfügbar ist, kann ein Anregungsfilter bei 490 nm zentriert verwendet werden. </li><li> Setzen Sie die Kammer mit Zellen auf dem Mikroskoptisch, Position der Elektroden am Boden der Kammer und verbinden sie mit dem Impulsgenerator. Das Medium sollte sich auf den Elektroden. </li><li> Zur Aufrechterhaltung Lebensfähigkeit der Zellen und reduziert die Erwärmung, sollte der Puls niedrig genug Amplitude und kurzer Dauer sein. In diesem Experiment ein Rechteckimpuls mit 35 V Amplitude und 50 ms Dauer (niedrig genug, um Elektroporation zu vermeiden) wird unter Verwendung einer DC-Spannungsversorgung und eine maßgeschneiderte Mikroprozessor gesteuerten Umschalter Gerät. Alternativ kann auch eine kommerzielle Impulsgeber, wie 33210A (Agilent, Santa Clara, CA, USA) an einen Verstärker oder eine kommerzielle Stimulator, wie S88 (Grass, West Warwick, RI, USA) verbunden sind, verwendet werden können. Der Puls ist auf zwei parallel Pt / Ir-Draht-Elektroden mit 0,8 mm Durchmesser und 4 mm Abstand zwischen ihnen ausgeliefert, wodurch ein elektrisches Feld von etwa 88 V / cm zwischen den Elektroden und induzieren ΔΦ. Die genaue Feldverteilung zwischen den Elektroden in diesem Experiment verwendet wird an anderer Stelle [1] beschrieben. </li><li> Finden Sie die Zellen von Interesse. Tragen Sie einen einzigen elektrischen Impuls oder eine Folge von Impulsen. Für jeden Impuls, die Übernahme zweier Fluoreszenzbilder: eine unmittelbar vor dem Puls (das Steuerelement Bild), und man während des Pulses (Puls-Bild). Aufgrund der geringen Reaktionszeit von di-8-ANEPPS, werden die Änderungen in der Fluoreszenz schwierig, mit bloßem Auge zu erkennen und sich zeigen only nach der Verarbeitung. Der Puls Lieferung muss mit der Bildaufnahme, die ein Merkmal der Akquisition Software synchronisiert werden. Um zu vermeiden, Ausbleichen und mögliche Erwärmung kann die Beleuchtung auf die Dauer des Impulses begrenzt werden. </li></ol> Teil IV: Bildverarbeitung und-analyse <ol><li> Öffnen Sie die Bilder in MetaMorph Software. Für jeden Impuls, subtrahieren dem Hintergrund sowohl in der Kontrolle und der Puls Bild. </li><li> Wählen Sie eine Zelle und stellen Sie die Region von Interesse, so dass es entspricht der Membran. Messen Sie die Fluoreszenz-Intensitäten entlang dieser Region in die Steuerungs-und Puls-Image und übertragen die Werte in eine Tabellenkalkulation. </li><li> Für jeden Impuls, subtrahieren die Steuerdaten aus dem Impuls-Daten und teilt das Ergebnis durch Daten zur Steuerung der relativen Fluoreszenz Änderungen zu erhalten. Wenn eine Folge von Impulsen angewandt wird, können die Werte der relativen Fluoreszenz Änderungen für jeden Impuls bestimmt gemittelt, um eine zuverlässige Messung zu erhalten. </li><li> Transform der relativen Fluoreszenz Änderungen in ΔΦ mit Hilfe einer Kalibrationskurve. Eine grobe Schätzung dieser Kurve kann aus der Literatur gewonnen werden, aber für höhere Genauigkeit, muss es für die jeweilige Einrichtung gemessen werden. Für die Zellen und das Setup in diesem Experiment verwendet eine 6% ige Abnahme der Fluoreszenz entspricht einem 100 mV Anstieg ΔΦ [2]. </li><li> Schließlich Plot die Spannung als Funktion der relativen Bogenlänge im Grafikrechner Software-Paket wie Excel (Microsoft Corp, Redmond, WA, USA), Sigma Plot (Systat Software Inc., San Jose, CA, USA), oder Origin (OriginLab Corp, Northampton, MA, USA). Die Kurve kann auch geglättet mit einem geeigneten Filter, wie der gleitende Durchschnitt zu filtern. </li></ol>

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Eng. 34, 642-652 (2006).</li><li>Huang, C. J., Harootunian, A., Maher, M. P., Quan, C., Raj, C. D., McCormack, K., Numann, R., Negulescu, P. A., Gonzalez, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+voltage-gated+sodium-channel+blockers+by+electrical+stimulation+and+fluorescence+detection+of+membrane+potential.">Characterization of voltage-gated sodium-channel blockers by electrical stimulation and fluorescence detection of membrane potential.</a> Nat. Biotechnol. 24, 439-446 (2006).</li><li>Bedlack, R. S., Wei, M., Fox, S. H., Gross, E., Loew, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+electric+potentials+in+soma+and+neurite+membranes.">Distinct electric potentials in soma and neurite membranes.</a> Neuron. 13, 1187-1193 (1994).</li><li>Cheng, D. K. 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>

Messungen der induzierten Membran (transmembrane) Spannung, ΔΦ, werden in verschiedenen experimentellen Settings, wie Studien von spannungsabhängigen Membran Kanäle, Aktionspotential Ausbreitung, Herz-Zell-Stimulation oder Zellmembran Elektroporation [3, 4, 5, 6 wichtige , 7]. Mit einfachen Zellformen kann ΔΦ analytisch berechnet werden. Zum Beispiel für eine kugelförmige Zelle, ist ΔΦ durch die Schwan-Gleichung, so dass die Spannung proportional zur Feldstärke und Zellgröße Staaten und folgt der Cosinus-Funktion entlang der Membran gegeben [8, 9]. Für kompliziertere Zellformen kann ΔΦ weichen deutlich von den Cosinus und müssen entweder numerisch bestimmt werden, mithilfe eines Computers [2, 10, 11], oder experimentell mit Hilfe eines potentiometrischen Farbstoff [12, 13, 14, 15]. Einer der potentiometrischen Farbstoffe häufig für diesen Zweck verwendet wird di-8-ANEPPS (di-8-Butyl-Amino-naphthyl-Ethylen-Pyridinium-propyl-sulfonat), eine schnelle Farbstoff mit Anregungs-und Emissionsspektren abhängig von der Membran-Spannung, welche ermöglicht nicht-invasive Beobachtung der Variationen des ΔΦ auf der Zellmembran und seinen Wert zu messen. In diesem Video zeigen wir einen experimentellen Ansatz zur Bestimmung von ΔΦ mit di-8-ANEPPS. Der Farbstoff wurde durch Professor Leslie Loew und Kollegen [13, 14] entwickelt an der University of Connecticut und gehört zur Klasse der schnellen Reaktion Farbstoffe. di-8-ANEPPS ist nichtfluoreszierenden in Wasser und wird stark fluoreszierenden, wenn es in die Lipid-Doppelschicht der Zellmembran integriert. Eine Änderung ΔΦ ergibt sich eine Änderung der intramolekularen Ladungsverteilung und entsprechende Änderungen in den spektralen Profil und Intensität der Farbstoff-Fluoreszenz. Die Fluoreszenz-Intensität des di-8-ANEPPS variiert proportional zur Änderung des ΔΦ; die Reaktion des Farbstoffs ist linear für Spannungen im Bereich von -280 mV bis +250 mV [4, 16]. Relativ kleine Änderungen in der Fluoreszenz des Farbstoffes, unebene Membranfärbung und Farbstoff Internalisierung machen di-8-ANEPPS weniger geeignet für absolute Messungen von Membran-Spannung, z. B. seine Ruhestätte Komponente, obwohl diese Anstrengungen waren auch [17] berichtet. Es ist jedoch für die Messung von größeren Veränderungen in der Membran-Spannung, wie das Auftreten von induzierten Membran Spannung in nonexcitable Zellen, die äußere elektrische Felder [12, 13], oder Aktionspotentiale in erregbaren Zellen [4, 5]. Obwohl nicht angewendet hier, di-8-ANEPPS ermöglicht auch die Bestimmung von ΔΦ durch ratiometrische Messungen der Fluoreszenz-Anregung [18] oder Emission [19], die die Empfindlichkeit der Reaktion erhöht und die oben genannten Effekte. Als di-8-ANEPPS Flecken der Membran kann es auch deutlich, wie eine Membran-Marker verwendet werden [2]. Einer der Nachteile des Farbstoffes ist, dass es anfällig gegen Ausbleichen ist, so dass eine längere Exposition gegenüber dem starken Licht sollte vermieden werden. Die Kalibrierung der Farbstoff entweder mit (i) Kalium-Ionophor Valinomycin und eine Reihe von verschiedenen Kalium-Konzentrationen in externen Medium durchgeführt [2,18], oder (ii) Patch-Clamp-in Voltage-Clamp-Modus [17]. Schließlich mit den Messungen von ΔΦ auf sphärische Zellen und Zellen von komplexeren Formen, zeigt das Video den Einfluss der Zellform auf die Amplitude und die räumliche Verteilung von ΔΦ. So kugelförmigen Zellen ΔΦ ist in der Nähe eines Cosinus, im Einvernehmen mit Schwan-Gleichung, während für kompliziertere Zellformen die räumliche Verteilung der ΔΦ ist komplizierter [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

Die Platzierung einer Zelle in ein äußeres elektrisches Feld bewirkt eine lokale Ladungsumverteilung innerhalb und außerhalb der Zelle in der Nähe der Zellmembran, wodurch eine Spannung über der Membran. Diese Spannung, bezeichnet die induzierte Membran-Spannung (auch induzierte transmembrane Spannung oder induzierten transmembrane Potentialdifferenz) und bezeichnet durch ΔΦ, existiert nur, solange das externe Feld vorhanden ist. Wenn der ruhende Spannung wird auf der Membran, überlagert die induzierte Spannung (ergänzt) auf sie. Durch die Verwendung einer der potentiometrischen Fluoreszenzfarbstoffe, wie Di-8-ANEPPS, ist es möglich, die Schwankungen der ΔΦ auf der Zellmembran zu beobachten und ihren Wert nicht-invasiv zu messen. di-8-ANEPPS wird stark fluoreszierenden, wenn die Lipid-Doppelschicht der Zellmembran gebunden ist, mit der Änderung der Fluoreszenzintensität proportional zur Änderung des ΔΦ. Dieses Video zeigt das Protokoll für die Messung ΔΦ mit di-8-ANEPPS und zeigt auch den Einfluss der Zellform auf die Amplitude und die räumliche Verteilung von 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

Äußeres elektrisches Feld induziert eine Spannung an der Membran einer Zelle, bezeichnet die induzierte Membran-Spannung (ΔΦ). Mit Hilfe der potentiometrischen Farbstoff di-8-ANEPPS, ist es möglich, die ΔΦ-invasiv messen. Dieses Video zeigt das Protokoll für die Messung ΔΦ mit di-8-ANEPPS.

Messung der induzierten Spannung Membran mit Di-8-ANEPPS

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

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

Research

JoVE Journal

Biology

17.3K Aufrufe.  University of Ljubljana. Äußeres elektrisches Feld induziert eine Spannung an der Membran einer Zelle, bezeichnet die induzierte Membran-Spannung (ΔΦ). Mit Hilfe der potentiometrischen Farbstoff di-8-ANEPPS, ist es möglich, die ΔΦ-invasiv messen. Dieses Video zeigt das Protokoll für die Messung ΔΦ mit di-8-ANEPPS.

Serie: 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.

Messen in der Nähe Plasmamembran und Global intrazellulären Calcium-Dynamics in Astrozyten

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

Forschung

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.

Mess-Plasmamembranprotein endocytischen Preise von Reversible Biotinylierung

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.

Crystallizing Membranproteinen zur Strukturaufklärung mittels Lipidische Mesophasen

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.

Die Kristallisation von Membranproteinen in Lipidische Mesophasen

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.

Die Untersuchung der Konformationsänderungen Dynamics von Membranproteinen In situ Mit Site-directed Fluoreszenzmarkierung

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.

Die<em&gt; Xenopus</em&gt; Eizelle Cut-open Vaseline Gap Voltage-Clamp-Technik Mit Fluorometrie

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 Plasmamembran Verformungen Mit 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.

Mess Glutathion-induzierte Fütterung Antwort in 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.

Mikroperfusion Technik zur Untersuchung von Mikrogefäßdurchlässigkeit Verordnung in Rattengekröse

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.

Grundzustand Erschöpfung Super-Resolution Imaging in Säugerzellen

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