This work was supported by NIH R01 EY020533, WSU Startup Fund, and RPB grants.

Ethikerklärung: Verfahren unter Verwendung tierischer Probanden wurden von der Institutional Animal Care und Verwenden Committee (IACUC) an der Wayne State University zugelassen. 1. Vorbereitung der Versuchslösung <ol><li> Vorbereiten des Präparationslösung 1 Tag bis 1 Woche vor dem eigentlichen Experiment. Verwenden Sie ein HEPES-Pufferlösung für die Netzhaut Dissektion wegen seiner starken Pufferfähigkeit bei niedrigen Temperaturen 16. Mischen Sie alle Chemikalien wie folgt (in mM): 115 NaCl, 2,5 KCl, 2,5 CaCl 2, 1,0 MgCl 2, 10 HEPES, 28 Glucose. PH-Wert auf 7,4 mit NaOH. Die Lösung wird in einem Kühlschrank bis zu 1 Woche. </li><li> Aufzeichnungslösung ist Ames &#39;Medium, das eine künstliche Rückenmarksflüssigkeit (aCSF) für die Netzhautpräparate 17 ausgelegt ist. Mit dem Tag des Experiments abzuwiegen Ames-Pulver (4,4 g für 500 ml Lösung) in einem Rohr. Auch abwiegen NaHCO 3 (0,95 g für 500 ml Lösung) und mit der A mischenmes &quot;Pulver. </li><li> Bereiten Sie die intrazellulären Pipette Lösung für Patch Clamp Messungen von der Tag des Experiments. Mischen Sie alle Chemikalien wie folgt (in mM): 111 K-Gluconat, 1,0 CaCl 2, 10 HEPES, 1,1 EGTA, 10 NaCl, 1.0 MgCl2, 5 ATP-Mg und 1,0 GTP-Na. PH-Wert auf 7,2 mit KOH. Filtern der Pipettenlösung mit einem herkömmlichen Spritzenfilter. Stellen Sie ~ 500 ul Aliquots und speichern sie in einem Gefrierschrank oder eine Tiefkühltruhe. </li></ol> 2. Vorbereitung für den Tag der Experiment <ol><li> In der Nacht vor dem Experiment, legen Sie eine Maus in einen Käfig (C57BL / 6J oder ähnlichen Hintergrund, 4 - 8 Wochen alt, männlich) in einem dunklen Raum für O / N Dunkeladaptation. </li><li> Vorbereitung für die Präparation: <ol><li> Füllen Sie sezieren Lösung (100 - 200 ml) in einem Becherglas, legen Sie es auf Eis und Blase mit Sauerstoff für mindestens 10 min. </li><li> Starten Sie die Sauerstoffversorgung des dunklen Kasten Netzhautvorbereitung Lagerung. </li><li> Bereiten Kunststoffdeckgläser mit Fett Schienen für WiederDarm-Schnittpräparate und legen Sie jeden von ihnen in einem 35-mm-Kunststoffschale. </li><li> Setzen Sie eine neue Rasierklinge zu einem Chopper (Gewebeschneidemaschine). </li><li> Schneiden Sie einen Filtermembran in zwei Hälften. Dies ist für die Netzhaut-Schneiden. </li><li> Richten Sie die Sezieren Werkzeuge und Transferpipetten in der Nähe des Binokular. Der Arbeitsbereich ist auch so gestaltet sein, dass jedes Werkzeug kann im Dunkeln leicht zugegriffen werden. </li></ol></li><li> Vorbereitung für die Patch-Clamp-Aufnahmen: <ol><li> Löse Ames Pulver und NaHCO 3 in destilliertem Wasser (Ames 4,4 g und 0,95 g NaHCO 3/500 ml ddH 2 O). Blasen AMES &#39;Medium mit gemischten Sauerstoff (95% O 2 und 5% CO 2) für mindestens 30 min unter Erwärmen auf einem Aufzeichnungstemperaturniveau (~ 35 ° C). Den pH-Wert auf 7,4 mit NaHCO 3, während immer noch sprudelt. </li><li> Tauen Sie den internen Pipettenlösung während der Präparation. Nachdem es aufgetaut ist, fügen 0,01% Sulforhodamin B für die intrazelluläre Färbung. </li></ol></li></ol> 3. Netzhaut Dissection <ol><li> In einem dunklen Raum, einschläfern der Maus unter Verwendung von Kohlendioxid und bilaterale Pneumothorax. Nach 2 - 3 min mit dem Kohlendioxid, wird die Maus das Bewusstsein verlieren. </li><li> Wenn die Maus reagiert nicht mehr auf einem Schwanz kneifen, schnell die Augen enukleiert und legen Sie sie in einem kalten Sektions Lösung in einer Kunststoffschale. Führen Sie die bilateralen Pneumothorax zum Tode des Tieres sicherzustellen. Schnell speichern das Auge in die Schüssel in der Dunkelheit ein. Verwenden Sie eine Infrarot-Viewer, um das Verfahren in der Dunkelheit zu führen. HINWEIS: Alternativ Isofluran, Enthauptung oder Genickbruch für Euthanasie verwendet werden. Die Methode der Euthanasie sollte die Leitlinien des National Institute of Health (NIH) und / oder der Institutional Animal Care und Verwenden Ausschuss folgen. </li><li> In einem dunklen Raum, stellen Sie den Binokular und Infrarotbeleuchtung. Unter dem Mikroskop über ein Auge aus seiner Schale in eine 10 cm Plastikschale wit dem Sezieren Lösung gekühlt. Kontinuierlich Blase Sauerstoff durch das Rohr auf dem Boden des Kunststoffschale gegeben. Die Durchflussvolumen, damit es nicht zu gering, aber sicherzustellen, dass sie nicht hoch genug ist, um die Dissektion zu stören. </li><li> Hornhaut und Linse entfernen: <ol><li> Einen Einschnitt auf der Oberseite der Hornhaut mit einem mikrochirurgischen Messer. Halten des Sehnervs am anderen Ende des Augapfels verhindert das Auge bewegt. Erweitern Sie den Schnitt mit einem Paar chirurgische Schere. Schneiden Sie die Hornhaut mit einigen Lederhaut um ihn herum. </li><li> Schnappen Sie sich die Linse mit einer feinen Pinzette und ziehen Sie das Objektiv langsam aus. Wenn der Glaskörper zu klebrig ist, um die Linse zu entfernen, schneiden Sie den Glaskörper mit einer Schere. </li><li> Sobald die Augenmuschel gemacht wird, sanft gießen Sie kaltes, sauerstoff Sezieren Lösung in die Augenmuschel mit einem kleinen Transferpipette. HINWEIS: Der Augapfel schnell entleert, nachdem ein Einschnitt in der Hornhaut vorgenommen. Es ist wichtig, die Form des Auges, wie Verformen des Auges während dieser Pro aufrechtzuerhaltenverfahren wird die Netzhaut beschädigen und zu einer Netzhautablösung. </li></ol></li><li> Die Ermittlung der dorsalen und ventralen Seiten der Netzhaut: HINWEIS: Aufgrund der ungleichen Verteilung der grünen und der ultravioletten (UV) Kegel 18, Identifizierung einer bestimmten Netzhautbereich ist wichtig für die Lichtreaktion Aufnahmen. <ol><li> Identifizieren Sie die Linie, die über die Netzhaut-Augenmuschel vorbei an der Papille. Diese Linie verläuft meist auf der Bauchseite des Sehnerven; die Seite inklusive der Sehnerv ist die Rückenseite, während der andere ventral 19. Sehen Sie dieses Wahrzeichen deutlich unter Infrarot Zuschauer. </li><li> Je nach dem Zweck des Versuchs, einen Schnitt an der unnötige Neben. Dies kann hilfreich nach Trennen der Netzhaut von der Augenmuschel ist. </li></ol></li><li> Glaskörperentfernung: <ol><li> Optional inkubieren Sie die Augenmuschel mit Hyaluronidase (0,5 mg / ml) für 15 min. </li><li> Entfernen Sie den Glaskörper mit extra feinen Pinzette. Der Glaskörper ist in erster Linieauf dem Boden und der Außenkante der Okularmuschel angebracht. Entfernen Sie sie ohne sie zu berühren oder stoßen die Netzhaut. Da die Glaskörper kann fest an der Augenmuschel befestigt werden, entfernen Sie vorsichtig, um eine Netzhautablösung von der Augenmuschel zu vermeiden. Weiter zu ziehen, bis keine Spannung erreicht ist. Anmerkung: Dieser Schritt ist besonders wichtig für Sezieren der Mausretina; es ist jedoch kein Problem für den Netzhäuten von anderen Spezies, wie Ratten und Salamander. </li></ol></li><li> Isolieren Sie die Netzhaut von der Augenmuschel. Besorgen Sie sich die Lederhaut und sanft ziehen Sie die Netzhaut unter Verwendung der Rückseite der Zange. HINWEIS: Whole-Mount-Netzhaut-Präparate können aus der isolierten Netzhaut durchgeführt werden. Stellen Sie drei vor vier Schnitte an den Kanten und glätten die Netzhaut, oder schneiden Sie die Netzhaut in mehrere Stücke. </li><li> Schneiden Sie die Netzhaut und entsorgen Sie die unnötige Hälfte der Netzhaut (Schritt 3.5). Im gleichen Kunststoffschale, schneiden Sie die restlichen Netzhaut in zwei Stücke. Für jedes Stück der Netzhaut, schneiden Sie die Ränder um ein zu machenNetzhaut-Platte, und schneiden Sie die Falzkanten. </li><li> Übertragen einer Netzhaut-Platte auf eine Glasplatte mit einem großen Transferpipette (~ 2 ml). Saugen Sie die überschüssige Lösung mit einer kleinen Pipette, mit einem Stück Filterpapier, um die Platte zu glätten, und dann schnell legen Sie ein Stück der Filtermembran auf der Oberseite des Gewebes. <ol><li> Mit einem kleinen Transfer Pipette einen Tropfen Sezieren Lösung auf der Filtermembran. Warten Sie ca. 15 Sekunden, bis die Lösung erstreckt sich über der Membran. Während dieser Zeit wird das Netzhautgewebe, um die Filtermembran haften. </li><li> Pour mehr kalte Sezieren Lösung unter und um den Filter. Das Retinagewebes mit der Filtermembran wird schwimmen. Besorgen Sie sich die Filtermembran mit einer Pinzette und legen Sie sie in der Schneidkammer. Gießen Sie sezieren Lösung über das Gewebe, und speichern Sie die Zubereitung in der Dunkelheit ein. HINWEIS: Das hier beschriebene Verfahren ist von entscheidender Bedeutung. Es ist wichtig, diese Schritte schnell durchzuführen, um sicherzustellen, daß die Netzhautgewebe klebt an den Filter memBrane und trocknet nicht aus. </li></ol></li><li> Mit einem Chopper, schneiden Sie das Netzhautgewebe in Scheiben schneiden. Rund 200 um Dicke machbar für Patch-Clamp-Studien. </li><li> Legen Sie eine Kunststoff-Deckglas mit Fett Schienen im Schneideraum. Übertragen einer Netzhaut-Scheibe auf der Oberseite der Fett Schienen auf einer Kunststoffdeckglas. Drehen Sie die Scheibe um 90 °, so dass der Querschnitt zu sehen ist. Drücken Sie die Filtermembran nach unten auf das Deckglas, dann decken Sie die Seiten des Filterpapier mit Fett. <ol><li> Nachdem die Scheibe auf einer Kunststoffdeck immobilisiert, schnappen Sie das Deckglas und überträgt es auf einer 35 mm Kunststoffschale. Gießen Sie kaltes Sezieren Lösung auf die Netzhaut-Vorbereitung und tauchen die Vorbereitung. Bewahren Sie die Schale in den dunklen Kasten, der kontinuierlich mit Sauerstoff angereichert werden soll. ANMERKUNG: Das gesamte Verfahren muss sorgfältig durchgeführt, so dass die Filtermembran nicht deformiert werden. Andernfalls wird der retinaler Scheibe leicht trennt von der Filtermembran. </li></ol></li></ol><ol><li> Aufnahme der Zubereitung: <ol><li> Prime die Perfusionsschläuche mit Ames &#39;Medium. Lassen Sie alle Blasen passieren beim Füllen der Schläuche. </li><li> Stellen Sie Patch-Clamp-Aufzeichnung Pipetten mit einem Abzieher. Um eine Pipette mit der Pipettenlösung, Sprung in die Rückseite einer Pipette 1 min zu füllen, bis die Pipettenspitze aufgefüllt. Als nächstes füllen ~ 1/3 der Pipette mit einer Mikropipette Füllstoff. Bewahren Sie die Pipette in einem feuchten Pipettenfeld. </li><li> Schalten Sie die Geräte für Patch-Clamp-Aufnahmen; einschließlich des Computers, Verstärker, CCD-Kamera und Mikroskop. </li></ol></li><li> Schalten Sie die Raumbeleuchtung. Setzen Sie ein Netzhaut-Schnittpräparat auf den Mikroskoptisch Kammer mit einem Infrarot-Betrachter. Nachdem es immobilisiert ist, beginnen kontinuierliche Perfusion. Stellen Sie die Perfusion Temperatur bei 33 bis 37 ° C. </li><li> Sehen Sie sich die Scheibenoberfläche mit der CCD-Kamera. Konzentrieren Sie sich auf die Zielposition, wo das Ziel cell Arten befinden. Wählen Sie eine gesund aussehende Zelle für Patch-Clamp-Aufzeichnung (dh es sollte glatt aussehende Oberfläche und eine gute Zellform haben). </li><li> Legen Sie eine Aufnahme Pipette in einer Pipettenhalter. Voraus die Pipette auf die Schnittpräparat. Wenn es in der Nähe der Schnittpräparat (~ 2 mm oben) finden die Spitze der Pipette mit dem Mikroskop. Sobald der Pipettenspitze ist unter dem Mikroskop sichtbar ist, bewegen Sie die Spitze nach unten in Richtung der Zielzelle. </li><li> Stellen Sie den Verstärker. Stellen Sie die Pipette bei 0 mV / pA 0 (Nullstellung) und starten Sie einen kontinuierlichen elektrischen Impuls von ca. 5 mV bei ~ 10 Hz. Prüfen Sie die Pipette Widerstand; idealen Widerstand zwischen 5 und 10 M für die meisten Netzhautneuronen. </li><li> Starten Ausblasen von der Spitze. Verwenden Sie ein Mundstück, oder verwenden Sie eine Spritze, um Überdruck anzuwenden, wenn die Pipette taucht in der Badlösung. Kontinuierlich Ausblasen der internen Lösung, bis die Spitze ist auf der Oberfläche der Zielzelle. <ol><li> Wenn der Überdruck macht einen kleinen Grübchen auf the Zelloberfläche, voranzutreiben die Spitze leicht und Stop Ausblasen. Prüfen Sie die Pipette Widerstand, der zu erhöhen sollte. Wenn es ständig zu, lassen Sie es und Überwachung der Resistenz bis&gt; 1 G (Giga) erreicht. Wenn der Widerstand nicht spontan zu erhöhen, leicht negativen Druck ausüben, bis es eine undurchlässige. </li></ol></li><li> Nach dem Gigaseal erreicht ist, ändern Sie die Haltepotential auf -70 mV. Dann intermittierenden negativen Druck, um die Membran im Inneren der Pipettenspitze Bruch anzuwenden. Wenn der Gesamtzellkonfiguration hergestellt ist, kann die Pipettenwiderstand zwischen 500 MOhm und 1 GOhm, und der kapazitive Strom zu sehen sein. Manchmal spontane postsynaptischen Ströme beobachtet werden kann. </li><li> Notieren Sie sich die IV-Beziehung von -80 bis +40 mV. Verschiedene Typen von spannungsabhängigen Kanäle aktiviert je nach Zelltyp. </li><li> Nehmen Licht-evozierte synaptische Ströme oder Spannungen. </li></ol>

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The authors have nothing to disclose.

Good recordings can only be achieved with good retinal preparations and well-designed patch clamp setups. Although all the steps described above are important, the discussion highlights some critical steps both for the dissection and recordings. 
For dissection, two things are especially important: cooling and oxygenation. After enucleating the eye, quickly remove the front part of the eye in a dissecting chamber with oxygen-bubbled, cooled dissecting solution, and pour cold solution into the eye-cup. When the retinal tissue is in the dissecting chamber, cold solution should be poured on the tissue every 10 min. The tissue should be sliced quickly without pulling or poking the retinal tissue during the dissection, which could easily damage its structure. Using good dissecting tools (sharp forceps and scissors) is also crucial. A chopper is equally as effective as a vibratome because it allows for speedy maneuvering. Extra-fine forceps should be used to remove vitreous matter as efficiently as possible because residual vitreous damages the retinal structure when slicing the tissue.
Beginners tend to damage the retina when removing the lens. The retina can detach from the eye-cup, or can even be accidentally removed with the lens. To avoid this issue, the sclera can be cut around the Ora Serrata which is located close to the extra-ocular muscle attachment point. An eye-cup with a wide opening makes it easier to remove the lens. Also, it is important not to deform the eye when removing the front portion. Another critical point is fixing the retinal slice onto a coverslip. This can be facilitated by practicing rotating the filter paper without distorting it to prevent the retinal slice from coming off during the experiment. 
A sturdy, well-built rig is crucial for patch clamp recordings. It must be free of electric noise, vibration, and pipette drifting. In addition, light response can be recorded in dark conditions to prevent desensitization. Our cage is made with a metal frame and aluminum panels with black plastic drapes that act as an electric shield while also completely blocking light from entering the cage; computer monitor illumination will not affect the retinal preparation. Use a remote controller for the pipette manipulator and close the cage while setting up the patch clamp configuration, which also avoid illuminating the preparation.
Although we have been tried to simplify the procedures involved, patch clamp study is still a difficult experiment that requires extensive knowledge and training. An automated patch clamp has been developed by some groups20; this system is applicable to dissociated cells and might be an efficient technique for pharmacological screenings. 
In summary, patch clamp recording in dark-adapted retinal slice preparations is a useful technique to investigate visual information processing. Compared to other methods, patch clamp offers better temporal and spatial resolution. Additional technical innovations will make the patch clamp method more user-friendly in the near future.

The retina is one of the unique parts of the nervous system. As an accessible part of the brain, its synaptic architecture has been well characterized. In addition, the functions of this neural network can be examined with a physiological stimulus: light. If the retinal tissue is isolated in a dark room with appropriate procedures, neurons in the tissue will respond to light. This preparation has been used to study visual signal processing and elucidate various synaptic mechanisms and neural network functions, as well as disease mechanisms. 
Light responses in retinal neurons have been recorded for decades. Early studies used sharp electrodes to make intracellular recordings from mudpuppy retinal neurons1. In the 1980s, the patch clamp technique was invented2, and soon became a popular method among vision researchers3,4. Single cell recordings from lower vertebrates, including mudpuppy and fish retinal neurons, were popular methods that contributed to the elucidation of visual signal processing mechanisms5,6. 
After genetic mutation techniques were developed, the mouse retina became a more popular model for vision researchers7-9. The mammalian retina is more attractive than that of lower vertebrates because it is evolutionarily closer to the human retina, and there is an opportunity to use disease models. However, mouse retinal cells are small and fragile10, and making retinal preparations and conducting patch clamp recordings in a dark room is challenging. As technology has improved, diverse approaches have become available to study visual signaling mechanisms such as imaging studies11 and the electroretinogram (ERG)12. Nevertheless, single cell recording with the patch clamp method is still important because it is highly temporally and spatially sensitive compared to other methods. Therefore, we have continuously conducted patch clamp recordings and improved our methods to investigate visual signal processing in mouse retinal slice preparations13-15.
In this video tutorial, the protocols are presented with important tips. Good recordings can only be achieved with good preparation. Practicing animal dissection and building a sturdy patch clamp rig will enable most researchers to achieve successful recordings.

<table border="0" cellpadding="0" cellspacing="0" width="886">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Name of Material/ Equipment</td>
			<td style="width:145px;">Company</td>
			<td style="width:203px;">Catalog Number</td>
			<td style="width:323px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">mice (28-60 days old, male)</td>
			<td style="width:145px;">Jackson laboratory</td>
			<td style="width:203px;">C57BL/6J strain</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Ames&#39; medium powder</td>
			<td style="width:145px;">Sigma</td>
			<td style="width:203px;">A1420</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Stereo microscope</td>
			<td style="width:145px;">Nikon</td>
			<td style="width:203px;">SMZ745</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">dissecting tool_forceps</td>
			<td style="width:145px;">Dumont</td>
			<td style="width:203px;">#4, #5, #55</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">dissecting tool_scissors</td>
			<td style="width:145px;">Roboz</td>
			<td style="width:203px;">RS-5605</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">dissecting tool_surgery knife</td>
			<td style="width:145px;">Surgistar</td>
			<td style="width:203px;">7514</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">razor blade (for chopper)</td>
			<td style="width:145px;">EMS</td>
			<td style="width:203px;">71970</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">chopper</td>
			<td style="width:145px;">handmade</td>
			<td style="width:203px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">infrared viewer</td>
			<td style="width:145px;">Night Owl Optics</td>
			<td style="width:203px;">NOBG1</td>
			<td>It shows bright view.&#160; Focusing small objects is an issue.</td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:216px;">infrared pocket scopes</td>
			<td style="width:145px;">B.E. Meyers</td>
			<td style="width:203px;">OWL Gen 3 NV pocketscope</td>
			<td>excellent view</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">puller</td>
			<td style="width:145px;">Sutter</td>
			<td style="width:203px;">P-1000</td>
			<td>excellent.&#160; Make consistent size pipettes.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">dark box</td>
			<td style="width:145px;">Pelican</td>
			<td style="width:203px;">dark box</td>
			<td>excellent</td>
		</tr>
		<tr height="67">
			<td height="67" style="height:67px;width:216px;">patch clamp system</td>
			<td style="width:145px;">Scientifica</td>
			<td style="width:203px;">slice scope 2000</td>
			<td style="width:323px;">Excellent setup.&#160; Most key components are included in one package.&#160; Micromanipulators are excellent.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">amplifier</td>
			<td style="width:145px;">Molecular Devices</td>
			<td style="width:203px;">multiclamp 700B</td>
			<td>Excellent and easy control.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">acquiring software</td>
			<td style="width:145px;">Molecular Devices</td>
			<td style="width:203px;">pClamp software</td>
			<td>Excellent and easy control.</td>
		</tr>
		<tr height="28">
			<td height="28" style="height:28px;width:216px;">light source (LED)</td>
			<td style="width:145px;">Cool LED</td>
			<td style="width:203px;">pE-2 4 channel system</td>
			<td>Excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">CCD camera</td>
			<td style="width:145px;">Q-imaging</td>
			<td style="width:203px;">Retiga 2000</td>
			<td>Excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Faraday cage</td>
			<td style="width:145px;">handmade</td>
			<td style="width:203px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

recording light-evoked postsynaptic responses in neurons in dark-adapted, mouse retinal slice preparations using patch clamp techniques

Die Netzhaut ist das Tor zu den visuellen Systems. Zur visuellen Signalverarbeitung Mechanismen zu verstehen, untersuchen wir retinale neuronalen Netzwerkfunktionen. Retinalen Neuronen in dem Netzwerk umfassen eine Reihe von Subtypen. Mehr als 10 Subtypen der bipolaren Zellen, Ganglienzellen und Amakrinzellen wurden durch morphologische Studien identifiziert. Mehrere Subtypen von retinalen Neuronen sind gedacht, um unterschiedliche Merkmale der visuellen Signalisierung, wie Bewegung und Farbe zu codieren, und bilden mehrere Nervenbahnen. Jedoch sind die funktionalen Rollen von jedem Neuron in der visuellen Signalverarbeitung nicht vollständig verstanden. Die Patch-Clamp-Methode ist hilfreich, diese grundlegende Frage zu beantworten. Hier ist ein Protokoll, um Licht-evozierte synaptische Antworten in Maus retinalen Neuronen mit Patch-Clamp-Aufnahmen im dunkeladaptierten Bedingungen aufnehmen wird. Die Maus Augen sind dunkeladaptierten O / N, und Netzhautschnittpräparate werden in einem dunklen Raum mit Infrarotbeleuchtung und Zuschauer seziert. Infrarotlicht nichtAktivieren Maus Photorezeptoren und damit bewahrt ihr Licht Reaktionsfähigkeit. Patch-Clamp verwendet, um Licht hervorgerufenen Reaktionen in retinalen Neuronen aufzuzeichnen. Ein Fluoreszenzfarbstoff während Aufnahmen auf neuronale morphologischen Subtypen charakterisieren injiziert. Dieses Verfahren ermöglicht es, die physiologischen Funktionen von jedem Neuron in der Maus-Retina zu bestimmen.

Eine repräsentative Schnittpräparat ist in Abbildung 1 dargestellt. Die Schnittpräparat in einer geraden Position, welche Photorezeptoren Ganglionzellen in einer flachen Oberfläche, und keine Ablösung vom Filterpapier. Wenn eine Scheibe geneigt ist, ist nur ein Teil der Vorbereitung im Fokus, was es schwierig macht, eine geeignete Zelle für Patch-Clamp identifizieren. Für Aufnahmen ist es wichtig, eine gute Soma, die in der Regel hat eine glänzende Oberfläche, eine gleichmäßige runde Form, und keine sichtbaren dunklen Plaques zu wählen. Nachdem der Ganzzellkonfiguration vorgenommen wurde, wurde die Scheibe von Hintergrundlicht zu einem Schritt-Licht (2) ausgesetzt und dann. Licht evozierten exzitatorischen postsynaptischen Potentiale (L-EPSPs) wurden in Reaktion auf einen Schritt lichtinduzierten (Timing ist gelb). Die Spitzenamplitude und die Zerfallszeit variiert in Abhängigkeit von dem Zelltyp und Untertyp. Eine Ganglienzelle erzeugte Aktionspotentialen als Reaktion auf helles Stimuli (Abbildung 2 D &amp; E). Die cell-Typ und seine morphologische Subtypen wurden von Sulforhodamin Kennzeichnung nach physiologischen Aufnahmen (Abbildung 2, rechts) enthüllt. <img alt="Figur 1" src="/files/ftp_upload/52422/52422fig1highres.jpg" width="650" /> Abbildung 1. Netzhautschnittpräparate. (A) Eine Netzhautschnittpräparat mit einer 10X-Objektiv betrachtet. Eine Netzhautschnittpräparat (oben) wurde in ein Stück Filterpapier (unten) angebracht ist. (B) Ein Vierbildzusammenstellung, die eine Netzhautschnittpräparat mit einem 60X Ziel angesehen. Jede Zelle Schicht deutlich zu beobachten (ONL: äußere Körnerschicht, OPL: plexiformen Schicht, INL: innere Körnerschicht, IPL: inneren plexiformen Schicht, GCL: Ganglienzellschicht) <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52422/52422fig1highres.jpg" target="_blank">Bitte klicken Sie hier, um eine größere Version dieses Bild anzuzeigen.</a> Abbildung 2. Licht hervorgerufenen exzitatorischen postsynaptischen Potentiale (L-EPSPs) von Netzhautneuronen. Step-Licht hervorgerufen L-EPSPs (links). Die Lichtintensität war 30-60% Weber Kontrast. Hintergrundlicht Anpassung Ebene war 4 x 10 4 Photonen / um 2 / s. Sulforhodamin B wurde im Patch-Clamp-Aufzeichnung, die aufgezeichneten Neuronen zu visualisieren (rechts) eingespritzt. Eine Aufzeichnung Pipette wurde noch mit dem Soma (A) L-EPSPs und Sulforhodamin Färbung von einem ON Kegel bipolaren Zelle angebracht. (B) OFF Kegel bipolaren Zelle. (C) Amakrinzell. (D) AUF Ganglienzellen. (E) OFF Ganglionzellen. Maßstabsbalken zeigt an, 2 oder 5 mV festgestellt. Lichtstimulation wurde für 1 Sekunde angewendet. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52422/52422fig2highres.jpg" target="_blank">Bitte klicken Sie hier</a>um eine größere Version dieser Figur zu sehen.

Watch this Scientific Journal Video about Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques at JoVE.com

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

Recording Licht-evozierte Postsynaptische Antworten in Neuronen im Dunkel-adaptierten, Maus retinalen Scheibe Vorbereitungen Verwendung Patch-Clamp-Techniken

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. ...

recording-light-evoked-postsynaptic-responses-neurons-dark-adapted

Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.4K Aufrufe.  Wayne State University School of Medicine. We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

Serie: Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

Analyzing Responses of Mouse Olfactory Sensory Neurons Using the Air-phase Electroolfactogram Recording

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and other social interactions. The electroolfactogram (EOG) recording is an informative, easy to conduct, and reliable method to assay olfactory function at the level of the olfactory epithelium. Since the 1956 description of the EOG by Ottoson in frogs1, the EOG recording has been applied in many vertebrates including salamanders, rabbits, rats, mice, and humans (reviewed by Scott and Scott-Johnson, 2002, ref. 2). The recent advances in genetic modification in mice have rekindled interest in recording the EOG for physiological characterization of olfactory function in knock-out and knock-in mice. EOG recordings have been successfully applied to demonstrate the central role of olfactory signal transduction components3-8, and more recently to characterize the contribution of certain regulatory mechanisms to OSN responses9-12.
Odorant detection occurs at the surface of the olfactory epithelium on the cilia of OSNs, where a signal transduction cascade leads to opening of ion channels, generating a current that flows into the cilia and depolarizes the membrane13. The EOG is the negative potential recorded extracellularly at the surface of the olfactory epithelium upon odorant stimulation, resulting from a summation of the potential changes caused by individual responsive OSNs in the recording field2. Comparison of the amplitude and kinetics of the EOG thus provide valuable information about how genetic modification and other experimental manipulations influence the molecular signaling underlying the OSN response to odor. 
Here we describe an air-phase EOG recording on a preparation of mouse olfactory turbinates. Briefly, after sacrificing the mouse, the olfactory turbinates are exposed by bisecting the head along the midline and removing the septum. The turbinate preparation is then placed in the recording setup, and a recording electrode is placed at the surface of the olfactory epithelium on one of the medial turbinates. A reference electrode is electrically connected to the tissue through a buffer solution. A continuous stream of humidified air is blown over the surface of the epithelium to keep it moist. The vapor of odorant solutions is puffed into the stream of humidified air to stimulate the epithelium. Responses are recorded and digitized for further analysis.

The electroolfactogram (EOG) recording is an informative, easy-to-conduct, and reliable way of assessing olfactory function at the level of the olfactory epithelium. This protocol describes a recording setup, mouse tissue preparation, data collection, and basic data analysis.

Analysieren Antworten der Maus Riechzellen Mit der Air-Phase Electroolfactogram Recording

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and ...

Forschung

Neurowissenschaften

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Patch Clamp Messungen von Mouse Netzhautneuronen in einem Dark-Scheiben angepasst Vorbereitung

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

Die Transfektion von Maus retinalen Ganglienzellen durch In vivo Elektroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 &mu;m diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Here we describe a protocol for the preparation of agar-embedded retinal slices that are suitable for electrophysiology and Ca2+ imaging. This method allows one to study ribbon-type synapses in retinal microcircuits using direct patch-clamp recordings of single presynaptic nerve terminals.

Patch-Clamp Kapazitätsmessungen und Ca 2 + Imaging bei Single Nervenendigungen in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

Imaging neuronalen Antworten in Scheiben Zubereitungen von Vomeronasalorgan Ausdruck eines genetisch kodierte Kalzium-Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

Retrograde Fluoreszenzmarkierung Ermöglicht Gezielte Extracellular Einzel-Einheit Aufnahme von identifizierten Neuronen<em&gt; In vivo</em

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Perforated Patch-clamp Recording of Mouse Olfactory Sensory Neurons in Intact Neuroepithelium: Functional Analysis of Neurons Expressing an Identified Odorant Receptor

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of olfactory-driven behaviors and their modulation. These coding properties depend heavily on the initial interaction between odor molecules and the olfactory receptor (OR) expressed in the OSNs. The identity, specificity and ligand spectrum of the expressed OR are critical. The probability to find the ligand of the OR expressed in an OSN chosen randomly within the epithelium is very low. To address this challenge, this protocol uses genetically tagged mice expressing the fluorescent protein GFP under the control of the promoter of defined ORs. OSNs are located in a tight and organized epithelium lining the nasal cavity, with neighboring cells influencing their maturation and function. Here we describe a method to isolate an intact olfactory epithelium and record through patch-clamp recordings the properties of OSNs expressing defined odorant receptors. The protocol allows one to characterize OSN membrane properties while keeping the influence of the neighboring tissue. Analysis of patch-clamp results yields&#160;a precise quantification of ligand/OR interactions, transduction pathways and pharmacology, OSNs&#39; coding properties and their modulation at the membrane level.&#160;

Analyzing the physiological properties of olfactory sensory neurons still faces technical limitations. Here we record them through perforated patch-clamp in an intact preparation of the olfactory epithelium in gene-targeted mice. This technique allows the characterization of membrane properties and responses to specific ligands of neurons expressing defined olfactory receptors.

Perforated-Patch-Clamp-Aufzeichnung der Maus Riechzellen in Intact Neuroepithel: Funktionelle Analyse von Neuronen, die eine bestimmte Geruchsrezeptor

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-Cell-Patch-Clamp-Aufnahmen in Hirnschnitten

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Patch Clamp Recording of Starburst Amacrine Cells in a Flat-mount Preparation of Deafferentated Mouse Retina

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic network, electrophysiological recordings are frequently used to study connections among individual neurons. We have optimized a flat-mount preparation for patch clamp recording of genetically marked neurons in both GCL (ganglion cell layer) and INL (inner nuclear layer) of mouse retinas. Recording INL neurons in flat-mounts is favored over slices because both vertical and lateral connections are preserved in the former configuration, allowing retinal circuits with large lateral components to be studied. We have used this procedure to compare responses of mirror-partnered neurons in retinas such as the cholinergic starburst amacrine cells (SACs).

This protocol demonstrates how to perform whole-cell patch clamp recording on retinal neurons from a flat-mount preparation.

Patch-Clamp-Aufzeichnung von Starburst-Amakrinzellen in einer Wohnung Montage Herstellung von Deafferentated Maus Retina

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic ...

Slice Patch Clamp Technique for Analyzing Learning-Induced Plasticity

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze motor-learning induced plasticity, we trained rats using an accelerated rotor rod task. Rats performed the task 10 times at 30-s intervals for 1 or 2 days. Performance was significantly improved on the training days compared to the first trial. We then prepared acute brain slices of the primary motor cortex (M1) in untrained and trained rats. Current-clamp analysis showed dynamic changes in resting membrane potential, spike threshold, afterhyperpolarization, and membrane resistance in layer II/III pyramidal neurons. Current injection induced many more spikes in 2-day trained rats than in untrained controls.
To analyze contextual-learning induced plasticity, we trained rats using an inhibitory avoidance (IA) task. After experiencing foot-shock in the dark side of a box, the rats learned to avoid it, staying in the lighted side. We prepared acute hippocampal slices from untrained, IA-trained, unpaired, and walk-through rats. Voltage-clamp analysis was used to sequentially record miniature excitatory and inhibitory postsynaptic currents (mEPSCs and mIPSCs) from the same CA1 neuron. We found different mean mEPSC and mIPSC amplitudes in each CA1 neuron, suggesting that each neuron had different postsynaptic strengths at its excitatory and inhibitory synapses. Moreover, compared with untrained controls, IA-trained rats had higher mEPSC and mIPSC amplitudes, with broad diversity. These results suggested that contextual learning creates postsynaptic diversity in both excitatory and inhibitory synapses at each CA1 neuron.
AMPA or GABAA receptors seemed to mediate the postsynaptic currents, since bath treatment with CNQX or bicuculline blocked the mEPSC or mIPSC events, respectively. This technique can be used to study different types of learning in other regions, such as the sensory cortex and amygdala.

The slice patch clamp technique is an effective method for analyzing learning-induced changes in the intrinsic properties and plasticity of excitatory or inhibitory synapses.