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

Déclaration éthique: procédures impliquant des sujets animaux ont été approuvés par le Comité institutionnel de protection des animaux et l&#39;utilisation (IACUC) à l&#39;Université Wayne State. 1. Préparation de la solution expérimentale <ol><li> Préparer la solution de dissection 1 jour à 1 semaine avant l&#39;expérience réelle. Utilisez une solution tampon HEPES pour la dissection de la rétine raison de sa capacité tampon forte à des températures inférieures 16. Mélanger tous les produits chimiques comme suit (en mM): 115 NaCl, KCl 2,5, 2,5 CaCl2, 1,0 MgCl2, 10 HEPES, 28 glucose. Ajuster le pH à 7,4 avec NaOH. Conserver la solution dans un réfrigérateur jusqu&#39;à 1 semaine. </li><li> La solution d&#39;enregistrement est un milieu d&#39;Ames, qui est un liquide céphalo-rachidien artificiel (aCSF) conçu pour des préparations 17 de la rétine. Le jour de l&#39;expérience, peser la poudre de Ames (4,4 g pour 500 ml de solution) dans un tube. En outre, peser NaHCO 3 (0,95 g pour 500 ml de solution) et mélanger avec le Ala poudre de mes. </li><li> Préparer la solution de la pipette intracellulaire pour les enregistrements de patch-clamp par le jour de l&#39;expérience. Mélanger tous les produits chimiques comme suit (en mm): 111 K-gluconate, 1,0 CaCl2, 10 HEPES, 1,1 EGTA, 10 NaCl, 1,0 MgCl 2, 5 ATP-Mg et 1,0 GTP-Na. Ajuster le pH à 7,2 avec KOH. On filtre la solution de la pipette avec un filtre à seringue classique. Assurez ~ 500 aliquotes et les stocker dans un congélateur ou un congélateur. </li></ol> 2. Préparation pour la Journée de l&#39;expérience <ol><li> La nuit avant l&#39;expérience, placer une souris dans une cage (C57BL / 6J ou des antécédents similaires, 4-8 semaines, mâle) dans un espace sombre pour O / N adaptation à l&#39;obscurité. </li><li> Préparation pour la dissection: <ol><li> Remplissez dissection solution (de 100 à 200 ml) dans un récipient en verre, placez-le sur la glace, et la bulle d&#39;oxygène pendant au moins 10 min. </li><li> Lancer l&#39;oxygénation de la boîte noire pour le stockage de la préparation de la rétine. </li><li> Préparer des lamelles de plastique avec des rails de graisse pour repréparations de tranche intestinaux et placez chacun d&#39;eux dans un plat en plastique de 35 mm. </li><li> Fixez une nouvelle lame de rasoir à un chopper (trancheuse de tissu). </li><li> Couper une membrane de filtre en deux moitiés. Il se agit pour le tranchage de la rétine. </li><li> Alignez les outils de dissection et pipettes de transfert près de la loupe binoculaire. La zone de travail doit être bien organisé de sorte que chaque outil peut être facilement accessible dans l&#39;obscurité. </li></ol></li><li> Préparation pour les enregistrements de patch-clamp: <ol><li> Dissoudre la poudre de Ames et NaHCO 3 dans l&#39;eau distillée (Ames 4,4 g et 0,95 g / NaHCO 3 500 ml ddH 2 O). Le milieu de la bulle Ames mélangé avec de l&#39;oxygène (95% O 2 et 5% CO 2) pendant au moins 30 minutes tout en chauffant à un niveau de température d&#39;enregistrement (~ 35 ° C). Ajuster le pH à 7,4 avec NaHCO 3 tout en bouillonnant. </li><li> Décongeler la solution de pipette interne lors de la dissection. Après il est décongelé, ajouter 0,01% sulforhodamine B pour la coloration intracellulaire. </li></ol></li></ol> 3. rétine Dissection <ol><li> Dans une pièce sombre, euthanasier la souris en utilisant du dioxyde de carbone et de pneumothorax bilatéral. Après 2 à 3 min d&#39;utiliser le dioxyde de carbone, la souris se perdre conscience. </li><li> Lorsque la souris ne répond plus à un pincement de la queue, énucléer rapidement les yeux et les placer dans une solution froide de dissection dans un plat en plastique. Effectuer pneumothorax bilatéral pour assurer la mort de l&#39;animal. Stocker rapidement l&#39;œil dans le plat dans la boîte noire. Utilisez une visionneuse infrarouge pour mener la procédure dans l&#39;obscurité. REMARQUE: Sinon, l&#39;isoflurane, la décapitation ou la dislocation cervicale peuvent être utilisés pour l&#39;euthanasie. La méthode d&#39;euthanasie doit suivre les directives de l&#39;Institut national de la santé (NIH) et / ou du Comité de protection des animaux et l&#39;utilisation institutionnelle. </li><li> Dans une pièce sombre, régler le microscope de dissection et l&#39;éclairage infrarouge. Sous le microscope, transférer un œil de son plat dans un plat en plastique de 10 cm wvec la solution dissection refroidi. En continu l&#39;oxygène bulle à travers le tubage placée sur le fond de la cuvette en plastique. Réglez le volume d&#39;écoulement de sorte qu&#39;il ne est pas trop faible, mais assurez-vous qu&#39;il ne est pas suffisamment élevée pour perturber la dissection. </li><li> Cornée et le cristallin retrait: <ol><li> Faire une découpe sur le dessus de la cornée avec une lame de micro-chirurgical. En tenant le nerf optique à l&#39;autre extrémité du globe oculaire de l&#39;œil empêche de se déplacer. Prolonger la coupe à l&#39;aide d&#39;une paire de ciseaux chirurgicaux. Découpez la cornée avec un certain sclérotique autour d&#39;elle. </li><li> Prenez l&#39;objectif avec une pince fine et tirer lentement sur la lentille. Si le corps vitré est trop collante pour retirer le cristallin, le vitré couper avec une paire de ciseaux. </li><li> Une fois l&#39;œilleton est fait, verser doucement solution froide, oxygénée dissection dans l&#39;œilleton avec une petite pipette de transfert. REMARQUE: le globe oculaire se dégonfle rapidement après une incision est faite dans la cornée. Il est important de maintenir la forme de l&#39;oeil que l&#39;œil se déformer au cours de cette proprocédure peut endommager la rétine et provoquer un décollement de la rétine. </li></ol></li><li> Identifier les dorsales et ventrales côtés de la rétine: NOTE: En raison de la répartition inégale du vert et de l&#39;ultraviolet (UV) cônes 18, indiquant une zone spécifique de la rétine est important pour les enregistrements de réponse à la lumière. <ol><li> Identifier la ligne traversant la coupe de la rétine passant près de la tête du nerf optique. Cette ligne passe la plupart du temps à travers la face ventrale du nerf optique; le côté comprenant le nerf optique est le côté dorsal, tandis que l&#39;autre est 19 ventral. Voir ce monument clairement sous téléspectateurs infrarouges. </li><li> Selon le but de l&#39;expérience, faire une coupe sur le côté inutile. Cela sera utile après avoir isolé la rétine de l&#39;œilleton. </li></ol></li><li> Élimination vitreuse: <ol><li> Eventuellement, incuber la œilleton avec la hyaluronidase (0,5 mg / ml) pendant 15 min. </li><li> Retirer le corps vitré avec une pince extra-fines. Le corps vitré est principalementfixée sur le fond et le bord externe de l&#39;œil-tasse. Retirez délicatement sans toucher ou piquer la rétine. Comme le corps vitré peut être étroitement attaché à l&#39;œilleton, retirez délicatement pour éviter décollement de la rétine de l&#39;œil-tasse. Continuez à tirer jusqu&#39;à ce qu&#39;aucune tension se fait sentir. NOTE: Cette étape est particulièrement importante pour disséquer la rétine de souris; cependant, ce ne est pas un problème pour les rétines d&#39;autres espèces telles que le rat ou salamandre. </li></ol></li><li> Isoler la rétine de l&#39;œil-tasse. Prenez la sclérotique et retirez délicatement la rétine en utilisant l&#39;arrière de forceps. REMARQUE: l&#39;ensemble du montage des préparations de la rétine peut être fabriqué à partir de la rétine isolée. Faire trois à quatre coupes sur les bords et aplatir la rétine, ou couper la rétine en plusieurs morceaux. </li><li> Coupez la rétine et jeter le moitié inutile de la rétine (étape 3.5). Dans le même plat en plastique, couper la rétine restante en deux morceaux. Pour chaque morceau de la rétine, couper les bords pour faire undalle rétinienne, et couper les bords de pliage. </li><li> Transfert d&#39;une dalle de la rétine sur une plaque de verre en utilisant une pipette de transfert à grande (~ 2 ml). Aspirer la solution en excès avec une petite pipette, utiliser un morceau de papier filtre pour aplatir la dalle, puis placer rapidement un morceau de la membrane du filtre sur le dessus du tissu. <ol><li> En utilisant une petite pipette de transfert, déposer une goutte de solution sur la membrane filtrante de dissection. Attendez ~ 15 sec jusqu&#39;à ce que la solution se répand sur la membrane. Pendant ce temps, le tissu rétinien se en tiendra à la membrane filtrante. </li><li> Verser la solution de dissection plus froid sous et autour du filtre. Le tissu de la rétine avec la membrane filtrante peut flotter. Saisir la membrane de filtre avec une pince et le placer dans la chambre de coupe. Verser la solution de dissection sur le tissu, et de stocker la préparation dans la boîte noire. NOTE: La procédure décrite ici est critique. Il est important d&#39;effectuer ces étapes rapidement pour se assurer que les bâtons de tissus de la rétine aux mem de filtreBrane et ne se dessèche pas. </li></ol></li><li> L&#39;utilisation d&#39;un hachoir, couper le tissu rétinien en tranches. Autour de 200 um d&#39;épaisseur est faisable pour les études de patch-clamp. </li><li> Déposer une lamelle couvre-objet en plastique avec des rails de graisse dans la chambre de coupe. Transférer une tranche de la rétine au-dessus des rails de graisse sur une lamelle en plastique. Faire tourner la tranche 90 ° de sorte que la section transversale est visible. Appuyez sur la membrane du filtre vers le bas sur la lamelle, puis, recouvrez les côtés du papier filtre avec de la graisse. <ol><li> Après la tranche est immobilisé sur une lamelle de plastique, saisir la lamelle et le transférer sur un plat en plastique de 35 mm. Verser la solution de dissection froide sur la préparation de la rétine et plonger la préparation. Stocker chaque plat dans la boîte noire, qui devrait être continuellement oxygéné. NOTE: L&#39;ensemble de la procédure qui doit être fait avec soin afin que la membrane du filtre ne est pas déformée. Dans le cas contraire, la tranche de la rétine se sépare facilement de la membrane filtrante. </li></ol></li></ol><ol><li> Enregistrement de la préparation: <ol><li> Prime les tubes de perfusion avec le milieu de Ames. Autoriser toutes les bulles de passer à travers lors du remplissage du tube. </li><li> Assurez-pipettes d&#39;enregistrement patch clamp avec un extracteur. Pour remplir une pipette avec la solution de la pipette, plongeon dans l&#39;arrière d&#39;une pipette pendant 1 min jusqu&#39;à ce que la pointe de la pipette est remblayé. Ensuite, remplissez ~ 1/3 de la pipette en utilisant une charge micro-pipette. Stocker chaque pipette dans une boîte de pipette humide. </li><li> Tournez sur l&#39;équipement pour les enregistrements de patch-clamp; y compris l&#39;ordinateur, un amplificateur, caméra CCD, et microscope. </li></ol></li><li> Éteignez la lumière ambiante. Placez une préparation de tranches de la rétine sur la chambre de platine de microscope en utilisant une visionneuse infrarouge. Après il est immobilisé, commencez perfusion continue. Réglez la température de perfusion à 33-37 ° C. </li><li> Voir la surface de tranche avec la caméra CCD. Concentrez-vous sur l&#39;emplacement cible où le CE ciblell types résident. Sélectionnez une cellule saine pour le patch clamp (ie, il devrait avoir une surface lisse d&#39;aspect et une forme bonne cellulaire). </li><li> Placer une pipette d&#39;enregistrement dans un support de pipette. Avancer la pipette à la préparation de la tranche. Quand il est proche de la préparation de tranche (~ 2 mm au-dessus), trouver la pointe de la pipette avec le microscope. Une fois la pointe de pipette est visible sous le microscope, déplacer la pointe vers le bas vers la cellule cible. </li><li> Réglez l&#39;amplificateur. Régler la pipette à 0 mV / 0 pA (réduction à zéro) et de commencer une impulsion électrique continu de ~ 5 mV à ~ 10 Hz. Vérifier la résistance de la pipette; la résistance idéale se situe entre 5 et 10 MQ pour la plupart des neurones rétiniens. </li><li> Lancer souffler de la pointe. Utilisation d&#39;un embout buccal, ou utiliser une seringue pour appliquer une pression positive lorsque la pipette plongeant dans la solution de bain. Souffler en continu la solution interne jusqu&#39;à la pointe est à la surface de la cellule cible. <ol><li> Lorsque la pression positive fait une petite fossette sur esurface de la cellule e, avancer la pointe légèrement et arrêter de souffler. Vérifier la résistance de la pipette, ce qui devrait augmenter. Se il ne cesse d&#39;augmenter, le quitter et surveiller la résistance jusqu&#39;à ce qu&#39;il atteigne&gt; 1 GQ (gigaseal). Si la résistance ne augmente pas spontanément, appliquer doucement pression négative jusqu&#39;à ce qu&#39;il devienne un gigaseal. </li></ol></li><li> Après la gigaseal est atteint, changer le potentiel de maintien de -70 mV. Ensuite, appliquer par intermittence pression négative pour rompre la membrane à l&#39;intérieur de la pointe de la pipette. Lorsque la configuration de cellule entière est faite, la résistance de la pipette peut être comprise entre 500 et 1 GQ MQ, et le courant capacitif est considérée. Parfois courants postsynaptiques spontanées peuvent être observées. </li><li> Enregistrez la relation IV -80 à 40 mV. Différents types de canaux voltage-dépendants sont activés en fonction du type de cellule. </li><li> Enregistrez lumière évoqué courants ou tensions synaptiques. </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 ...

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

La rétine est la porte d&#39;entrée du système visuel. Pour comprendre les mécanismes de traitement des signaux visuels, nous étudions les fonctions de réseau de neurones de la rétine. Neurones rétiniens dans le réseau comprennent de nombreux sous-types. Plus de 10 sous-types de cellules bipolaires, les cellules ganglionnaires, les cellules amacrines et ont été identifiés par des études morphologiques. Sous-types multiples de neurones rétiniens sont pensés pour coder des caractéristiques distinctes de signalisation visuelle, telles que le mouvement et la couleur, et de former de multiples voies neurales. Cependant, les rôles fonctionnels de chaque neurone en traitement du signal visuel sont pas pleinement compris. La méthode patch clamp est utile de répondre à cette question fondamentale. Ici, un protocole d&#39;enregistrer les réponses synaptiques lumière évoqués dans les neurones de la rétine de souris en utilisant des enregistrements de patch clamp dans des conditions sombres adapté est fourni. Les yeux de souris sont adaptés à l&#39;obscurité O / N, et les préparations de tranche de la rétine sont disséqués dans une pièce sombre en utilisant un éclairage infrarouge et les téléspectateurs. La lumière infrarouge ne est pasactiver photorécepteurs de la souris et préserve ainsi leur réactivité de lumière. Patch clamp est utilisé pour enregistrer les réponses de lumière évoquée dans les neurones de la rétine. Un colorant fluorescent est injecté pendant les enregistrements pour caractériser les sous-types morphologiques neuronales. Cette procédure permet de déterminer les fonctions physiologiques de chaque neurone dans la rétine de souris.

Une préparation représentative de tranche est représenté sur la Figure 1. La préparation de la tranche se trouve dans une position droite, montrant photorécepteurs aux cellules ganglionnaires dans une surface plane, et aucun détachement du papier filtre. Si une tranche est incliné, seule une partie de la préparation est au point, ce qui rend difficile d&#39;identifier une cellule appropriée pour le patch serrage. Pour les enregistrements, il est important de choisir un bon soma, qui a généra...

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.

Enregistrement Light-réponses évoquées dans les neurones post-synaptiques dans Dark-adaptés, Préparations rétine de la souris tranche en utilisant des techniques de patch clamp

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

Research

JoVE Journal

Neuroscience

8.3K vues.  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.

Vidéo: 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.

Réponses Analyse des souris neurones sensoriels olfactifs Utilisation de l&#39;enregistrement Electroolfactogram Air-phase

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

Recherche

Neurosciences

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.

Enregistrements Patch Clamp à partir de neurones rétiniens souris dans une préparation adaptés à l&#39;obscurité Slice

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

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 mesures de capacité et de Ca 2 + Imagerie à terminaisons nerveuses rétiniennes unique en tranches

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.

Réponses d&#39;imagerie neuronale dans les préparations d&#39;organe voméronasal Slice Exprimant un capteur de calcium codées génétiquement

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

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.

Perforé patch-clamp enregistrement des souris neurones sensoriels olfactifs dans neuroépithélium Intact: Analyse fonctionnelle des neurones exprimant un récepteur odorant Identifié

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.

Recordings cellules entières Patch-clamp en tranches cérébrales

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 Enregistrement de cellules amacrines Starburst dans un plat de montage Préparation de Deafferentated Souris 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.

Tranche Patch Clamp Technique d’analyse de plasticité induite par l’apprentissage

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze ...

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

Préparations et protocoles pour toute cellule Patch Clamp enregistrement des neurones Tectal Xenopus laevis

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...

Preparation of Acute Spinal Cord Slices for Whole-cell Patch-clamp Recording in Substantia Gelatinosa Neurons

Recent whole-cell patch-clamp studies from substantia gelatinosa (SG) neurons have provided a large body of information about the spinal mechanisms underlying sensory transmission, nociceptive regulation, and chronic pain or itch development. Implementations of electrophysiological recordings together with morphological studies based on the utility of acute spinal cord slices have further improved our understanding of neuronal properties and the composition of local circuitry in SG. Here, we present a detailed and practical guide for the preparation of spinal cord slices and show representative whole-cell recording and morphological results. This protocol permits ideal neuronal preservation and can mimic in vivo conditions to a certain extent. In summary, the ability to obtain an in vitro preparation of spinal cord slices enables stable current- and voltage-clamp recordings and could thus facilitate detailed investigations into the intrinsic membrane properties, local circuitry and neuronal structure using diverse experimental approaches.

Here, we describe the essential steps for whole-cell patch-clamp recordings made from substantia gelatinosa (SG) neurons in the in vitro spinal cord slice. This method allows the intrinsic membrane properties, synaptic transmission and morphological characterization of SG neurons to be studied.

Préparation de tranches médullaires aiguës pour l’enregistrement de Patch-clamp de la cellule entière dans les neurones de Substantia Gelatinosa

Recent whole-cell patch-clamp studies from substantia gelatinosa (SG) neurons have provided a large body of information about the spinal mechanisms ...