The authors sincerely thank the Canadian Institutes of Health Research MOP 130258 and the Star Foundation for Children&#8217;s Health Research, along with the Molly Towell Foundation, for the provision of the research facility and financial support. The authors also sincerely thank Dr. Kinkead Richard for manuscript proofreading and advice.

Cette méthode nécessitait l&#39;utilisation de sujets animaux, autorisés par le Comité d&#39;éthique animale de l&#39;Université Laval (protocole N ° 2012-170). 1. Configuration et préparation <ol><li> Solutions <ol><li> Préparer des solutions de ACSF selon les recettes 7,17 suivantes. D&#39;autres recettes avec des variations de concentration sont disponibles dans la littérature. Solutions magasin d&#39;achat d&#39;actions à 4 ° C pour jusqu&#39;à un mois. <ol><li> Sel solution: ajouter 75,39 g de NaCl (129 mM final); 2,5 g de KCl (3,35 mM final); 0,81 g de NaH2PO4 (0.58 mM final); 2,33 g de MgCl2 (1,15 mM final); 1,85 g de CaCl2 (1,26 mM). Dissoudre dans 800 ml d&#39;eau distillée puis remplir à 1 L avec de l&#39;eau distillée. </li><li> Une solution de bicarbonate: ajouter 17,65 g de NaHCO 3 (21 mM final). Dissoudre dans 900 ml d&#39;eau distillée puis remplir à 1 L avec de l&#39;eau distillée. Les variations de la concentration en bicarbonate se traduiront par des variations de pH. </li><li> Une solution de glucose: ajouter 54.06g de glucose (30 mM final). Dissoudre dans 400 ml d&#39;eau distillée puis remplir à 500 ml avec de l&#39;eau distillée. </li></ol></li><li> Préparer aCSF en diluant 100 ml d&#39;une solution de sel, 100 ml d&#39;une solution de bicarbonate et 50 ml de solution de glucose dans 1 litre d&#39;eau distillée. Conserver à 4 ° C jusqu&#39;à utilisation. </li><li> Préparer une électrode de verre par chauffage et étirage d&#39;un tube de verre jusqu&#39;à ce qu&#39;elle se casse. Poncer la pointe avant de l&#39;utiliser. L&#39;électrode peut être réutilisé plusieurs fois aussi longtemps qu&#39;il reste propre. </li></ol></li><li> Installation expérimentale Remarque: Les détails de mise en place sont présentés dans la figure 2. <ol><li> Mettez l&#39;amplificateur, le déplacement de calcul de moyenne, le système d&#39;acquisition de données, le contrôleur de température et la pompe. Vérifiez l&#39;amplification du signal (gain = 10 000), la filtration (seuil bas, 10 Hz; seuil haut, 5 kHz), et le taux d&#39;échantillonnage de la conversion analogique-numérique du signal brut (2,5 kHz). </li><li> Remplissez une bouteille avec aCSF et bulle avec carbogène (95% O 2 </sub&gt;, 5% CO 2). Induire une bubbled- et réchauffé aCSF écoulement dans la chambre d&#39;enregistrement (volume 5 ml) à 4 ml / min. Maintenir la température dans la chambre d&#39;enregistrement à 26 (± 1) ° C. pH devrait être de 7,4 dans ces conditions standard. </li><li> Maintenir 50 ml de carbogène et RT-aCSF barboter dans une seringue de 50 ml à proximité de la chambre de dissection. </li><li> Allumez l&#39;ordinateur et lancez le logiciel d&#39;enregistrement. </li></ol></li></ol> 2. Dissection <ol><li> Peser et de déterminer visuellement le sexe de l&#39;animal (les mâles ont une ano-génitale plus longue distance, tandis que les femelles ont une distance ano-génitale court; les mâles les organes génitaux sont souvent sombres tandis que les organes génitaux féminins sont de couleur rose). </li><li> Anesthésier les rongeurs nouveau-nés par l&#39;une des options suivantes: cryoanesthesia (immerger complètement l&#39;animal dans la glace pendant 4 - 5 min 18), l&#39;injection (equitensine à 4 ml / kg) 19 ou par inhalation d&#39;anesthésiques volatiles (isoflurane 20 ou éther 21). Confirm un plan adéquat de l&#39;anesthésie par l&#39;absence d&#39;un réflexe de retrait de la patte. </li><li> Placez l&#39;animal sur le banc, face ventrale vers le bas. Section de la partie rostrale de la tête coronaire avec un scalpel au niveau du bregma (visible à travers la peau et le crâne). Effectuez cette étape immédiatement après que l&#39;animal est anesthésié. </li><li> Section du corps coronaire avec un scalpel dans les membres antérieurs. </li><li> Enlever la peau, les muscles, les tissus adipeux et viscère avec des ciseaux et des pinces chirurgicales et minces. Depuis os sont mous à cet âge, être prudent pour ne pas endommager le système nerveux. Effectuez cette étape sur le banc. </li><li> Placer la préparation dans la chambre de dissection. Utilisez la aCSF stocké dans la seringue pour oxygéner la préparation. De ce point à la fin de l&#39;enregistrement, utiliser un microscope. </li><li> Sur la face dorsale de la préparation, couper le crâne et les vertèbres de la rostrale à la partie caudale long de l&#39;axe moyen avec des ciseaux et des pinces chirurgicales et minces. Ouvrez le crâne de coupeet vertèbres afin d&#39;exposer le tissu nerveux. Utilisez la aCSF strored dans la seringue pour oxygéner la préparation. </li><li> Avec la pince, retirer l&#39;arachnoïde, un tissu mince recouvrant la surface du tissu nerveux. Conserver la pie-mère et les vaisseaux sanguins contre le tissu nerveux. Utilisez la aCSF stocké dans la seringue pour oxygéner la préparation. </li><li> Placer la préparation avec la dorsale face vers le bas, et soigneusement abattre le tronc cérébral et la moelle épinière en coupant les nerfs et du tissu conjonctif avec les ciseaux tout en maintenant la préparation en place. Une approche dorsale est également possible. Garder les racines et les nerfs aussi longtemps que possible. Retirez les os pour isoler le tronc cérébral et la moelle épinière. Utilisez la aCSF stocké dans la seringue pour oxygéner la préparation. </li><li> Retirer le cervelet et le reste des structures rostrales en les sectionnant avec le scalpel. Utilisez le aCSF stockée dans l&#39;aiguille pour oxygéner la préparation. </li><li> Eventuellement fonction sur expériconception tal, retirez le pont avec le scalpel par une section coronale antérieure à l&#39;artère cérébelleuse inférieure 22. Notez que le maintien de l&#39;ensemble des pons va ralentir le rythme chez les rats et entièrement inhiber chez les souris. Les préparatifs qui ne comprennent que la partie caudale du pont affichent une activité respiratoire-like avec une fréquence stable à la racine de la C4. </li></ol> 3. Enregistrement <ol><li> Placez la préparation dans la chambre d&#39;enregistrement, ventrale face vers le haut. Fixer la préparation avec des épingles dans la partie inférieure de la moelle épinière et rostrale-plus partie du tronc cérébral. </li><li> En utilisant une seringue reliée à l&#39;électrode par son aiguille, induire une dépression dans l&#39;électrode (diamètre de la pointe de verre: de 150 à 225 um) en tirant sur le piston de la seringue, afin de combler partiellement l&#39;électrode avec aCSF. </li><li> Utilisation d&#39;un micromanipulateur, placer soigneusement l&#39;électrode à proximité de l&#39;un des quatrième racines ventrales. Autres racines ventrales peuvent aussi présenter une respiratoires comme l&#39;activité (e.g., nerf crânien XII, C1 racine ventrale). </li><li> Provoquer une dépression dans le réservoir d&#39;électrode via la seringue en tirant le piston afin d&#39;aspirer délicatement le radicelles de nerf. Puis déplacez doucement l&#39;électrode pour l&#39;appliquer contre la moelle épinière. Remarque: Idéalement, la taille des radicelles correspond à la taille de l&#39;ouverture de l&#39;électrode et crée un joint d&#39;étanchéité entre les compartiments interne et externe de l&#39;électrode ainsi. Depuis amplificateurs différentiels sont couramment utilisés pour de tels enregistrements, toute ouverture entre l&#39;intérieur de l&#39;électrode et la chambre d&#39;enregistrement réduit la qualité du signal et il est difficile d&#39;éliminer les bruits de fond. </li><li> Commencez l&#39;enregistrement. </li><li> Enregistrer le rythme produit par la préparation dans des conditions normoxiques (c.-à barboter avec du carbogène aCSF: 95% O 2 et 5% CO 2) pendant au moins 20 min afin de déterminer les paramètres de base de la préparation. </li><li> Mettez la perfusion de carbogène-barboter à aCSFaCSF stimulus (par exemple, par barbotage avec 95% de N 2 et 5% de CO2 pendant l&#39;hypoxie stimulus) pendant 15 min. Modifier la durée d&#39;exposition en fonction du protocole expérimental. Consigner la durée de l&#39;exposition sur l&#39;enregistrement et animale fiche. Utilisation tubes en acier inoxydable entre la bouteille et le ACSF chambre chaque fois que possible pour éviter la diffusion de gaz vers l&#39;extérieur du tube. </li><li> Mettez la perfusion retour à aCSF carbogène-barboter norme pendant au moins 15 minutes pour un enregistrement de récupération. Inscrire ce sur l&#39;enregistrement et animale fiche. </li><li> Terminez l&#39;enregistrement. </li></ol> 4. Analyse statistique <ol><li> A partir du signal intégré, le calcul de la fréquence que le nombre de paquets par minute (exprimée en salves / min), l&#39;amplitude de la différence entre la ligne de base et le pic de la salve (exprimée en mV), la durée de salve que la durée de du début à la fin de la salve (exprimé en s), la région d&#39;éclatement comme l&#39;aire sous la curve de la salve sur le signal intégré (exprimée en mV · s) (Figure 3). </li><li> Calculez la fréquence, l&#39;amplitude, la durée et la zone éclater éclater sous normoxique (de base), hypoxique (stimulus) et post-hypoxiques conditions (post-stimulus) de récupération comme la moyenne des les 5 dernières minutes des enregistrements pour chaque condition. Ne pas analyser la première partie de chaque condition, car il ya un délai entre la commutation de la perfusion et aCSF homogénéisation dans la chambre d&#39;enregistrement. </li><li> Exprimez puis stimulus (ie, l&#39;hypoxie) et post-stimulus (ie, post-hypoxiques) valeurs de récupération en pourcentage des valeurs de référence pour l&#39;enregistrement correspondant. Exprimer les résultats en tant que moyen ± SD </li></ol>

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M., Paton, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizations+of+eupnea,+apneusis+and+gasping+in+a+perfused+rat+preparation.">Characterizations of eupnea, apneusis and gasping in a perfused rat preparation.</a> Respir Physiol. 123, 201-213 (2000).</li></ol>

The authors have no competing financial interests or conflicts of interests to disclose.

Quantification précise de l&#39;activité respiratoire peut être difficile. En effet, la respiration est une fonction qui peut être à la fois automatique et volontaire, et que est modulée en fonction de l&#39;environnement, les besoins de l&#39;organisme, l&#39;état émotionnel et le comportement. L&#39;avantage de cette technique est l&#39;isolement des éléments neuronaux responsables de la production de la commande respiratoire. Ainsi, les enregistrements électrophysiologiques de préparations pléthysmograph...

La respiration est une activité complexe et vital commandé par le cerveau, ce qui permet dioxygène (O 2) et l&#39;absorption du dioxyde de carbone (CO 2) d&#39;élimination. L&#39;entraînement respiratoire centrale est généré par un réseau complexe situé dans le tronc cérébral dans les deux mammifères 1, 2 amphibiens, reptiles, oiseaux 3 4 et 5 poissons. Même si l&#39;étude de la respiration peut être traitée in vivo, les enquêtes mécanistes précises nécessitent un accès direct au réseau de contrôle respiratoire. À cette fin, Adrian et Buytendijk développé une préparation de poisson rouge réduite, dans laquelle des électrodes placées sur le record de surface du tronc cérébral le rythme généré associé à la ventilation des branchies 5. Cette approche a ensuite été adapté par Suzue en 1984 6 pour une utilisation chez les rongeurs nouveau-nés. L&#39;avènement de cette préparation a conduit à des avancées significatives dans la neurobiologie respiratoire. Étant donné qu&#39;il est relativement simple, la technique présentée here se prête à un large éventail d&#39;enquêtes de base de comportements moteurs rythmiques et leurs origines chez les rongeurs nouveau-nés. L&#39;objectif global de cette méthode est d&#39;enregistrer le corrélat neuronal de l&#39;activité inspiratoire, un rythme respiratoire de type appelé respiration fictive, produite par le réseau des voies respiratoires. Ce procédé peut être utilisé dans un large éventail d&#39;objectifs de recherche, visant les réponses inspiratoires aux variations ou la pharmacologie respiratoires dans les deux sauvages de type 7 et 8 animaux transgéniques. Étant donné que les expériences sont réalisées à basse température, sans afférences sensorielles, et dans des conditions où les concentrations de glucose et de O 2 dans le aCSF sont élevés, des questions ont été soulevées quant à la pertinence physiologique du signal enregistré. Bien qu&#39;il existe des différences claires entre in vivo et in vitro de conditions (par exemple., La fréquence de salves inspiratoire) le fait demeure que la présence deles éléments de base du réseau des voies respiratoires 6 permettent d&#39;étudier un rythme robuste associée à une fonction homéostatique essentiel 9,10. La raison derrière le développement et l&#39;utilisation de cette technique est de faciliter l&#39;accès direct aux éléments du tronc cérébral du réseau des voies respiratoires, qui sont difficilement accessibles in vivo, en particulier dans les nouveaux-nés. Le tronc cérébral est placé dans des conditions strictement contrôlées: le rythme enregistré est pas modulée par des apports afférents périphériques des poumons ou les organes de la carotide, ce qui permet l&#39;étude de se concentrer sur l&#39;entraînement respiratoire centrale elle-même 11. Ainsi, cet accès est utilisé pour appliquer des stimuli et enregistrer le signal de sortie. Contrairement à la pléthysmographie enregistrements, le rythme respiratoire est modulée par la totalité de ses composants dans tout le corps (par ex., Distension pulmonaire, senseurs chimiques périphériques), ce qui rend difficile l&#39;application de stimuli précis. Dans unewborn rat, le protocole consiste à enregistrer le quatrième signal de la racine ventrale sur un tronc cérébral isolé et une moelle épinière tronqué, maintenue dans le liquide céphalo-rachidien artificiel (aCSF). Le rythme généré par les préparations de la moelle du tronc cérébral-spinal est constituée de salves lentes individuels qui sont liés au signal inspiratoire 9. Préparations isolées de la moelle épinière tronc cérébral sont facilement enregistrables chez les rats de jour postnatal 0 à 4 (P0 - P4) 7. Cette approche est couramment utilisé pour évaluer la réponse hypoxique du réseau des voies respiratoires, ainsi que la réponse à l&#39;hypercapnie, acidose ou de drogues. Un protocole d&#39;hypoxie aiguë est présenté ici. Cette stimulation est obtenue par retrait de O 2 dans le aCSF; cette approche est couramment utilisé pour évaluer la tolérance et de la réactivité à des insultes hypoxiques. Le protocole induit une dépression du rythme dès la première minute jusqu&#39;à la fin de l&#39;exposition de l&#39;hypoxie (figure 1) 12. Cette dépression est inversépendant la post-hypoxique 12 de récupération. En ce qui concerne la conception expérimentale, il est important de noter que le pont, situé à la partie rostrale du tronc cérébral, a une action inhibitrice sur le générateur de rythme 8. Ainsi, les préparations de pleine tronc cérébral et la moelle épinière rostrale affichent un rythme plus faible. L&#39;inclusion de la protubérance dans l&#39;échantillon isolé de l&#39;enregistrement est déterminée en fonction du but de l&#39;expérience 13; l&#39;étude de l&#39;influence pontique sur le réseau bulbe rachidien de exigerait enregistrements avec et sans les pons de comparer les résultats 14. En outre, l&#39;un des avantages de cette technique est la possibilité d&#39;étendre la partie rostrale de la préparation d&#39;inclure mésencéphalique et / ou régions diencéphaliques 15,16, ce qui permet d&#39;évaluer l&#39;effet de ces régions sur le réseau des voies respiratoires ponto-médullaire.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Sylgard</td>
			<td>Sigma Aldrich</td>
			<td>761036-5EA</td>
			<td>Use under hood</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Bioshop</td>
			<td>SOD002</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Bioshop</td>
			<td>POC888</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Bioshop</td>
			<td>CCL444</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Bioshop</td>
			<td>MAG510</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Bioshop</td>
			<td>SOB999</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Bioshop</td>
			<td>SPM306</td>
			<td></td>
		</tr>
		<tr>
			<td>D-glucose</td>
			<td>Bioshop</td>
			<td>GLU501</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbogen</td>
			<td>Linde</td>
			<td>343-02-0006&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature Controller</td>
			<td>Warner Instruments, Hamden, CT, USA</td>
			<td>TC-324B</td>
			<td></td>
		</tr>
		<tr>
			<td>Suction electrode</td>
			<td>A-M Systems, Everett, WA, USA</td>
			<td>model 573000</td>
			<td></td>
		</tr>
		<tr>
			<td>Differential AC amplifier</td>
			<td>A-M Systems, Everett, WA, USA</td>
			<td>model 1700</td>
			<td></td>
		</tr>
		<tr>
			<td>Moving averager</td>
			<td>CWE, Ardmore, PA, USA</td>
			<td>model MA-821</td>
			<td></td>
		</tr>
		<tr>
			<td>Data acquisition system</td>
			<td>Dataq Instruments, Akron, OH, USA</td>
			<td>model DI-720</td>
			<td></td>
		</tr>
		<tr>
			<td>LabChart software</td>
			<td>ADInstruments, Colorado Springs, CO, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Prism sofware</td>
			<td>Graphpad, La Jolla, CA, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection chamber</td>
			<td>Plastic box (e.g. petri box) will do</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Recording chamber</td>
			<td>Home made</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Base</td>
			<td>Kanetec, Bensenville, IL, USA</td>
			<td>MB</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>World Precision Instrument Inc, Sarasota, FL, USA</td>
			<td>KITE-R</td>
			<td></td>
		</tr>
		<tr>
			<td>Base</td>
			<td>Kanetec, Bensenville, IL, USA</td>
			<td>MB</td>
			<td></td>
		</tr>
		<tr>
			<td>Peristaltic pump</td>
			<td>Gilson, Middleton, WI, USA</td>
			<td>MINIPULS 3</td>
			<td></td>
		</tr>
		<tr>
			<td>Faraday Cage</td>
			<td>Home made</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Computer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

electrophysiology on isolated brainstem-spinal cord preparations from newborn rodents allows neural respiratory network output recording

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to stimuli remain to be fully understood. To overcome the difficulty of accessing the brainstem in the whole animal, isolation of the brainstem and part of the spinal cord is performed. This preparation is maintained in artificial cerebro-spinal fluid where gases, concentrations, and temperature are controlled and monitored. The output signal from the respiratory network is recorded by a suction electrode placed on the fourth ventral root. In this manner, stimuli can be directly applied onto the brainstem, and the effect can be recorded directly. The signal recorded is linked to the inspiratory signal sent to the diaphragm via the phrenic nerve, and can be described as bursts (around 8 bursts per minute). Analysis of these bursts (frequency, amplitude, length, and area under the curve) allows precise characterization of the stimulus effect on the respiratory network. The main limitation of this method is the viability of the preparation beyond the early post-natal stages. Thus, this method greatly focuses on the study of the whole network without the peripheral inputs in the newborn rat.

Comme mentionné dans l&#39;introduction, l&#39;un des avantages les plus importants de cette technique est l&#39;accès direct au tronc cérébral pour appliquer divers stimuli. A titre d&#39;exemple, l&#39;hypoxie a été appliqué ici. La Figure 1. AB affiche un enregistrement de protocole complète, avec les deux conditions normoxiques et hypoxiques. Figure 1.CE affiche le rythme enregistrés dans des conditions normoxiques (par exemple, aCSF barboter avec 95% O 2</sub...

Watch this Scientific Journal Video about Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording at JoVE.com

Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording

The central respiratory drive is located in the brainstem. Spontaneous respiratory motor output from an isolated brainstem-spinal cord is recorded by placing an electrode on the fourth ventral root. This experimental approach is valuable for pharmacological investigations or the assessment of respiratory challenges and genetic manipulations on rhythmic motor behavior.

Électrophysiologie sur isolés Cord Préparatifs du tronc cérébral-spinal de la naissance Rongeurs Permet Neural respiratoire Réseau d&#39;enregistrement de sortie

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to ...

electrophysiology-on-isolated-brainstem-spinal-cord-preparations-from

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Neuroscience

8.4K vues.  Laval University. The central respiratory drive is located in the brainstem. Spontaneous respiratory motor output from an isolated brainstem-spinal cord is recorded by placing an electrode on the fourth ventral root. This experimental approach is valuable for pharmacological investigations or the assessment of respiratory challenges and genetic manipulations on rhythmic motor behavior.

Vidéo: Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Dissection et la culture de neurones de la moelle épinière commissurales embryonnaires

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

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Neurosciences

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

La moelle épinière électrophysiologie II: fabrication des électrodes extracellulaires d&#39;aspiration

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

Examining Local Network Processing using Multi-contact Laminar Electrode Recording

Cortical layers are ubiquitous structures throughout neocortex1-4 that consist of highly recurrent local networks. In recent years, significant progress has been made in our understanding of differences in response properties of neurons in different cortical layers5-8, yet there is still a great deal left to learn about whether and how neuronal populations encode information in a laminar-specific manner.
Existing multi-electrode array techniques, although informative for measuring responses across many millimeters of cortical space along the cortical surface, are unsuitable to approach the issue of laminar cortical circuits. Here, we present our method for setting up and recording individual neurons and local field potentials (LFPs) across cortical layers of primary visual cortex (V1) utilizing multi-contact laminar electrodes (Figure 1; Plextrode U-Probe, Plexon Inc).
The methods included are recording device construction, identification of cortical layers, and identification of receptive fields of individual neurons. To identify cortical layers, we measure the evoked response potentials (ERPs) of the LFP time-series using full-field flashed stimuli. We then perform current-source density (CSD) analysis to identify the polarity inversion accompanied by the sink-source configuration at the base of layer 4 (the sink is inside layer 4, subsequently referred to as granular layer9-12). Current-source density is useful because it provides an index of the location, direction, and density of transmembrane current flow, allowing us to accurately position electrodes to record from all layers in a single penetration6, 11, 12.

A fundamental issue in our understanding of cortical circuitry is how networks in different cortical layers encode sensory information. Here we describe electrophysiological techniques utilizing multi-contact laminar electrodes to record single-units and local field potentials and present analyses to identify cortical layers.

L&#39;examen de transformation réseau local en utilisant l&#39;enregistrement multi-contact d&#39;électrode laminaire

Cortical layers are ubiquitous structures throughout neocortex1-4 that consist of highly recurrent local networks. In recent years, significant progress ...

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.

Marquage fluorescent rétrograde Permet ciblée extracellulaire seule unité d&#39;enregistrement de neurones identifiés<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 ...

Neural Circuit Recording from an Intact Cockroach Nervous System

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological effects of insecticides. This article describes the scope of cockroach sensory modalities that can be used to assay how an insect nervous system responds to environmental perturbations. Emphasis here is on the escape behavior mediated by cerci to giant fiber transmission in Periplaneta americana. This in situ preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of neuronal activity. Peptides or other chemical reagents can then be applied directly to the nervous system in solution with the physiological saline. Insecticides could also be administered prior to dissection and the escape circuit can serve as a proxy for the excitable state of the central nervous system. In this context the assays described herein would also be useful to researchers interested in limb regeneration and the evolution of nervous system development for which P. americana is an established model organism.

This article describes the cockroach ventral nerve cord dissection and extracellular recordings from the cercal nerve and connectives. Evoked responses are generated by electrical stimulation of the cercal nerve or direct mechanical stimulation of the cerci.

Des circuits neuronaux enregistrement à partir d&#39;un cancrelat système nerveux Intact

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Augmentation de la réponse du réseau à la stimulation des cultures cellulaires neuronales sur les tableaux de microélectrodes

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...

Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a signal driving the rhythmic contraction of inspiratory muscles. Rhythmic neural activity generated in the pBC and carried to other neuronal pools to drive the musculature of breathing may be studied using various approaches, including en bloc nerve recordings and transverse slice recordings. However, previously published methods have not extensively described the brainstem-spinal cord dissection process in a transparent and reproducible manner for future studies. Here, we present a comprehensive overview of a method used to reproducibly cut rhythmically-active brainstem slices containing the necessary and sufficient neuronal circuitry for generating and transmitting inspiratory drive. This work builds upon previous brainstem-spinal cord electrophysiology protocols to enhance the likelihood of reliably obtaining viable and rhythmically-active slices for recording neuronal output from the pBC, hypoglossal premotor neurons (XII pMN), and hypoglossal motor neurons (XII MN). The work presented expands upon previous published methods by providing detailed, step-by-step illustrations of the dissection, from whole rat pup, to in vitro slice containing the XII rootlets.

This protocol both visually communicates the brainstem-spinal cord preparation and clarifies the preparation of brainstem transverse slices in a comprehensive step-by-step manner. It was designed to increase reproducibility and enhance the likelihood of obtaining viable, long lasting, rhythmically-active slices for recording neural output from the respiratory regions of the brainstem.

Préparation de rythmiquement actifs In Vitro néonatale rongeurs tronc cérébral-moelle épinière et fine tranche

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Enregistrement optique à grand champ à un seul photon dans des tranches de cerveau à l'aide de teinture sensible à la tension

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an increasingly detailed view of the circuits underpinning spinal pain processing. However, the effects of these connections for broader network activity in the DH remain less well understood because most studies focus on the activity of single neurons and small microcircuits. Alternatively, the use of microelectrode arrays (MEAs), which can monitor electrical activity across many cells, provides high spatial and temporal resolution of neural activity. Here, the use of MEAs with mouse spinal cord slices to study DH activity induced by chemically stimulating DH circuits with 4-aminopyridine (4-AP) is described. The resulting rhythmic activity is restricted to the superficial DH, stable over time, blocked by tetrodotoxin, and can be investigated in different slice orientations. Together, this preparation provides a platform to investigate DH circuit activity in tissue from na&#239;ve animals, animal models of chronic pain, and mice with genetically altered nociceptive function. Furthermore, MEA recordings in 4-AP-stimulated spinal cord slices can be used as a rapid screening tool to assess the capacity of novel antinociceptive compounds to disrupt activity in the spinal cord DH.

The combined use of microelectrode array technology and 4-aminopyridine-induced chemical stimulation for investigating network-level nociceptive activity in the spinal cord dorsal horn is outlined.

Enregistrement de l’activité du réseau dans les circuits nociceptifs rachidiens à l’aide de réseaux de microélectrodes

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an ...

Comprendre les réponses des motoneurones à la stimulation de la moelle épinière

The Ex vivo Preparation of Spinal Cord Slice for the Whole-Cell Patch-Clamp Recording in Motor Neurons During Spinal Cord Stimulation

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to execute sensorimotor behaviors, directly studying the electrical responses of motor neurons with SCS can help us understand the underlying logic of spinal motor modulation. To simultaneously record diverse stimulus characteristics and cellular responses, a patch-clamp is a good method to study the electrophysiological characteristics at a single-cell scale. However, there are still some complex difficulties in achieving this goal, including maintaining cell viability, quickly separating the spinal cord from the bony structure, and using the SCS to successfully induce action potentials. Here, we present a detailed protocol using patch-clamp to study the electrical responses of motor neurons to SCS with high spatiotemporal resolution, which can help researcher improve their skills in separating the spinal cord and maintaining the cell viability at the same time to smoothly study the electrical mechanism of SCS on motor neuron and avoid unnecessary trial and mistake.

Pleins feux sur l’auteur : Faire progresser la stimulation de la moelle épinière - Explorer les réponses cellulaires des motoneurones grâce à l’électrophysiologie par patch-clamp 

This protocol describes a method using a patch-clamp to study the electrical responses of motor neurons to spinal cord stimulation (SCS) with high spatiotemporal resolution, which can help researchers improve their skills in separating the spinal cord and maintaining cell viability simultaneously.

Préparation ex vivo d’une tranche de moelle épinière pour l’enregistrement d’un patch-clamp de cellules entières dans les motoneurones lors d’une stimulation de la moelle épinière

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to ...