Les auteurs remercient Giorgio Carlini pour le soutien technique dans l&#39;élaboration de la structure de confinement et Dott. Ornella LoBrutto pour la révision approfondie du manuscrit. La recherche menant à ces résultats a reçu un financement de 7 ème programme-cadre de l&#39;Union européenne (TIC-FET FP7 / 2007-2013, FET Jeunes Explorateurs régime) sous convention de subvention 284772 CERVEAU BOW (www.brainbowproject.eu).

Le protocole expérimental a été approuvé par la législation européenne de protection des animaux (2010/63 / UE), par le ministère italien de la Santé, conformément à la DL 116/1992 et par les lignes directrices de l&#39;Université de Gênes (Prot. N. 13130, mai 2011). Tous les efforts ont été faits pour réduire le nombre d&#39;animaux pour le projet et pour minimiser leur souffrance. 1. Préparation des matériaux et supports <ol><li> Construire le moule pour construire les PDMS (Poly-diméthyl-siloxane) contrainte au moyen d&#39;un CNC (commande numérique par ordinateur) fraiseuse. Un tel moule est réalisé en polycarbonate avec le cylindre central en polytétrafluoroéthylène (PTFE). Ce matériau permet une extraction plus facile du masque une fois que le PDMS a durci. REMARQUE: La conception du moule a été réalisé par Computer-Aided-Design (CAD), puis livré à la fraiseuse CNC. </li><li> Préparer l&#39;élastomère PDMS. PDMS est un polymère organique. Il est composé de deux éléments, Un agent de durcissement et un polymère. Mélangez par un rapport en volume de 1:10, une partie d&#39;agent de durcissement et neuf parties de polymère. Mettez les deux composants dans une boîte de Pétri, secouez et insérez le mélange dans la chambre à vide pendant 10 min pour éliminer les bulles d&#39;air. 3 g de PDMS sont suffisantes pour une contrainte. </li><li> Construire la contrainte PDMS. Insérez le matériau PDMS dans le mouleur et le mettre dans le four pendant 30 min à 120 ° C. La contrainte PDMS a la forme d&#39;un cylindre idéal avec un diamètre externe et interne de 22,0 mm et 3,0, respectivement, et une hauteur de 650 pm. </li><li> Le jour avant le placage, coupler le masque à la zone active des réseaux de micro-électrodes (AME) à l&#39;aide d&#39;une pince à épiler sous un stéréomicroscope et stériliser le masque PDMS dans le four à 120 ° C. </li><li> Stériliser les microbilles de verre (diamètre nominal de 40 ± 2 um; diamètre moyen de 42,3 ± certifié 1,1 pm) dans 70% d&#39;éthanol pendant 2 heures dans un flacon conique et tourner evEry 30 min pour exposer tous les microbilles. </li><li> Retirer la solution d&#39;éthanol à partir de la fiole et les microbilles rincer deux fois avec de l&#39;eau stérilisée. </li><li> Conditionner la partie centrale de la zone délimitée par la MEA masque PDMS (diamètre égal à 3,0 mm) avec 24 ul d&#39;une solution mixte de Poly-D-Lysine et laminine à 0,05 ug / ml. </li><li> Enduire les microbilles avec des protéines d&#39;adhérence, de la laminine et de poly-D-lysine à 0,05 ug / ml et de les laisser pendant une nuit à l&#39;étuve à 37 ° C, pour obtenir un réseau neuronal stable et de longue durée. </li><li> Retirer les facteurs d&#39;adhésion à la fois à partir de la surface du verre de la MEA et de microbilles de verre. Aspirer environ 95% de la solution de revêtement avec une pipette et appliquer une petite 24μl- en volume d&#39;eau stérile à la surface du MEA. Aspirer à nouveau plus de 95% de l&#39;eau et laissez la MEA à sec sous le capot laminaire pendant 1 h avant l&#39;ensemencement des cellules. NOTE: Dans le cas de microbilles place, aspirer le revêtementsolution et faire un premier lavage avec de l&#39;eau stérilisée et un second avec le milieu de base et son supplément comme B27, afin de conditionner la surface de verre où les neurones seront étalées. Ne laissez pas les microbilles sec. Les laisser en suspension dans le milieu de culture à l&#39;intérieur du flacon. </li><li> Distribuer, au moyen d&#39;une pipette -200 μl-, la suspension de microbilles traités (environ 32 000) sur une plaque à puits multiples avec insert de membrane (diamètre des pores 0,4 pm), où ils auto-assemblage formant une couche uniforme. </li><li> Remplir chaque puits de la membrane avec 0,5 ml de milieu et le mettre dans l&#39;incubateur (T = 37,0 ° C, CO 2 = 5%) jusqu&#39;à ce que les neurones sont prêts à être plaqué (3.2). </li></ol> 2. Dissection d&#39;embryons et de la dissociation des tissus <ol><li> Maison rats femelles adultes (200-250 g) à une température constante (22 ± 1 ° C) et l&#39;humidité relative (50%) selon un calendrier clair-obscur régulière (la lumière sur les sept heures-19 heures) dans leanimalerie. Veiller à ce que la nourriture et l&#39;eau sont disponibles gratuitement. </li><li> Anesthésier un rat femelle enceinte après 18 jours de développement (E18) en utilisant 3% d&#39;isoflurane. Ensuite, sacrifier l&#39;animal par dislocation cervicale. </li><li> Retirer hippocampes de l&#39;embryon de rat et les placer dans une solution saline équilibrée de la glace froide de Hank sans Ca 2+ et Mg 2+. Au jour 18 de l&#39;hippocampe de développement et de tissus de cortex sont très doux et ne nécessitent pas d&#39;être coupé en petits morceaux. De plus amples détails peuvent être trouvés 18,19. </li><li> Dissocier le tissu à 0,125% de solution de trypsine / 0,05% de Hank contenant de 18 à 20 pour la DNAse min à 37 ° C. </li><li> Retirer la solution de surnageant avec une pipette Pasteur et arrêter la digestion enzymatique par addition de milieu avec 10% de sérum bovin fœtal (FBS) pendant 5 min. </li><li> Retirer le moyen de FBS et laver une fois avec du milieu avec son complément, 1% de L-glutamine, la gentamicine 10 ug / ml. Retirer à nouveau le milieu avec un pip Pasteurette et remplir à nouveau avec une petite quantité (500 pi) de milieu de croissance avec son complément, 1% de L-glutamine, la gentamicine 10 ug / ml. </li><li> Dissocier le culot tissulaire mécaniquement avec une pipette Pasteur étroite jusqu&#39;à une suspension laiteuse de cellules est apparent. Il est pas nécessaire de centrifuger la suspension cellulaire. </li><li> Diluer le faible volume de suspension de cellules avec le milieu de croissance pour obtenir un volume final de 2,0 ml. Compter le concentration cellulaire obtenue avec une chambre de hémocytomètre. Diluer cette concentration à 1: 5 pour obtenir la concentration souhaitée de cellules de 600-700 cellules / ul. </li></ol> 3. Cellule Placage <ol><li> Cellules de la plaque à une densité d&#39;environ 2000 cellules / mm 2 sur la zone active de la MEA, définis par la contrainte de PDMS pour créer un réseau neuronal 2D. NOTE: Chaque hippocampe contient environ 5 x 10 5 cellules, (1 x 10 6 pour un seul embryon). Disséquer 6 embryons pour obtenir un montant total de 6 x 106 cellules dans 2 ml, et une concentration d&#39;environ 3.000 cellules / ul. Diluer cette concentration à 1: 5 pour obtenir la concentration souhaitée de la cellule et la plaque environ 600-700 cellules / ul. Si la zone de MEA délimitée par la contrainte est d&#39;environ 7,065 PDMS mm 2, et le nombre total de cellules étalées à 600 cellules / ul ul x 24 = 14.400 cellules, la densité finale de 2.038 cellules / mm 2. </li><li> Placez les dispositifs de MEA dans l&#39;incubateur humidifié avec atmosphère de CO 2 (5%) à 37 ° C. </li><li> Distribuer 160 pl de la suspension à une concentration cellulaire de 600-700 cellules / ul (environ 100 000 cellules) sur la surface de la monocouche de microbilles placée à l&#39;intérieur des plaques multipuits pour terminer l&#39;étape préliminaire à la construction de culture en trois dimensions. Mettre les plaques à puits multiples dans l&#39;incubateur humidifié avec atmosphère de CO 2 (5%) à 37 ° C. </li></ol> 4. Construction 3D Neuronal réseau <ol><li> 6-8 heures après le placage, transférer la suspension (microbilles avec les neurones) des plaques à puits multiples très soigneusement l&#39;intérieur de la zone délimitée par la contrainte PDMS en utilisant une pipette fixé pour un volume d&#39;environ 30-40 ul. Après chaque transfert, attendez pendant environ une demi-minute, pour permettre aux microbilles se auto-assembler dans une structure hexagonale compacte. </li><li> Une fois toutes les couches sont déposées et spontanément assemblés, remplir avec une forte baisse d&#39;environ 300 pi de milieu du haut de la zone délimitée par la contrainte PDMS. </li><li> Mettre la structure 3D couplée à la MEA dans l&#39;incubateur (T = 37,0 ° C, CO 2 = 5%) pendant 48 heures avant d&#39;ajouter un volume final (environ 1 ml) de milieu de croissance avec la culture de son complément. NOTE: Considérons le nombre total de perles sur les plaques à puits multiples avec insert de membrane (30 000) et le nombre de perles sur une seule couche sur le dispositif de MEA (6000). La structure 3D résultante se compose de 5 couches de microbeads et des cellules. Tenant compte du fait que le réseau neuronal 3D est pas géométriquement parfaite, la structure 3D qui en résulte pourrait être composé de 5-8 couches. </li><li> Le jour après l&#39;étalement, ajouter soigneusement le volume final du milieu (environ 900 pi) à l&#39;intérieur de l&#39;anneau de MEA. Maintenir les cultures 3D dans une atmosphère humidifiée de CO2 (5%) à 37 ° C pendant 4-5 semaines. Remplacer la moitié du milieu une fois par semaine. </li></ol> 5. microscopie confocale Acquisition <ol><li> Fixer les échantillons biologiques à l&#39;étape du microscope dans un support. Réglez le laser (Argon, de 496 à 555 nm), la vitesse de balayage (400 Hz) à laquelle d&#39;acquérir l&#39;image, le gain et l&#39;offset (-0,3%) de PMT pour chaque image pour capturer le filtre bande passante d&#39;émission correcte et dans l&#39;ordre pour éviter la saturation et à augmenter le rapport signal à bruit. La section Résultats rapporte les valeurs utilisées pour acquérir les images de la figure 4. </li><li> Pour l&#39;acquisition de la séquence z -stack, sapinst sélectionner, la valeur de z position de la partie supérieure et la couche inférieure de l&#39;échantillon, puis sélectionnez la taille de l&#39;étape pour faire l&#39;acquisition. </li></ol>

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The authors declare that they have no competing financial interests.

Dans ce travail, un roman expérimental dans la plateforme vitro constitué de 3D ​​conçu cultures de neurones couplés à AME pour électrophysiologie de réseau a été présenté. L&#39;utilisation des microbilles comme échafaudage pour permettre au long de l&#39;excroissance neuritique axe des z a été adapté pour être intégré avec le MEA plane. De cette façon, les résultats micro-système obtenus dans un modèle in vitro 3D valide et fiable pour étudier la dynamique électro...

In vitro en deux dimensions (2D), les réseaux de neurones couplés à micro-électrodes (AME) Arrays arethe modèle expérimental étalon-or adoptée pour étudier l&#39;interaction entre les dynamiques neuronales et de la connectivité sous-jacente. Lors du développement, les neurones de recréer des réseaux complexes qui affichent ainsi l&#39;activité definedspatio-temporalpatternsof 1,2 (ie, des éclats, des éclats de réseau, l&#39;activité de dopage aléatoire). AME enregistrent l&#39;activité électrophysiologique de nombreux sites (de quelques dizaines à des milliers de microélectrodes), permettant une étude détaillée de la dynamique exprimées au niveau du réseau. En outre, l&#39;utilisation de cultures dissociées rend possibile de concevoir-réseaux d&#39;ingénierie. Il est plus facile de comprendre de cette manière les relations fonctionnelles entre l&#39;activité électrophysiologique enregistrées et les paramètres de l&#39;organisation de réseau comme la densité cellulaire 3, degré de modularité 4,5, la présence de la pop neuronale hétérogèneglement 6, etc. Cependant, toutes les études in vitro sur des cellules en culture dissociés sont basés sur des réseaux neuronaux 2D. Cette approche conduit à des simplifications excessives par rapport à la (intrinsèquement 3 dimensions 3D) système in vivo: (i) dans un modèle 2D, Somata et de croissance cônes sont aplaties et l&#39;excroissance des axones-dendrites peut pas se propager dans toutes les directions 7. (ii) 2D in vitro réseaux exposition stéréotypé dynamique électrophysiologiques dominés par l&#39;éclatement activité impliquant la plupart des neurones du réseau 8. Récemment, différentes solutions ont été développées pour permettre la construction d&#39;dissocié in vitro 3D réseaux neuronaux. L&#39;idée commune consiste à créer un échafaudage où les neurones peuvent se développer dans un mode 3D. Un tel échafaudage peut être réalisé avec des gels de polymères et de matrices poreuses solides 9-13. En exploitant les propriétés mécaniques des polymères, il est possible d&#39;incorporer des cellules à l&#39;intérieur thesstructures de e en définissant un bloc uniforme des cultures 3D de neurosphères 11. La principale caractéristique de cette approche est la propriété mécanique rigide des neurosphères 9,12. Cependant, ces matériaux ont une porosité limitée, et ils ne garantissent pas la migration des cellules à l&#39;intérieur de la matrice. Pour pallier cet inconvénient, une solution possible consiste à découper la matrice en modules de «base». Malheureusement, la taille et la forme de la diversité des particules pourrait entraver l&#39;emballage dans des structures en couches régulières. Dans 7, Cullen et ses collègues ont conçu une construction neuronale 3D constitué de neurones et / ou des astrocytes dans un extracellulaire échafaudage à base de matrice bioactive. Un tel tissu neural conçu permis dans les enquêtes in vitro pour étudier et manipuler les réponses neurobiologiques dans des micro-environnements 3D. Ce modèle est composée de neurones et de cellules gliales distribuées dans toute la matrice extracellulaire (ECM) et / ou des échafaudages d&#39;hydrogel (500 à 600 pm d&#39;épaisseur). Dans ce condition, une viabilité cellulaire optimale (supérieur à 90%) a été trouvée en étalant les cellules à une densité finale d&#39;environ 3750 - 5000 cellules / mm 3. Il est à noter qu&#39;une telle valeur de densité est beaucoup plus faible que celle de la condition in vivo, où la densité de cellules du cortex cérébral de souris est d&#39;environ 90 000 cellules / mm 3 14. Pour surmonter cette limitation Pautot et ses collaborateurs 15 réalisé un système 3D in vitro où la densité cellulaire et la connectivité réseau sont contrôlés à ressembler à des conditions in vivo, tout en permettant l&#39;imagerie en temps réel du réseau. En pratique, ce procédé est basé sur le concept que dissociées sont des neurones en culture capable de croître sur des microbilles de silice. Ces perles offrent une surface assez grande pour les corps cellulaires neuronaux à respecter et pour leurs arborisations à grandir, mûrir, étendre, et de définir des contacts synaptiques avec d&#39;autres neurones croissance. Cette méthode exploite les propriétés d&#39;assemblage spontané de perles mono-dispersées à form 3D couches réseaux hexagonaux contenant des sous-ensembles distincts de neurones sur des couches différentes avec la connectivité entre les neurones contraint sur différentes perles. La densité cellulaire obtenue avec cette méthode était d&#39;environ 75 000 cellules / mm 3. Récemment, nous avons adapté la méthode de Pautot aux AME 16: les résultats obtenus montrent que l&#39;activité électrophysiologique 3D présente un répertoire plus large d&#39;activités que celui exprimé par des réseaux 2D. Cultures matures 3D présentent une dynamique améliorée dans laquelle les deux rafale de réseau et aléatoire l&#39;activité de pointe coexistent. De même, Tang-Schomer et ses collaborateurs ont réalisé 17 un échafaudage poreux à base de protéine de soie qui maintient une culture corticale primaire in vitro pour plusieurs mois, et enregistré l&#39;activité électrophysiologique à l&#39;aide d&#39;une électrode en tungstène. Dans ce travail, les procédures expérimentales de construire des réseaux neuronaux 3D couplés à AME sera décrite.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Laminin</td>
			<td>Sigma-Aldrich</td>
			<td>L2020</td>
			<td>0.05 &#181;g/ml</td>
		</tr>
		<tr>
			<td>Poly-D-lysine</td>
			<td>Sigma-Aldrich</td>
			<td>P6407</td>
			<td>0.05 &#181;g/ml</td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Gibco</td>
			<td>25050-014</td>
			<td align="left">0.125% diluted 1:2 in HBSS w\o CA++, MG++</td>
		</tr>
		<tr>
			<td>DNAase</td>
			<td>Sigma-Aldrich</td>
			<td>D5025</td>
			<td>0.05% diluted&#160; in Hanks solution</td>
		</tr>
		<tr>
			<td>Neurobasal</td>
			<td>Gibco Invitrogen</td>
			<td>21103049</td>
			<td>culture medium</td>
		</tr>
		<tr>
			<td>B27</td>
			<td>Gibco Invitrogen</td>
			<td>17504044</td>
			<td>2% medium supplent</td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>Sigma-Aldrich</td>
			<td>F-2442</td>
			<td>10%</td>
		</tr>
		<tr>
			<td>Glutamax-I&#160;</td>
			<td>Gibco</td>
			<td>35050038</td>
			<td>0.5 mM</td>
		</tr>
		<tr>
			<td>gentamicin</td>
			<td>Sigma-Aldrich</td>
			<td>G.1272</td>
			<td>5mg/liter</td>
		</tr>
		<tr>
			<td>Poly-Dimethyl-Siloxane (PDMS)</td>
			<td>Corning Sigma</td>
			<td>481939</td>
			<td>curing agent and the polymer</td>
		</tr>
		<tr>
			<td>Micro-Electrode Arrays</td>
			<td>Multi Channel Systems (MCS)</td>
			<td>60MEA200/30-Ti-pr</td>
			<td>MEA with: Electrode grid 8x8; Electrode spacing and diameter 200 and 30 &#181;m, respectively; plastic ring without thread</td>
		</tr>
		<tr>
			<td>Microbead</td>
			<td>Distrilab-Duke Scientific</td>
			<td>9040</td>
			<td>&#160;1gr Glass Part.Size Stds 40 &#181;m</td>
		</tr>
		<tr>
			<td>Transwell</td>
			<td>Costar Sigma</td>
			<td>CLS 3413</td>
			<td>multiwell plates with membrane insert (6.5 mm diameter porous 0.4 &#181;m)</td>
		</tr>
		<tr>
			<td>HBSS w\o Ca++ ,Mg++&#160;</td>
			<td>Gibco Invitrogen</td>
			<td>14175-052</td>
			<td></td>
		</tr>
		<tr>
			<td>Hanks Buffer Solution&#160;</td>
			<td>Sigma&#160;</td>
			<td>H8264</td>
			<td></td>
		</tr>
		<tr>
			<td>Teflon</td>
			<td>Sigma</td>
			<td>430935-5G</td>
			<td>polytetrafluoroethylene</td>
		</tr>
		<tr>
			<td>Rat</td>
			<td>Sprague Dawley</td>
			<td>Wistar Rat</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal Microscopy Upright</td>
			<td>Leica</td>
			<td>TCS SP5 AOBS</td>
			<td></td>
		</tr>
		<tr>
			<td>20.0x0.50 WATER objective</td>
			<td>Leica</td>
			<td>Leica HCX APO L U-V-I&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>40.0x0.80 WATER objective</td>
			<td>Leica</td>
			<td>Leica HCX APO L U-V-I&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>25.0x0.95 WATER objective</td>
			<td>Leica</td>
			<td>HCX IRAPO L</td>
			<td></td>
		</tr>
	</tbody>
</table>

interfacing 3d engineered neuronal cultures to micro-electrode arrays: an innovative in vitro experimental model

Currently, large-scale networks derived from dissociated neurons growing and developing in vitro on extracellular micro-transducer devices are the gold-standard experimental model to study basic neurophysiological mechanisms involved in the formation and maintenance of neuronal cell assemblies. However, in vitro studies have been limited to the recording of the electrophysiological activity generated by bi-dimensional (2D) neural networks. Nonetheless, given the intricate relationship between structure and dynamics, a significant improvement is necessary to investigate the formation and the developing dynamics of three-dimensional (3D) networks. In this work, a novel experimental platform in which 3D hippocampal or cortical networks are coupled to planar Micro-Electrode Arrays (MEAs) is presented. 3D networks are realized by seeding neurons in a scaffold constituted of glass microbeads (30-40 &#181;m in diameter) on which neurons are able to grow and form complex interconnected 3D assemblies. In this way, it is possible to design engineered 3D networks made up of 5-8 layers with an expected final cell density. The increasing complexity in the morphological organization of the 3D assembly induces an enhancement of the electrophysiological patterns displayed by this type of networks. Compared with the standard 2D networks, where highly stereotyped bursting activity emerges, the 3D structure alters the bursting activity in terms of duration and frequency, as well as it allows observation of more random spiking activity. In this sense, the developed 3D model more closely resembles in vivo neural networks.

. Dans ce mode opératoire, un rôle important est joué par les microbilles qui définissent l&#39;échafaudage mécanique pour la croissance du réseau neuronal 3D figures affichage 1A et B la disposition des microbilles (diamètre nominal de 40 ± 2 um; diamètre moyen certifiée de 42,3 ± 1,1 pm) dans le plan. La particularité de ces structures est que microbilles auto assembler spontanément définissant une géométrie hexagonale compacte (figure 1B et C). L&#39...

Watch this Scientific Journal Video about Interfacing 3D Engineered Neuronal Cultures to Micro-Electrode Arrays: An Innovative In Vitro Experimental Model at JoVE.com

Interfacing 3D Engineered Neuronal Cultures to Micro-Electrode Arrays: An Innovative In Vitro Experimental Model

In this work, a novel experimental model in which 3D neuronal cultures are coupled to planar Micro-Electrode Arrays (MEAs) is presented. 3D networks are built by seeding neurons in a scaffold made up of glass microbeads on which neurons grow and form interconnected 3D structures.

Cultures interface 3D Engineered neuronales à des micro-électrodes tableaux: Une innovante<em&gt; In Vitro</em&gt; Modèle expérimental

Currently, large-scale networks derived from dissociated neurons growing and developing in vitro on extracellular micro-transducer devices are the ...

interfacing-3d-engineered-neuronal-cultures-to-micro-electrode-arrays

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JoVE Journal

Neuroscience

Interfacing 3D Engineered Neuronal Cultures to Micro-Electrode Arrays: An Innovative In Vitro Experimental Model

9.9K vues.  University of Genova. In this work, a novel experimental model in which 3D neuronal cultures are coupled to planar Micro-Electrode Arrays (MEAs) is presented. 3D networks are built by seeding neurons in a scaffold made up of glass microbeads on which neurons grow and form interconnected 3D structures.

Vidéo: Interfacing 3D Engineered Neuronal Cultures to Micro-Electrode Arrays: An Innovative In Vitro Experimental Model

Multi-electrode Array Recordings of Neuronal Avalanches in Organotypic Cultures

The cortex is spontaneously active, even in the absence of any particular input or motor output. During development, this activity is important for the migration and differentiation of cortex cell types and the formation of neuronal connections1. In the mature animal, ongoing activity reflects the past and the present state of an animal into which sensory stimuli are seamlessly integrated to compute future actions. Thus, a clear understanding of the organization of ongoing i.e. spontaneous activity is a prerequisite to understand cortex function. 
Numerous recording techniques revealed that ongoing activity in cortex is comprised of many neurons whose individual activities transiently sum to larger events that can be detected in the local field potential (LFP) with extracellular microelectrodes, or in the electroencephalogram (EEG), the magnetoencephalogram (MEG), and the BOLD signal from functional magnetic resonance imaging (fMRI). The LFP is currently the method of choice when studying neuronal population activity with high temporal and spatial resolution at the mesoscopic scale (several thousands of neurons). At the extracellular microelectrode, locally synchronized activities of spatially neighbored neurons result in rapid deflections in the LFP up to several hundreds of microvolts. When using an array of microelectrodes, the organizations of such deflections can be conveniently monitored in space and time. 
Neuronal avalanches describe the scale-invariant spatiotemporal organization of ongoing neuronal activity in the brain2,3. They are specific to the superficial layers of cortex as established in vitro4,5, in vivo in the anesthetized rat 6, and in the awake monkey7. Importantly, both theoretical and empirical studies2,8-10 suggest that neuronal avalanches indicate an exquisitely balanced critical state dynamics of cortex that optimizes information transfer and information processing.
In order to study the mechanisms of neuronal avalanche development, maintenance, and regulation, in vitro preparations are highly beneficial, as they allow for stable recordings of avalanche activity under precisely controlled conditions. The current protocol describes how to study neuronal avalanches in vitro by taking advantage of superficial layer development in organotypic cortex cultures, i.e. slice cultures, grown on planar, integrated microelectrode arrays (MEA; see also 11-14).

A robust way to study neuronal avalanches, i.e. scale-invariant spatio-temporal activity bursts, indicative of critical state dynamics in cortex. Avalanches emerge spontaneously in developing superficial layers of cultured cortex which allows for long-term measurements of the activity with planar integrated multi-electrode arrays (MEA) under precisely controlled conditions.

Enregistrements tableau multi-électrodes des Avalanches neuronale dans des cultures organotypiques

The cortex is spontaneously active, even in the absence of any particular input or motor output.  During development, this activity is important for the ...

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Neurosciences

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

The mouse is an excellent model organism to study mammalian brain development due to the abundance of molecular and genetic data. However, the developing mouse brain is not suitable for easy manipulation and imaging in vivo since the mouse embryo is inaccessible and opaque. Organotypic slice cultures of embryonic brains are therefore widely used to study murine brain development in vitro. Ex-vivo manipulation or the use of transgenic mice allows the modification of gene expression so that subpopulations of neuronal or glial cells can be labeled with fluorescent proteins. The behavior of labeled cells can then be observed using time-lapse imaging. Time-lapse imaging has been particularly successful for studying cell behaviors that underlie the development of the cerebral cortex at late embryonic stages 1-2. Embryonic organotypic slice culture systems in brain regions outside of the forebrain are less well established. Therefore, the wealth of time-lapse imaging data describing neuronal cell migration is restricted to the forebrain 3,4. It is still not known, whether the principles discovered for the dorsal brain hold true for ventral brain areas. In the ventral brain, neurons are organized in neuronal clusters rather than layers and they often have to undergo complicated migratory trajectories to reach their final position. The ventral midbrain is not only a good model system for ventral brain development, but also contains neuronal populations such as dopaminergic neurons that are relevant in disease processes. While the function and degeneration of dopaminergic neurons has been investigated in great detail in the adult and ageing brain, little is known about the behavior of these neurons during their differentiation and migration phase 5. We describe here the generation of slice cultures from the embryonic day (E) 12.5 mouse ventral midbrain. These slice cultures are potentially suitable for monitoring dopaminergic neuron development over several days in vitro. We highlight the critical steps in generating brain slices at these early stages of embryonic development and discuss the conditions necessary for maintaining normal development of dopaminergic neurons in vitro. We also present results from time lapse imaging experiments. In these experiments, ventral midbrain precursors (including dopaminergic precursors) and their descendants were labeled in a mosaic manner using a Cre/loxP based inducible fate mapping system 6.

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

A method to generate organotypic slices from the E12.5 murine embryonic midbrain is described. The organotypic slice cultures can be used to observe the behavior of dopaminergic neurons or other ventral midbrain neurons.

Cultures organotypique de mésencéphale ventral embryonnaire: Un système pour étudier le développement neuronal dopaminergique In vitro</em

The  mouse is an excellent model organism to study mammalian brain development due  to the abundance of molecular and genetic data. However, the ...

VisioTracker, an Innovative Automated Approach to Oculomotor Analysis

Investigations into the visual system development and function necessitate quantifiable behavioral models of visual performance that are easy to elicit, robust, and simple to manipulate. A suitable model has been found in the optokinetic response (OKR), a reflexive behavior present in all vertebrates due to its high selection value. The OKR involves slow stimulus-following movements of eyes alternated with rapid resetting saccades. The measurement of this behavior is easily carried out in zebrafish larvae, due to its early and stable onset (fully developed after 96 hours post fertilization (hpf)), and benefitting from the thorough knowledge about zebrafish genetics, for decades one of the favored model organisms in this field. Meanwhile the analysis of similar mechanisms in adult fish has gained importance, particularly for pharmacological and toxicological applications.
Here we describe VisioTracker, a fully automated, high-throughput system for quantitative analysis of visual performance. The system is based on research carried out in the group of Prof. Stephan Neuhauss and was re-designed by TSE Systems. It consists of an immobilizing device for small fish monitored by a high-quality video camera equipped with a high-resolution zoom lens. The fish container is surrounded by a drum screen, upon which computer-generated stimulus patterns can be projected. Eye movements are recorded and automatically analyzed by the VisioTracker software package in real time.
Data analysis enables immediate recognition of parameters such as slow and fast phase duration, movement cycle frequency, slow-phase gain, visual acuity, and contrast sensitivity.
Typical results allow for example the rapid identification of visual system mutants that show no apparent alteration in wild type morphology, or the determination of quantitative effects of pharmacological or toxic and mutagenic agents on visual system performance.

The VisioTracker is an automated system for the quantitative analysis of visual performance of larval and small adult fish based on the recording of eye movements. It features full control over visual stimulus properties and real-time analysis, enabling high-throughput research in fields such as visual system development and function, pharmacology, neural circuit studies and sensorimotor integration.

VisioTracker, une approche novatrice automatisé pour l&#39;analyse oculomotrice

Investigations into the visual system development and function necessitate quantifiable behavioral models of visual performance that are easy to elicit, ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

Rapide Micro-iontophorèse de glutamate et le GABA: un outil utile d&#39;étudier l&#39;intégration synaptique

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Sex Stratified Neuronal Cultures to Study Ischemic Cell Death Pathways

Sex differences in neuronal susceptibility to ischemic injury and neurodegenerative disease have long been observed, but the signaling mechanisms responsible for those differences remain unclear. Primary disassociated embryonic neuronal culture provides a simplified experimental model with which to investigate the neuronal cell signaling involved in cell death as a result of ischemia or disease; however, most neuronal cultures used in research today are mixed sex. Researchers can and do test the effects of sex steroid treatment in mixed sex neuronal cultures in models of neuronal injury and disease, but accumulating evidence suggests that the female brain responds to androgens, estrogens, and progesterone differently than the male brain. Furthermore, neonate male and female rodents respond differently to ischemic injury, with males experiencing greater injury following cerebral ischemia than females. Thus, mixed sex neuronal cultures might obscure and confound the experimental results; important information might be missed. For this reason, the Herson Lab at the University of Colorado School of Medicine routinely prepares sex-stratified primary disassociated embryonic neuronal cultures from both hippocampus and cortex. Embryos are sexed before harvesting of brain tissue and male and female tissue are disassociated separately, plated separately, and maintained separately. Using this method, the Herson Lab has demonstrated a male-specific role for the ion channel TRPM2 in ischemic cell death. In this manuscript, we share and discuss our protocol for sexing embryonic mice and preparing sex-stratified hippocampal primary disassociated neuron cultures. This method can be adapted to prepare sex-stratified cortical cultures and the method for embryo sexing can be used in conjunction with other protocols for any study in which sex is thought to be an important determinant of outcome.

Primary disassociated embryonic hippocampal neuronal cultures are useful for investigating the signaling mechanisms involved in neuron death. Sexing the embryos before the isolation and dissociation of the hippocampus allows the preparation of separate male and female cultures, which enables the researcher to identify and investigate sex-specific cell signaling.

Sex differences in neuronal susceptibility to ischemic injury and neurodegenerative disease have long been observed, but the signaling mechanisms ...

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

Une procédure d&#39;implantation de tableaux organisés de microfils pour enregistrements unitaires dans Awake, se comportant Animaux

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Manipulation génétique des neurones du cervelet granules<em&gt; In Vitro</em&gt; Et<em&gt; In Vivo</em&gt; À étudier neuronale Morphologie et migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Neural Activity Propagation in an Unfolded Hippocampal Preparation with a Penetrating Micro-electrode Array

This protocol describes a method for preparing a new in vitro flat hippocampus preparation combined with a micro-machined array to map neural activity in the hippocampus. The transverse hippocampal slice preparation is the most common tissue preparation to study hippocampus electrophysiology. A longitudinal hippocampal slice was also developed in order to investigate longitudinal connections in the hippocampus. The intact mouse hippocampus can also be maintained in vitro because its thickness allows adequate oxygen diffusion. However, these three preparations do not provide direct access to neural propagation since some of the tissue is either missing or folded. The unfolded intact hippocampus provides both transverse and longitudinal connections in a flat configuration for direct access to the tissue to analyze the full extent of signal propagation in the hippocampus in vitro. In order to effectively monitor the neural activity from the cell layer, a custom made penetrating micro-electrode array (PMEA) was fabricated and applied to the unfolded hippocampus. The PMEA with 64 electrodes of 200 &#181;m in height could record neural activity deep inside the mouse hippocampus. The unique combination of an unfolded hippocampal preparation and the PMEA provides a new in-vitro tool to study the speed and direction of propagation of neural activity in the two-dimensional CA1-CA3 regions of the hippocampus with a high signal to noise ratio.

We have developed an in vitro unfolded hippocampus which preserves CA1-CA3 array of neurons. Combined with the penetrating micro-electrode array, neural activity can be monitored in both the longitudinal and transverse orientations. This method provides advantages over hippocampal slice preparations as the propagation in the entire hippocampus can be recorded simultaneously.

Neural activité Propagation dans un hippocampe Préparation déplié avec un pénétrant Micro-réseau d&#39;électrodes

This protocol describes a method for preparing a new in vitro flat hippocampus preparation combined with a micro-machined array to map neural activity in ...

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 ...

In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively thin to properly visualize and access neurons for patch-clamping or imaging, and in vitro examination of brain circuitry is limited to only what is physically present in the acute slice. To maintain the benefits of in vitro slice experimentation while preserving a larger portion of presynaptic nuclei, we developed a novel slice preparation. This &#8220;wedge slice&#8221; was designed for patch-clamp electrophysiology recordings to characterize the diverse monaural, sound-driven inputs to medial olivocochlear (MOC) neurons in the brainstem. These neurons receive their primary afferent excitatory and inhibitory inputs from neurons activated by stimuli in the contralateral ear and corresponding cochlear nucleus (CN). An asymmetrical brain slice was designed which is thickest in the rostro-caudal domain at the lateral edge of one hemisphere and then thins towards the lateral edge of the opposite hemisphere. This slice contains, on the thick side, the auditory nerve root conveying information about auditory stimuli to the brain, the intrinsic CN circuitry, and both the disynaptic excitatory and trisynaptic inhibitory afferent pathways that converge on contralateral MOC neurons. Recording is performed from MOC neurons on the thin side of the slice, where they are visualized using DIC optics for typical patch-clamp experiments. Direct stimulation of the auditory nerve is performed as it enters the auditory brainstem, allowing for intrinsic CN circuit activity and synaptic plasticity to occur at synapses upstream of MOC neurons. With this technique, one can mimic in vivo circuit activation as closely as possible within the slice. This wedge slice preparation is applicable to other brain circuits where circuit analyses would benefit from preservation of upstream connectivity and long-range inputs, in combination with the technical advantages of in vitro slice physiology.

Integration of diverse synaptic inputs to neurons is best measured in a preparation that preserves all pre-synaptic nuclei for natural timing and circuit plasticity, but brain slices typically sever many connections. We developed a modified brain slice to mimic in vivo circuit activity while maintaining in vitro experimentation capability.

Préparation in vitro en tranches compensées pour imiter la connectivité du circuit neuronal In Vivo

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively ...