This work was partially supported by National Institute of Deafness and Other Communications Disorders Awards R03-DC-012125 to D. A. Llano and F31-DC-013501 to B. J. Slater as well as the Carver Foundation.
The authors would like to thank Jason MacLean and Matthew Banks for technical advice with calcium imaging.

Toutes les procédures ont été approuvées par le Comité de protection des animaux et l&#39;utilisation institutionnelle à l&#39;Université de l&#39;Illinois. Tous les animaux ont été logés dans des installations de soins des animaux approuvées par l&#39;Association américaine pour l&#39;accréditation du Laboratoire de protection des animaux. Tout a été fait pour réduire au minimum le nombre d&#39;animaux utilisés et de réduire la souffrance à toutes les étapes de l&#39;étude. 1. Préparation et suppression de Brain de souris pour le tranchage <ol><li> Préparez-vous à perfusion et une tranche incubation. <ol><li> Environ 30 min avant de trancher, préparer une solution de saccharose de coupe haute et basse aCSF de calcium pour l&#39;incubation de la tranche avant l&#39;enregistrement ou l&#39;imagerie. </li><li> Disposez tous les outils nécessaires (grands ciseaux émoussés finaux, petits ciseaux à ressort, forceps anatomiques, de grands ciseaux ou d&#39;une guillotine, iris ciseaux, bijoutiers forceps, seringue de 10 ml avec une aiguille 27G x 1/2 pouces, petit morceau de 1 cm x 2 cm filtres papier, et un bout pipette Pasteur rompu pour sltransfert de la glace) pour perfusion et tranchage, et préparer plateau de perfusion. </li><li> Mettre en place la chambre avec oxygénée aCSF incubation dans un bain-marie à 32 o et une boîte de culture rempli de solution de coupe. </li></ol></li><li> Préparer étape de coupe pour trancher. <ol><li> Coupez un petit morceau de 3% de gélose environ 1 cm 3 pour l&#39;utiliser comme un filet de sécurité pour le cerveau et 1,5 cm x 1,5 mm x 3 mm pour une bosse à être utilisé pour soutenir l&#39;IC. </li><li> Coller la butée sur la scène avec de la colle cyanoacrylate ainsi que la bosse sur le côté droit de la butée de telle sorte qu&#39;elles forment un angle de 80 o. </li></ol></li><li> Retirer le cerveau. <ol><li> Profondément anesthésier une p12-p20 (idéalement p14-18) de la souris avec la kétamine 100 mg / kg et xylaxine 3 mg / kg (ou comité d&#39;éthique comparables procédure approuvée). Remarque: Confirmation du niveau d&#39;anesthésie appropriée par l&#39;absence de réponse à pincement de l&#39;orteil. Comme l&#39;animal est sous anesthésie brièvement, pommade ophtalmique est inutile. </li><li> Utilisation émousséefin ciseaux, exposer la cage thoracique de la pointe du sternum à la nuque. <ol><li> Trouver la pointe du sternum, et faire une coupe horizontale d&#39;environ 1,5 cm. </li><li> Faites deux coupes verticales à partir des extrémités de la coupe horizontale précédente sur les épaules, d&#39;environ 1,5 cm chaque. </li></ol></li><li> Couper à travers le diaphragme et les jonctions costochondral pour exposer le coeur. </li><li> Avec de petits ciseaux à ressort, faire une petite, 2-5 mm couper dans l&#39;oreillette droite, à partir de la dorsale ventrale à côté du cœur. </li><li> En utilisant une seringue de 10 ml avec une aiguille 27G x 1/2 pouces, injecter le ventricule gauche et perfuser rapidement l&#39;animal avec une solution de saccharose coupe élevé. </li><li> Une fois que le sang soit claire, utiliser de plus grandes ciseaux pour enlever la tête. Note: Nous avons pas systématiquement évalué l&#39;utilité de la perfusion. Cependant, dans notre expérience, transcardiaque perfusion chez les souris jeunes ce qui peut être fait avec près de 100% de réussite, et a l&#39;avantage d&#39;éliminer bl rougecellules Ood, peut fluo, et des interférences avec l&#39;imagerie. </li><li> Couper la peau vers le bas de la ligne médiane pour exposer le crâne. </li><li> Utilisation iris pliés ciseaux, couper le crâne entre les yeux, puis à partir de la coupure entre les yeux, couper soigneusement d&#39;avant en arrière le long de la ligne médiane de suture en prenant soin de ne pas endommager le cerveau en dessous. Remarque: La dure vient habituellement hors du crâne. Si nécessaire, retirez la dure-mère avant de retirer soigneusement le cerveau. </li><li> Utilisation bijoutiers pince, levier ouvrir le crâne et retirez soigneusement le cerveau, puis placez le cerveau dans une solution de coupe. Prenez soin de ne pas endommager le cortex. </li></ol></li></ol> 2. Préparation de cerveau pour le tranchage <ol><li> Préparation cerveau pour le montage sur vibratome scène. <ol><li> En utilisant une lame marquée par deux lignes à 90 ° et une ligne diagonale à 17 o à partir du haut de gauche à droite en bas, coupant où les deux lignes se rencontrent, placez le côté cérébral dorsale jusqu&#39;à, en utilisantune lame de rasoir, retirer 2-4 mm de l&#39;extrémité rostrale du cerveau créer une surface plane. </li><li> Placez le côté caudal du cerveau sur la surface plane nouvellement créée, et aligner la surface dorsale du cerveau avec l&#39;horizontale et la ligne médiane du cerveau avec la ligne verticale. </li><li> Aligner la lame de rasoir avec la ligne 17 de o, inclinez le rasoir à un angle de 30 o et enlever environ 3 mm du cortex droit dans une coupe en diagonale à double (17 o de plan horizontal et 60 o de la coronale). </li></ol></li><li> Montage sur le cerveau vibratome scène. <ol><li> Placer un petit morceau de papier filtre de 1 cm x 2 cm sur la face ventrale du cerveau de sorte que la dimension longue est perpendiculaire à la ligne médiane. </li><li> Soigneusement appliquer une petite quantité de colle cyanoacrylate à la zone en face de la butée arrière (et à gauche de la bosse). </li><li> Placez le double visage du cerveau en diagonale coupé à la colle de sorte que la partie caudale du cerveau et du cerveau postérieur sont proppé sur la bosse et le côté droit du cerveau est contre la butée. </li><li> Délicatement Appuyez sur le cerveau, veiller à ce que toute la surface est en contact avec la chambre de tranchage. </li></ol></li></ol> 3. Obtenir l&#39;Colliculo-thalamocortical Slice <ol><li> De prendre rapidement l&#39;étape de coupe et le placer dans le vibratome, remplir la scène avec une solution de coupe. </li><li> Alignez la lame avec le haut (côté ventral) du cerveau. </li><li> Retirer 1-1,5 mm de la partie supérieure du cerveau, retirez 300-500 um tranches et d&#39;évaluer la profondeur après l&#39;enlèvement. Remarque: Lors de la CI, le MGB et le corps genouillé latéral (CGL) sont tous visibles, et le gyrus denté forme une forme de «C» de prendre une à deux tranches de 600 um. Voir la figure 3A. </li><li> Placer les tranches de chambre chauffée (32 ° C) tenant. </li></ol> 4. Imagerie de la tranche <ol><li> Préparation pour l&#39;imagerie flavoprotéine d&#39;autofluorescence. <ol> &lt;li&gt; Préparez aCSF pour la perfusion de la tranche dans une chambre d&#39;enregistrement électrophysiologique standard, bulle aCSF avec 95% de O 2/5% de CO 2. </li><li> Tirez électrode de verre pour la stimulation. Notez que plusieurs différentes électrodes de stimulation et de configurations (verre, le tungstène, la fibre de carbone, monopolaire, bipolaire) tous les travaux très bien en conjonction avec l&#39;imagerie flavoprotéine d&#39;autofluorescence. </li><li> Tournez sur le logiciel de capture d&#39;image logiciel de stimulation ordinateur, caméra, micromanipulateur, source de lumière,,. </li><li> Perfuser la chambre d&#39;enregistrement avec oxygénée aCSF, placez pont personnalisé (de conception disponibles dans des matériaux supplémentaires) pour élever tranche dans la chambre. Remarque: le taux de perfusion peut varier avec tout individu mis en place; 5-15 ml / min est utilisé ici. </li><li> Placez une tranche dans la chambre, abaisser le niveau de liquide dans la chambre pour empêcher la tranche de levage du pont tout en plaçant la harpe spécialement construit sur tranche, en prenant soin d&#39;éviter de placer cordes de la harpe sur les voies d&#39;interest (figure 1A). Augmenter le niveau de liquide à environ 1-2 mm au-dessus tranche pour maintenir le débit aCSF dessus et en dessous tranche. Remarque: Si un repositionnement est nécessaire, utiliser la fin pipette Pasteur cassé ou laisser le niveau de aCSF se lever et utiliser une pince avec un soin extrême. </li><li> Insérez argent électrode de chlorure dans électrode de verre rempli de aCSF, insérez l&#39;électrode de verre dans micromanipulateur et se connecter à un stimulus isolateur, et placer soigneusement l&#39;électrode de verre dans IC / matière blanche conduisant à thalamus (voir la figure 1A). </li></ol></li><li> Flavoprotéine acquisition d&#39;image d&#39;autofluorescence. <ol><li> Recueillir des images à 4 Hz (en utilisant un 2X macro objectif corrigé à l&#39;infini (NA 0,13)) et une caméra pour 105 sec tout en stimulant électriquement (chaque stimulation se compose de 1 sec de 40 Hz 2 ms impulsions tissus à 0,05 Hz cinq fois) tout en éclairant le tissu avec la lumière bleue de 470 à 490 nm et la capture ci-dessus 515 nm. Assurez-vous que l&#39;image est ni àVoir o sombre, ni soufflé Figure 3 colonne de gauche pour référence. Remarque: le temps de la collection, la fréquence de collecte, et la fréquence de stimulation peuvent être modifiés en fonction de l&#39;expérience, le programme personnalisé inclus dans du matériel supplémentaire peut être modifiée en tant que telle. </li><li> Exporter des images vers Matlab, et en utilisant le programme écrit sur mesure disponibles dans des matériaux supplémentaires pour analyser la puissance spectrale des images à 0,05 Hz. L&#39;image résultante montrera voies connectés. Remarque: Le programme personnalisé utilise une transformée de Fourier rapide des valeurs de pixels dans la série de temps pour produire l&#39;image. </li></ol></li><li> Préparation pour l&#39;imagerie de calcium. <ol><li> Préparer petite chambre d&#39;incubation à l&#39;aide d&#39;une boîte de culture de 3 cm et la membrane de culture soulevé pour permettre aCSF pour atteindre à la fois le haut et le bas de la tranche, et le placer sur un coussin chauffant pour maintenir la température à environ 32 ° C </li><li> Remplissez petite chambre d&#39;incubation avec aCSF chaude et lentement bulle avec 95% d&#39;O2/5% de CO 2. </li><li> Mélanger 2 pi de pluronique acide F-127 et 50 pg de Fura-2 heures dissous dans 48 pi de DMSO. </li><li> Lieu tranche dans la petite chambre d&#39;incubation sur la membrane soulevée et soigneusement à l&#39;aide d&#39;une micropipette Ajouter le mélange de coloration directement au-dessus de la BTP (ou autre structure d&#39;intérêt). </li><li> Couvrir de tranches pour prévenir le blanchiment avant l&#39;imagerie, et permettent tranches à incuber pendant 45 min - 1 h. </li><li> Retirer les tranches de chambre de retenue chauffée pendant 10-15 min pour laver l&#39;excès de matériau. </li><li> Suivez les étapes 4.1.1 à 4.1.6 pour la préparation à stimuler et à l&#39;image de la tranche de colliculo-thalamocortical. </li></ol></li><li> Calcium acquisition d&#39;image. <ol><li> Recueillir des images à 10 Hz (en utilisant un objectif à immersion 20X d&#39;eau) pendant 25 secondes tout en stimulant électriquement (chaque stimulation se compose d&#39;une impulsion de 2 ms) tissu à 0,2 Hz cinq fois tout en éclairant le tissu avec 365 nm de lumière et la capture de fluorescence au-dessus de 510 nm. </li&gt; <li> Exporter des images vers Matlab et en utilisant le programme écrit sur mesure disponibles dans des matériaux supplémentaires pour analyser la puissance spectrale des images à 0,2 Hz. L&#39;image résultante montrera cellules actives. </li></ol></li></ol>

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The authors have nothing to disclose.

This protocol describes improvements upon a previously described colliculo-thalamocortical brain slice in p12-20 mouse to study information flow in the auditory system1. This method has a number of advantages over other, similar, brain slice preparations by retaining connections between more brain areas in a single slice, which gives investigators new tools to understand the interaction and interplay between auditory nuclei in the forebrain. There have been a few key modifications in this protocol, compared to...

Dans le système auditif, bien qu&#39;il y ait une transformation substantielle d&#39;informations entre la périphérie sensorielle et le colliculus inférieur, il existe un traitement supplémentaire considérable avant d&#39;atteindre le cortex auditif. Nous savons très peu sur la façon dont la transformation est effectuée et donc peu sur la façon dont cette transformation permet au cerveau pour interpréter l&#39;information sensorielle entrante. À l&#39;exception de l&#39;olfaction, chacun des sens a une organisation très similaire avec les signaux périphériques étant initialement relayés avec une haute fidélité qui diminue à mesure que le signal monte au cortex. Le cortex envoie alors projections aux structures inférieures à moduler davantage l&#39;information entrante. Ce système complexe a été étudiée dans une variété de manières in vivo, ainsi que dans un certain nombre de préparations in vitro. Dans le premier, toutes les connexions sont intactes, ce qui permet au chercheur de sonder tout ensemble de connexions, tout en contrôlant l&#39;entrée sensorielle et mesuresortie dans une zone donnée. Avec cette approche, il ya peu ou pas de contrôle de la grande variété d&#39;autres intrants, y compris d&#39;autres entrées sensorielles, l&#39;excitation et l&#39;attention, donnant lieu à une sortie intensément complexe. In vitro, des tranches de cerveau ont été coupés pour capturer soit un ensemble unique des projections, ou deux zones du cerveau connectés, qui permettent aux chercheurs de stimuler et évaluer afférences ou zones cérébrales différentes. Ce sont souvent des tranches soit thalamocorticales ou tectothalamic où soit l&#39;entrée du thalamus ou du thalamus et sa sortie vers le cortex sont conservés 5.2. Ces préparations permettent une grande variété de manipulations pharmacologiques, électriques et optogénétiques. Cependant, avec seulement deux régions du cerveau, ils évaluent essentiellement le transfert d&#39;informations et manquent de la capacité d&#39;évaluer la transformation de l&#39;information qui passe dans le thalamus. Aussi la projection de réticulo-thalamique, qui peuvent jouer un rôle dans l&#39;attention modulation 6-9 est prESENT dans cette tranche. Ici, nous démontrons améliorations sur notre préparation précédente 1, qui permet le contrôle de l&#39;investigateur de diverses entrées du thalamus pour donner une perspective unique de la façon dont les portes thalamus et filtres informations. Nous coupler cette nouvelle préparation de tranche avec l&#39;imagerie flavoprotéine d&#39;autofluorescence pour évaluer la connectivité de tranche et l&#39;analyse d&#39;activation à grande échelle, l&#39;imagerie de calcium dans le thalamus pour neuronale analyse de la population, et l&#39;enregistrement d&#39;une cellule unique de mesurer l&#39;impact des différentes entrées au niveau de la cellule unique. Pour aider à maintenir ces relations étendues, nous avons également développé un certain nombre de modifications de l&#39;ancrage de la tranche normale (aka &quot;harpe&quot;) pour maintenir la tranche de cerveau en place et un pont pour élever la tranche pour perfusion améliorée. La harpe est conçu en forme de fer à cheval modifié pour entourer la tranche et permettrait aux points de fixation personnalisables pour les cordes de la harpe. Trois chaînes sontattaché de telle sorte que i) une se trouve à l&#39;horizontale le long du bord interne de la tranche, ii) on étend à partir du bord extrême de la puce vers le bord caudal du secteur et iii) une prolonge en diagonale à partir du bord interne de la tranche à une zone rostrale à l&#39;AC (voir la figure 1A). Petites indentations dans le cadre pour le collage (avec de la colle cyanoacrylate) de la harpe cordes permettent une diminution de la quantité de pression sur la tranche pour aider à maintenir l&#39;intégrité de la tranche (voir la figure 1B). En utilisant l&#39;impression en trois dimensions, nous sommes en mesure de harpes de conception personnalisée à nos spécifications uniques, ainsi que des ponts qui permettent l&#39;écoulement idéal de liquide céphalo-rachidien artificiel (aCSF) dessus et en dessous du tissu. Cela maintient également de grandes zones à la lumière pour pénétrer dans le tissu pour le patch clamp électrophysiologie.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>High sucrose cutting solution</td>
			<td></td>
			<td></td>
			<td>in mM: 206 sucrose, 10.0 MgCl2, 11.0 glucose, 1.25 NaH2PO4, 26 NaHCO3, 0.5 CaCl2, 2.5 KCl, pH 7.4</td>
		</tr>
		<tr>
			<td>Low calcium aCSF</td>
			<td></td>
			<td></td>
			<td>in mM: 126 NaCl, 3.0 MgCl2, 10.0 glucose, 1.25 NaH2PO4, 26 NaHCO3, 1.0 CaCl2, 2.5 KCl, pH 7.4</td>
		</tr>
		<tr>
			<td>aCSF</td>
			<td></td>
			<td></td>
			<td>in mM: 126 NaCl, 2.0 MgCl2, 10.0 glucose, 1.25 NaH2PO4, 26 NaHCO3, 2.0 CaCl2, 2.5 KCl, pH 7.4</td>
		</tr>
		<tr>
			<td>Stimulus Isolator</td>
			<td>World Precision Instruments</td>
			<td>A360</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Life Technologies</td>
			<td>D12345</td>
			<td>Lot: 1572C502</td>
		</tr>
		<tr>
			<td>Fura-2AM</td>
			<td>Life Technologies</td>
			<td>F1201</td>
			<td>Lot: 144912</td>
		</tr>
		<tr>
			<td>Pluronic F-127</td>
			<td>Life Technologies</td>
			<td>P3000MP</td>
			<td>Lot: 1499369</td>
		</tr>
		<tr>
			<td>Large culture dish</td>
			<td>Fisherbrand</td>
			<td>08-757-13</td>
			<td>100x15mm culture dish</td>
		</tr>
		<tr>
			<td>Small culture dish</td>
			<td>Falcon</td>
			<td align="right">353001</td>
			<td>35x10mm culture dish</td>
		</tr>
		<tr>
			<td>Raised culture membrane</td>
			<td>Millicell</td>
			<td>PICMORG50</td>
			<td>Used to maintain oxygenated fluid perfusion on both sides of slice.</td>
		</tr>
		<tr>
			<td>Flavoprotein imaging fluorescence cube</td>
			<td>Olympus</td>
			<td>UMNIB</td>
			<td>470&#8211;490 nm excitation, 505 nm dichroic, 515 nm emission long pass.&#160; We have found that virtually any green fluorescence protein filter cube will work here.</td>
		</tr>
		<tr>
			<td>Calcium imaging fluorescence cube</td>
			<td>Omega Optical</td>
			<td>BX-18</td>
			<td>XF1005 365nm exitation, XF2001 400nm dichroic, XF3080 510nm emission</td>
		</tr>
		<tr>
			<td>Agar for blocking brain</td>
			<td></td>
			<td></td>
			<td>3% by weight in water</td>
		</tr>
		<tr>
			<td>Viper si Stereo Lithography Apparatus</td>
			<td>3D Systems</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

modification of a colliculo-thalamocortical mouse brain slice, incorporating 3-d printing of chamber components and multi-scale optical imaging

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study these two systems and how they interact has been with the use of in vivo preparations. Major advances have been made with acute brain slices containing the thalamocortical and cortico-thalamic pathways in the somatosensory, visual, and auditory systems. With key refinements to our recent modification of the auditory thalamocortical slice1, we are able to more reliably capture the projections between most of the major auditory midbrain and forebrain structures: the inferior colliculus (IC), medial geniculate body (MGB), thalamic reticular nucleus (TRN), and the auditory cortex (AC). With portions of all these connections retained, we are able to answer detailed questions that complement the questions that can be answered with in vivo preparations. The use of flavoprotein autofluorescence imaging enables us to rapidly assess connectivity in any given slice and guide the ensuing experiment. Using this slice in conjunction with recording and imaging techniques, we are now better equipped to understand how information processing occurs at each point in the auditory forebrain as information ascends to the cortex, and the impact of descending cortical modulation. 3-D printing to build slice chamber components permits double-sided perfusion and broad access to networks within the slice and maintains the widespread connections key to fully utilizing this preparation.

Un exemple de cerveau de souris tranche de colliculo-thalamocortical obtenu dans P15 souris est présentée dans la figure 2. La tranche idéal contiendra les quatre grandes structures auditives mésencéphale et du cerveau antérieur IC, MGB, TRN, et AC, qui sont tous activés lorsque l&#39;IC est stimulée (figure 2A). En utilisant l&#39;analyse de Fourier, la puissance spectrale est mesurée à la fréquence de la stimulation électrique, avec des régions cérébral...

Watch this Scientific Journal Video about Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging at JoVE.com

Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging

L&#39;utilisation de plusieurs angles à couper le cerveau des petits de la souris, nous améliorons sur une tranche de cerveau aiguë décrite précédemment qui capture les liens entre la plupart des grands du mésencéphale et du cerveau antérieur structures auditives.

Modification d&#39;un Colliculo-thalamocortical cerveau de souris Slice, Intégrer l&#39;impression 3-D de la Chambre des composants et multi-échelle imagerie optique

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study ...

modification-colliculo-thalamocortical-mouse-brain-slice

Research

JoVE Journal

Neuroscience

8.2K vues.  University of Illinois Urbana-Champaign.  L'utilisation de plusieurs angles à couper le cerveau des petits de la souris, nous améliorons sur une tranche de cerveau aiguë décrite précédemment qui capture les liens entre la plupart des grands du mésencéphale et du cerveau antérieur structures auditives. 

Vidéo: Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging

Scalable Fluidic Injector Arrays for Viral Targeting of Intact 3-D Brain Circuits

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted in neurological and psychiatric disorders--would be greatly facilitated by a technology for rapidly targeting genes to complex 3-dimensional neural circuits, enabling fast creation of "circuit-level transgenics." We have recently developed methods in which viruses encoding for light-sensitive proteins can sensitize specific cell types to millisecond-timescale activation and silencing in the intact brain. We here present the design and implementation of an injector array capable of delivering viruses (or other fluids) to dozens of defined points within the 3-dimensional structure of the brain (Figure. 1A, 1B). The injector array comprises one or more displacement pumps that each drive a set of syringes, each of which feeds into a polyimide/fused-silica capillary via a high-pressure-tolerant connector. The capillaries are sized, and then inserted into, desired locations specified by custom-milling a stereotactic positioning board, thus allowing viruses or other reagents to be delivered to the desired set of brain regions. To use the device, the surgeon first fills the fluidic subsystem entirely with oil, backfills the capillaries with the virus, inserts the device into the brain, and infuses reagents slowly (&lt;0.1 microliters/min). The parallel nature of the injector array facilitates rapid, accurate, and robust labeling of entire neural circuits with viral payloads such as optical sensitizers to enable light-activation and silencing of defined brain circuits. Along with other technologies, such as optical fiber arrays for light delivery to desired sets of brain regions, we hope to create a toolbox that enables the systematic probing of causal neural functions in the intact brain. This technology may not only open up such systematic approaches to circuit-focused neuroscience in mammals, and facilitate labeling of brain regions in large animals such as non-human primates, but may also open up a clinical translational path for cell-specific optical control prosthetics, whose precision may enable improved treatment of intractable brain disorders. Finally, such devices as described here may facilitate precisely-timed fluidic delivery of other payloads, such as stem cells and pharmacological agents, to 3-dimensional structures, in an easily user-customizable fashion.

Controlling and analyzing neural circuits in vivo would be facilitated by a technology for delivery of viruses and other reagents to desired 3-dimensional sets of brain regions. We demonstrate customized fluidic injector array fabrication, and delivery of virally-encoded optical sensitizers, enabling optical manipulation of complex brain circuits.

Évolutif fluidique Arrays Injecteur pour virale ciblage d&#39;Intact circuits du cerveau en 3-D

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted ...

Recherche

Neurosciences

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

Multiple-Mouse Imaging neuroanatomiques par résonance magnétique

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Imaging Pheromone Sensing in a Mouse Vomeronasal Acute Tissue Slice Preparation

Peter Karlson and Martin L&uuml;scher used the term pheromone for the first time in 19591 to describe chemicals used for intra-species communication. Pheromones are volatile or non-volatile short-lived molecules2 secreted and/or contained in biological fluids3,4, such as urine, a liquid known to be a main source of pheromones3. Pheromonal communication is implicated in a variety of key animal modalities such as kin interactions5,6, hierarchical organisations3 and sexual interactions7,8 and are consequently directly correlated with the survival of a given species9,10,11. In mice, the ability to detect pheromones is principally mediated by the vomeronasal organ (VNO)10,12, a paired structure located at the base of the nasal cavity, and enclosed in a cartilaginous capsule. Each VNO has a tubular shape with a lumen13,14 allowing the contact with the external chemical world. The sensory neuroepithelium is principally composed of vomeronasal bipolar sensory neurons (VSNs)15. Each VSN extends a single dendrite to the lumen ending in a large dendritic knob bearing up to 100 microvilli implicated in chemical detection16. Numerous subpopulations of VSNs are present. They are differentiated by the chemoreceptor they express and thus possibly by the ligand(s) they recognize17,18. Two main vomeronasal receptor families, V1Rs and V2Rs19,20,21,22, are composed respectively by 24023 and 12024 members and are expressed in separate layers of the neuroepithelium. Olfactory receptors (ORs)25 and formyl peptide receptors (FPRs)26,27 are also expressed in VSNs.
Whether or not these neuronal subpopulations use the same downstream signalling pathway for sensing pheromones is unknown. Despite a major role played by a calcium-permeable channel (TRPC2) present in the microvilli of mature neurons28 TRPC2 independent transduction channels have been suggested6,29. Due to the high number of neuronal subpopulations and the peculiar morphology of the organ, pharmacological and physiological investigations of the signalling elements present in the VNO are complex.
Here, we present an acute tissue slice preparation of the mouse VNO for performing calcium imaging investigations. This physiological approach allows observations, in the natural environment of a living tissue, of general or individual subpopulations of VSNs previously loaded with Fura-2AM, a calcium dye. This method is also convenient for studying any GFP-tagged pheromone receptor and is adaptable for the use of other fluorescent calcium probes. As an example, we use here a VG mouse line30, in which the translation of the pheromone V1rb2 receptor is linked to the expression of GFP by a polycistronic strategy.

In mice, the ability to detect pheromones is principally mediated by the vomeronasal organ (VNO). Here, an acute tissue slice preparation of VNO for performing calcium imaging is described. This physiological approach allows observations of subpopulations and/or individual neurons in a living tissue and is convenient for receptor-ligand identification.

Imagerie phéromones détection dans une souris Vomeronasal tissus Préparation Trancher aiguë

Peter Karlson and Martin L&uuml;scher used the term pheromone for the first time in 19591 to describe chemicals used for intra-species communication. ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Une procédure pour l&#39;implantation d&#39;une chambre pour la colonne vertébrale longitudinale<em&gt; In Vivo</em&gt; Imagerie du cordon spinal de souris

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Whole-mount Imaging of Mouse Embryo Sensory Axon Projections

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we describe a method to label in situ a subset of dorsal root ganglion (DRG) axon projections to assess their phenotypic characteristics using several genetically manipulated mouse lines. The TrkA-positive neurons are nociceptor neurons, dedicated to the transmission of pain signals. We utilize a TrkAtaulacZ mouse line to label the trajectories of all TrkA-positive peripheral axons in the intact mouse embryo. We further breed the TrkAtaulacZ line onto a Bax null background, which essentially abolishes neuronal apoptosis, in order to assess growth-related questions independently of possible effects of genetic manipulations on neuronal survival. Subsequently, genetically modified mice of interest are bred with the TrkAtaulacZ/Bax null line and are then ready for study using the techniques described herein. This presentation includes detailed information on mouse breeding plans, genotyping at the time of dissection, tissue preparation, staining and clearing to allow for visualization of full-length axonal trajectories in whole-mount preparation.

We present here an optimized protocol to genotype, stain and prepare fetal mice for the imaging of peripheral nociceptor axon projections in the whole animal, as an effective method to assess sensory axon growth phenotypes in developing genetically engineered mice.

Whole-monter imagerie de l&#39;embryon de souris projections sensorielles Axon

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we ...

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma establishes a lifelong persistent infection in the brain. While this brain infection is asymptomatic in most immunocompetent people, in the developing fetus or immunocompromised individuals such as acquired immune deficiency syndrome (AIDS) patients, this predilection for and persistence in the brain can lead to devastating neurologic disease. Thus, it is clear that the brain-Toxoplasma interaction is critical to the symptomatic disease produced by Toxoplasma, yet we have little understanding of the cellular or molecular interaction between cells of the central nervous system (CNS) and the parasite. In the mouse model of CNS toxoplasmosis it has been known for over 30 years that neurons are the cells in which the parasite persists, but little information is available about which part of the neuron is generally infected (soma, dendrite, axon) and if this cellular relationship changes between strains. In part, this lack is secondary to the difficulty of imaging and visualizing whole infected neurons from an animal. Such images would typically require serial sectioning and stitching of tissue imaged by electron microscopy or confocal microscopy after immunostaining. By combining several techniques, the method described here enables the use of thick sections (160 &#181;m) to identify and image whole cells that contain cysts, allowing three-dimensional visualization and analysis of individual, chronically infected neurons without the need for immunostaining, electron microscopy, or serial sectioning and stitching. Using this technique, we can begin to understand the cellular relationship between the parasite and the infected neuron.

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Using this protocol, we were able to image 160 &#181;m thick brain sections from mice infected with the parasite Toxoplasma gondii, which enables visualization and analysis of the spatial relationship between the encysting parasite and the infected neuron.

Imagerie 3-D et de l&#39;analyse des neurones infecté<em&gt; In Vivo</em&gt; Avec<em&gt; Toxoplasma gondii</em

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma ...

A Large Lateral Craniotomy Procedure for Mesoscale Wide-field Optical Imaging of Brain Activity

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, typically 3 mm x 3 mm. This protocol introduces a method for creating a substantially larger 7 mm x 6 mm cranial window exposing most of a cerebral hemisphere over the mouse temporal and parietal cortices (e.g., bregma 2.5 - 4.5 mm, lateral 0 - 6 mm). To perform this surgery, the head must be tilted approximately 30&#176; and much of the temporal muscle must be retracted. Due to the large amount of bone removal, this procedure is intended only for acute experiments with the animal anesthetized throughout the surgery and experiment.
The main advantage of this innovative large lateral cranial window is to provide simultaneous access to both medial and lateral areas of the cortex. This large unilateral cranial window can be used to study the neural dynamics between cells, as well as between different cortical areas by combining multi-electrode electrophysiological recordings, imaging of neuronal activity (e.g., intrinsic or extrinsic imaging), and optogenetic stimulation. Additionally, this large craniotomy also exposes a large area of cortical blood vessels, allowing for direct manipulation of the lateral cortical vasculature.

This protocol presents a method for creating a large unilateral craniotomy over the temporal and parietal regions of the mouse cerebral cortex. This is especially useful for real time imaging over an expansive area of a cortical hemisphere.

Un grand latéral Craniotomie Procédure pour Mesoscale grand champ d&#39;imagerie optique de l&#39;activité cérébrale

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, ...

Live Imaging of the Ependymal Cilia in the Lateral Ventricles of the Mouse Brain

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various neurological deficits. The current ex vivo live imaging of motile ependymal cilia technique allows for a detailed study of ciliary dynamics following several steps. These steps include: mice euthanasia with carbon dioxide according to protocols of The University of Toledo&#8217;s Institutional Animal Care and Use Committee (IACUC); craniectomy followed by brain removal and sagittal brain dissection with a vibratome or sharp blade to obtain very thin sections through the brain lateral ventricles, where the ependymal cilia can be visualized. Incubation of the brain&#8217;s slices in a customized glass-bottom plate containing Dulbecco&#8217;s Modified Eagle&#8217;s Medium (DMEM)/High-Glucose at 37 &#176;C in the presence of 95%/5% O2/CO2 mixture is essential to keep the tissue alive during the experiment. A video of the cilia beating is then recorded using a high-resolution differential interference contrast microscope. The video is then analyzed frame by frame to calculate the ciliary beating frequency. This allows distinct classification of the ependymal cells into three categories or types based on their ciliary beating frequency and angle. Furthermore, this technique allows the use of high-speed fluorescence imaging analysis to characterize the unique intracellular calcium oscillation properties of ependymal cells as well as the effect of pharmacological agents on the calcium oscillations and the ciliary beating frequency. In addition, this technique is suitable for immunofluorescence imaging for ciliary structure and ciliary protein localization studies. This is particularly important in disease diagnosis and phenotype studies. The main limitation of the technique is attributed to the decrease in live motile cilia movement as the brain tissue starts to die.

Using high-resolution differential interference contrast (DIC) microscopy, an ex vivo observation of the beating of motile ependymal cilia located within the mouse brain ventricles is demonstrated by live-imaging. The technique allows a recording of the unique ciliary beating frequency and beating angle as well as their intracellular calcium oscillation pacing properties.

Imagerie en direct de la épendymaire Cilia dans les ventricules latéraux du cerveau de souris

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various ...

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It has helped to describe the different steps of radial migration and characterize the molecular mechanisms controlling this process. To directly and dynamically analyze migrating neurons they have to be traced over time. This protocol describes a workflow that combines in utero electroporation with organotypic slice culture and time-lapse confocal imaging, which allows for a direct examination and dynamic analysis of radially migrating cortical neurons. Furthermore, detailed characterization of migrating neurons, such as migration speed, speed profiles, as well as radial orientation changes, is possible. The method can easily be adapted to perform functional analyses of genes of interest in radially migrating cortical neurons by loss and gain of function as well as rescue experiments. Time-lapse imaging of migrating neurons is a state-of-the-art technique that once established is a potent tool to study the development of the cerebral cortex in mouse models of neuronal migration disorders.

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

This protocol provides instructions for direct observation of radially migrating cortical neurons. In utero electroporation, organotypic slice culture, and time-lapse confocal imaging are combined to directly and dynamically study the effects of overexpression or downregulation of genes of interest in migrating neurons and to analyze their differentiation during development.

L&#39;imagerie confocale temporelle des neurones migrateurs dans la culture de tranches organotypiques du cerveau embryonnaire de souris en utilisant<em&gt; Dans Utero</em&gt; Electroporation

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Enregistrement de la plasticité synaptique dans des tranches d’hippocampe aiguës maintenues dans un petit volume recyclage - Perfusion- et système de chambre de Submersion-type

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...