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.

Tutte le procedure sono state approvate dalla cura e l&#39;uso degli animali Comitato Istituzionale presso la University of Illinois. Tutti gli animali sono stati alloggiati in strutture di cura degli animali approvato dalla Associazione Americana per l&#39;accreditamento di cura degli animali da laboratorio. Ogni tentativo è stato fatto per ridurre al minimo il numero di animali utilizzati e di ridurre la sofferenza in tutte le fasi dello studio. 1. La preparazione e la rimozione di Brain Mouse affettare <ol><li> Prepararsi per la perfusione e la fetta di incubazione. <ol><li> Circa 30 minuti prima di tagliare, preparare ad alta soluzione di taglio di saccarosio e basso aCSF calcio per l&#39;incubazione della fetta prima della registrazione o l&#39;imaging. </li><li> Disporre tutti gli strumenti necessari (grandi forbici spuntate finali, piccole forbici primavera, pinze anatomiche, grandi forbici o ghigliottina, iride forbici, pinze, gioiellerie 10 ml siringa con ago 27G x 1/2 pollice, piccolo pezzo di 1 cm di filtro x 2 cm carta, e un rotto punta pipetta Pasteur per sltrasferimento ghiaccio) per la perfusione e affettare, e preparare vassoio di perfusione. </li><li> Istituito incubazione da camera con ossigenato aCSF in un bagno d&#39;acqua C 32 o e un piatto di cultura piena di soluzione di taglio. </li></ol></li><li> Preparare fase di taglio per affettare. <ol><li> Tagliare un piccolo pezzo di 3% agar circa 1 cm 3 da utilizzare come arresto per il cervello e 1.5 cm x 1.5 mm x 3 mm per un urto da utilizzare per sostenere l&#39;IC. </li><li> Colla l&#39;arresto sul palco con cianoacrilato nonché l&#39;urto sul lato destro del blocco d&#39;inversione tale da formare un angolo di 80 °. </li></ol></li><li> Rimuovere il cervello. <ol><li> Profondamente anestetizzare un p12-P20 (idealmente p14-18) mouse con ketamina 100 mg / kg e xylaxine 3 mg / kg (o comitato etico comparabili procedura approvata). Nota: Verificare il corretto livello di anestesia con mancata risposta alla pizzico piedi. Poiché l&#39;animale è sotto anestesia brevemente, pomata oftalmica è inutile. </li><li> Utilizzando smussatoterminare forbici, esporre la gabbia toracica dal processo xifoideo al collo. <ol><li> Trovare il processo xifoideo, e fare un taglio orizzontale di circa 1,5 centimetri. </li><li> Fare due tagli verticali dalle estremità del precedente taglio orizzontale alle spalle, circa 1,5 cm ciascuno. </li></ol></li><li> Tagliare attraverso il diaframma e le giunzioni costocondrali per esporre il cuore. </li><li> Con piccole forbici primavera, fare un piccolo, 2-5 tagliare nell&#39;atrio destro mm, dal lato dorsale ventrale del cuore. </li><li> Usando una siringa da 10 ml con un ago 27G x 1/2 pollice, iniettare il ventricolo sinistro e profumato rapidamente l&#39;animale con alta soluzione di taglio saccarosio. </li><li> Una volta che il sangue scorre chiaro, usare le forbici più grandi per rimuovere la testa. Nota: Non abbiamo sistematicamente valutato l&#39;utilità di perfusione. Tuttavia, nella nostra esperienza, transcardiac perfusione nei topi giovani questo può essere fatto con quasi il 100% di successo, e che hanno il vantaggio di eliminare bl rossoOOD cellule, che possono interferire con fluorescenza e di imaging. </li><li> Tagliare la pelle verso il basso la linea mediana per esporre il cranio. </li><li> Utilizzando iris piegate forbici, tagliare il cranio in mezzo agli occhi, poi a partire dal taglio tra gli occhi, accuratamente tagliato da anteriore a posteriore lungo la linea mediana di sutura facendo attenzione a non danneggiare il cervello sotto. Nota: La dura di solito viene fuori con il cranio. Se necessario, rimuovere la dura prima di rimuovere accuratamente il cervello. </li><li> Utilizzando gioiellerie pinze, leva aprire il cranio e rimuovere con attenzione il cervello, quindi inserire il cervello in soluzione di taglio. Fare attenzione a non danneggiare le cortecce. </li></ol></li></ol> 2. Preparazione del cervello per affettare <ol><li> Preparazione del cervello per il montaggio su vibratome palco. <ol><li> Utilizzando una diapositiva segnato con due linee a 90 ° e una linea diagonale a 17 o dalla parte superiore sinistra a destra in basso, si intersecano in cui le due linee si incontrano, posizionare il lato dorsale del cervello fino, utilizzandouna lama di rasoio, rimuovere 2-4 mm del cervello fine rostrale creando una superficie piana. </li><li> Posizionare il lato caudale del cervello sulla superficie piatta di nuova creazione, e allineare la superficie dorsale del cervello con l&#39;orizzontale e la linea mediana del cervello con la linea verticale. </li><li> Allineare la lama di rasoio con la linea 17 o, inclinare il rasoio in un angolo di 30 ° e di rimuovere circa 3 mm della corteccia destra in un doppio taglio diagonale (17 o dal piano orizzontale e 60 o dalla coronale). </li></ol></li><li> Cervello Montaggio su vibratome palco. <ol><li> Mettere un piccolo pezzo di carta da filtro 1 cm x 2 cm sul lato ventrale del cervello in modo che la dimensione lunga è perpendicolare alla linea mediana. </li><li> Applicare delicatamente una piccola quantità di adesivo cianoacrilato alla zona di fronte al blocco d&#39;inversione (ea sinistra del urto). </li><li> Posizionare la doppia faccia del cervello in diagonale taglio al colla in modo che la parte caudale del cervello e romboencefalo sono propped sulla urto e il lato destro del cervello è contro l&#39;arresto. </li><li> Delicatamente premere sul cervello, garantendo che l&#39;intera superficie è a contatto con la camera di affettatura. </li></ol></li></ol> 3. Ottenere la Colliculo-thalamocortical Slice <ol><li> Prendere rapidamente la fase di taglio e posto nel vibratome, riempire il palco con la soluzione di taglio. </li><li> Allineare la lama con la parte superiore (lato ventrale) del cervello. </li><li> Rimuovere 1-1,5 mm dalla parte superiore del cervello, rimuovere 300-500 micron fette e valutare la profondità dopo la rimozione. Nota: Quando l&#39;IC, il MGB, e il nucleo genicolato laterale (LGN) sono tutte visibili, e il giro dentato costituisce una forma di &#39;C&#39; prendere uno a due 600 micron fette. Vedere la Figura 3A. </li><li> Mettere le fette in camera riscaldata (32 ° C) che tiene. </li></ol> 4. Imaging del Slice <ol><li> Preparazione per l&#39;imaging flavoproteina autofluorescenza. <ol> &lt;li&gt; Preparare aCSF per la perfusione di fetta in una camera di registrazione elettrofisiologica standard bolla aCSF con il 95% O 2/5% di CO 2. </li><li> Tirare elettrodo di vetro per la stimolazione. Si noti che più elettrodi stimolanti diversi e configurazioni (vetro, tungsteno, fibra di carbonio, monopolare, bipolare) tutto il lavoro molto bene in congiunzione con l&#39;imaging flavoproteina autofluorescenza. </li><li> Accendere il computer, la macchina fotografica, micromanipolatore, fonte di luce, software stimolazione, software per immagini. </li><li> Profumato camera di registrazione con ossigenato aCSF, posizionare il ponte personalizzato (disegno disponibile in Materiali Supplementari) per elevare fetta nella camera. Nota: tasso di perfusione può variare con qualsiasi individuo istituito; 5-15 ml / min è usato qui. </li><li> Mettere fetta nella camera, abbassare il livello del liquido nella camera per evitare che la fetta da sollevare dal ponte mentre si posiziona arpa appositamente costruito su fetta, avendo cura di evitare di mettere corde dell&#39;arpa su percorsi di interrelazioneest (Figura 1A). Aumentare il livello del liquido a circa 1-2 mm al di sopra fetta di mantenere il flusso aCSF sopra e sotto la fetta. Nota: se riposizionamento è necessario, utilizzare l&#39;estremità pipetta Pasteur rotto o consentire il livello aCSF a salire e utilizzare pinze con estrema cura. </li><li> Inserire l&#39;elettrodo cloruro d&#39;argento in elettrodo di vetro riempita con aCSF, inserire l&#39;elettrodo di vetro in micromanipolatore e connettersi allo stimolo isolante, e posizionare con cura elettrodo di vetro in IC / sostanza bianca che porta al talamo (vedere Figura 1A). </li></ol></li><li> Flavoprotein autofluorescenza acquisizione delle immagini. <ol><li> Raccogliere immagini a 4 Hz (utilizzando un 2X macro obiettivo infinito corretta (NA 0,13)) e una macchina fotografica per 105 secondi mentre elettricamente stimolante (ogni stimolazione è composto da 1 sec di 40 Hz 2 msec impulsi tessuto a 0,05 Hz cinque volte), mentre l&#39;illuminazione il tessuto con luce blu 470-490 nm e catturare sopra 515 nm. Assicurarsi che l&#39;immagine non è né dio scuro, né spente vedi figura 3 colonna sinistra per riferimento. Nota: il tempo Collection, frequenza di raccolta, e la frequenza di stimolazione possono essere modificate per soddisfare l&#39;esperimento, il programma personalizzato incluso nel materiale supplementare può essere modificata in quanto tale. </li><li> Esportare immagini da Matlab, e utilizzando il programma scritto personalizzato disponibile in materiali supplementari per analizzare spettrale di potenza delle immagini a 0,05 Hz. L&#39;immagine risultante mostrerà percorsi collegati. Nota: Il programma personalizzato utilizza una trasformata di Fourier dei valori dei pixel nella serie storica per produrre l&#39;immagine. </li></ol></li><li> Preparazione per l&#39;imaging del calcio. <ol><li> Preparare piccola camera di incubazione con un piatto di coltura 3 centimetri e la membrana cultura sollevato per consentire aCSF di raggiungere sia la parte superiore e inferiore della fetta, e posto su una piastra elettrica per mantenere la temperatura a circa 32 ° C. </li><li> Riempire piccola camera di incubazione con aCSF caldo e lentamente la bolla con il 95% O2/5% di CO 2. </li><li> Mescolare 2 ml di acido Pluronic F-127 e 50 microgrammi di Fura-02:00 sciolto in 48 ml di DMSO. </li><li> Luogo fetta piccola camera di incubazione sulla membrana rialzato e con cura utilizzando una micropipetta aggiungere miscela colorazione direttamente sopra la MGB (o altra struttura di interesse). </li><li> Coprite le fette per prevenire lo sbiancamento prima di imaging, e permettono fette di incubare per 45 minuti - 1 ora. </li><li> Rimuovere le fette della camera di tenuta riscaldato per 10-15 minuti per lavare via il materiale in eccesso. </li><li> Seguire i passaggi da 4.1.1 a 4.1.6 per la preparazione di stimolare e immagine la fetta colliculo-talamo-corticale. </li></ol></li><li> Calcium acquisizione dell&#39;immagine. <ol><li> Raccogliere le immagini a 10 Hz (con un obiettivo ad immersione 20X acqua) per 25 sec stimolando elettricamente (ogni stimolazione è costituito da un impulso 2 msec) tessuto a 0,2 Hz cinque volte mentre illumina il tessuto con 365 luce nm e catturare la fluorescenza sopra 510 nm. </li&gt; <li> Esportare immagini da Matlab e utilizzando il programma scritto personalizzato disponibile in materiali supplementari per analizzare spettrale di potenza delle immagini a 0.2 Hz. L&#39;immagine risultante mostrerà cellule attive. </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...

Nel sistema uditivo, anche se vi è sostanziale elaborazione di informazioni tra la periferia sensoriale e collicolo inferiore, vi è una notevole ulteriore elaborazione prima che raggiunga la corteccia uditiva. Sappiamo molto poco su come che l&#39;elaborazione è fatto e quindi poco su come queste trasformazioni permette al cervello di interpretare le informazioni sensoriali in entrata. Con l&#39;eccezione dell&#39;olfatto, ognuno dei sensi ha un&#39;organizzazione molto simile con segnali periferici inizialmente inoltrato ad alta fedeltà che diminuisce all&#39;aumentare del segnale di sale alla corteccia. La corteccia invia proiezioni alle strutture inferiori per modulare ulteriormente le informazioni in entrata. Questo complesso sistema è stato studiato in una varietà di modi in vivo così come in un certo numero di preparati in vitro. Nel primo caso, tutte le connessioni sono intatte, consentendo al ricercatore di sondare qualsiasi insieme di collegamenti, controllando l&#39;input sensoriale e misurandouscita in una determinata area. Con questo approccio, non vi è poco o nessun controllo della grande varietà di altri fattori, inclusi altri input sensoriali, eccitazione, e attenzione, dando luogo ad una uscita intensamente complesso. In vitro, fettine cerebrali sono stati tagliati per catturare un singolo set di proiezioni, o due aree cerebrali connesse, che permettono ai ricercatori di stimolare e valutare afferenze o aree cerebrali diverse. Si tratta spesso di fette o talamocorticali o tectothalamic in cui o l&#39;ingresso al talamo o il talamo e la sua uscita alla corteccia sono conservati 2-5. Queste preparazioni consentono un&#39;ampia varietà di manipolazioni farmacologiche, elettrici, e optogenetic. Tuttavia con solo due regioni del cervello, essi sono principalmente valutano il trasferimento di informazioni e non hanno la capacità di valutare la trasformazione delle informazioni che passa attraverso il talamo. Anche la proiezione reticolo-talamo, che possono svolgere un ruolo nella modulazione attenzione 6-9 è prESENT in questa fetta. Qui mostriamo miglioramenti su nostra precedente preparazione 1, che permette il controllo investigatore di vari ingressi al talamo di dare una prospettiva unica di come le porte talamo e filtri informazioni. Abbiamo paio questo romanzo preparazione fetta con immagini flavoproteina autofluorescenza per valutare la connettività fetta e l&#39;analisi di attivazione su larga scala, l&#39;imaging calcio nel talamo per neuronale analisi di popolazione, e la registrazione singola cellula per misurare l&#39;impatto dei vari ingressi su un livello di singola cellula. Per aiutare a mantenere queste connessioni diffuse abbiamo anche sviluppato una serie di modifiche del normale ancoraggio fetta (alias &quot;arpa&quot;) per la tenuta della fetta cervello in luogo e un ponte di elevare la fetta per una migliore perfusione. L&#39;arpa è stato progettato in un ferro di cavallo modificata per circondare la fetta e consentire punti di fissaggio personalizzabili per le corde dell&#39;arpa. Tre stringhe sonoallegata tale che i) uno si trova orizzontalmente lungo il bordo mediale della fetta, ii) uno estende dal bordo caudale del IC all&#39;estremità caudale della CA e iii) uno estende diagonalmente dal bordo mediale della fetta un&#39;area rostrale alla CA (vedere Figura 1A). Piccoli tacche del telaio per incollaggio (con colla cianoacrilato) dell&#39;arpa stringhe permettono una ridotta quantità di pressione sulla fetta per aiutare a mantenere l&#39;integrità fetta (vedere Figura 1B). Utilizzando stampa tridimensionale, possiamo arpe progettazione personalizzata alle nostre specifiche uniche, così come i ponti che consentono il flusso ideale di liquido cerebrospinale artificiale (aCSF) sopra e sotto il tessuto. Questo mantiene anche ampie aree per la luce di penetrare il tessuto per la patch clamp elettrofisiologia.

<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 esempio della fetta cervello di topo colliculo-talamocorticale ottenuta nel topo P15 è mostrata in figura 2. La fetta ideale conterrà i quattro principali strutture uditive mesencefalo e prosencefalo IC, MGB, TRN, e AC, che sono tutti attivati ​​quando il circuito integrato è stimolata (Figura 2A). Utilizzando l&#39;analisi di Fourier, la potenza spettrale viene misurato alla frequenza di stimolazione elettrica, con regioni cerebrali collegate mostrano attivit...

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;utilizzo di più angoli per tagliare il cervello di topo cucciolo, miglioriamo su una fetta cerebrale acuto precedentemente descritto, che cattura le connessioni tra le maggiori strutture mesencefalo e proencefalo uditive.

Modifica di un Colliculo-talamo-corticale mouse Cervello Fetta, Incorporando stampa 3-D della Camera componenti e multi-scala 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 ...

modification-colliculo-thalamocortical-mouse-brain-slice

Research

JoVE Journal

Neuroscience

8.2K Views.  University of Illinois Urbana-Champaign.  L'utilizzo di più angoli per tagliare il cervello di topo cucciolo, miglioriamo su una fetta cerebrale acuto precedentemente descritto, che cattura le connessioni tra le maggiori strutture mesencefalo e proencefalo uditive. 

Video: 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.

Gli array scalabile fluidico Iniettore per Viral targeting di intatto 3-D circuiti cerebrali

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

Ricerca

Neuroscienze

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-topo Imaging neuroanatomici Risonanza Magnetica

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.

Imaging Pheromone Sensing in preparazione dei tessuti Vomeronasal acuta Slice mouse

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.

Una procedura di impianto del Camera spinale per longitudinale<em&gt; In Vivo</em&gt; Imaging del cavo del mouse spinale

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-mount Imaging di embrione di topo Sensory Axon Proiezioni

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.

3-D imaging e analisi di neuroni infetti<em&gt; In Vivo</em&gt; Con<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.

Una procedura Grande laterale Craniotomia per Mesoscale Wide-field Optical Imaging di 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, ...

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.

Immagini dal vivo del ependimale Cilia nei ventricoli laterali del cervello di topo

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.

Impressione confocale dei migratori dei neuroni nella cultura di fetta organotipica del cervello embrionale del mouse usando<em&gt; In Utero</em&gt; Elettroporazione

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.

Registrazione di plasticità sinaptica in fettine ippocampali Acute mantenuti in un piccolo volume riciclaggio - aspersione- e sistema di immersione-tipo camera

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