Name: imaging serotonergic fibers in the mouse spinal cord using the clarity/cubic technique
Uploaded: 2016-02-26T14:00:00.000+00:00
Duration: 9 min 54 s
Description: Watch this Scientific Journal Video about Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique at JoVE.com

This work was supported by the Australian Research Council Centre of Excellence for Integrative Brain Function (ARC Centre Grant CE140100007), an NHMRC project grant (#1086643). Prof. George Paxinos is supported by a Senior Principal Research Fellow NHMRC grant (#1043626).

Etica Dichiarazione: Tutte le procedure che coinvolgono soggetti animali seguono le linee guida del Comitato Animal Care and Ethics (ACEC) presso l&#39;Università del New South Wales (il numero ACEC approvato è di 14 / 94A). 1. Preparazione del cavo del mouse trasparente spinale <ol><li> Preparazione di Ice Cold Hydrogel Soluzione <ol><li> Preparazione della soluzione di paraformaldeide al 16% (PFA) <ol><li> Aggiungere 16 g di polvere di paraformaldeide in 70 ml-pre riscaldato acqua distillata (50-55 ° C) e mescolare su un agitatore magnetico riscaldata fino paraformaldeide si scioglie. Nota: Non lasciare che la soluzione per il riscaldamento oltre 55 ° C ed essere consapevoli del fatto che paraformaldeide è tossico. </li><li> Trasferire la soluzione paraformaldeide ad un cilindro e aggiungere acqua distillata fino a 100 ml. Raffreddare la soluzione paraformaldeide utilizzando un C frigorifero a 4 °. </li></ol></li><li> Sciogliere 125 mg VA-044 iniziatore 26,25 ml di acqua distillata e raffreddare la soluzione in un 4° C frigo. </li><li> Raffreddare 5 ml di soluzione di acrilammide (40%), 1,25 ml di soluzione di bis (2%) (ATTENZIONE: Bis e le soluzioni di acrilamide sono tossici, può causare difetti genetici o il cancro), 5 ml di 10X PBS, 12,5 ml al 16% soluzione PFA, e la soluzione di iniziatore VA-044 su ghiaccio. </li><li> Mescolare le soluzioni sopra sul ghiaccio. Nota: il volume totale è di 50 ml. </li></ol></li><li> Perfusione e Raccolta del cavo del mouse spinale <ol><li> Anestetizzare un mouse con un&#39;iniezione intraperitoneale di ketamina (80 mg / kg) e xyzaline (5 mg / kg) diluito in soluzione fisiologica 0,9% normale. Il dosaggio per mouse può essere inferiore a quello dei ratti. </li><li> Su una superficie di plastica adatta in una cappa aspirante, fissare gli arti del mouse lontano dal corpo (nastro adesivo è efficace) una volta che il mouse non risponde a una presa per la sua pelle del piede. </li><li> Tagliare la pelle del mouse sopra il petto e poi il torace osso per esporre il cuore con una forbice. </li><li> Utilizzando una pompa peristaltica, con un ago 25 G collegato a the estremità del tubo, inserire l&#39;ago nel ventricolo sinistro del cuore, snip l&#39;atrio destro per creare un punto di uscita per il sangue e quindi avviare il lavaggio con 40 ml 0,9% soluzione fisiologica una velocità di circa 10 ml / min. </li><li> Al termine, profumato il mouse con 35 ml di ghiaccio soluzione idrogel freddo. Nota: profumato ad una velocità di 10 ml / min per evitare gonfiore del tessuto nervoso. </li><li> Incidere la pelle sul dorso con una lama e quindi rimuovere i muscoli vicino al vertebrati. Dopo il taglio archi vertebrali da un lato e facendoli ruotare verso l&#39;altro lato, prendere il midollo spinale dalla colonna vertebrale, con una multa forbice. </li></ol></li><li> Cancellazione del cavo del mouse spinale <ol><li> Posizionare il midollo spinale in 15 ml di soluzione di idrogel notte a 4 ° C. </li><li> Rimuovere 10 ml della soluzione di idrogel e versare il resto della soluzione con i segmenti del midollo spinale in una provetta 5 ml. Riempire il tubo 5 ml con la soluzione di idrogel fino a quando non è completamente pieno. </li><li> Stretch un piccolo pezzo di parafilm sopra la parte superiore del tubo 5 ml e poi avvolgere parafilm intorno al collo del tubo. Nota: Assicurarsi che i contatti parafilm la soluzione idrogel e non vi siano bolle tra la soluzione idrogel e il parafilm. </li><li> Inserire il tubo 5 ml a 37 &#160; ° C forno durante la notte. Osservare la soluzione idrogel fino a farla diventare un gel. </li><li> Prendere il tubo 5 ml dal forno e rimuovere il gel dal tubo con una chiave (come un shoveled 1 ml puntale). Rimuovere il gel dal tessuto del midollo spinale da attaccare il tessuto grossolano al gel e poi prendere il tessuto grossolano di distanza (il gel si attacca al tessuto e quindi sarà rimosso dal tessuto). </li><li> Dopo aver rimosso il gel dal tessuto del midollo spinale, lavare il midollo spinale 4 volte con PBS 1x (pH 7,4) per 24 ore su un agitatore. </li><li> Preparare la soluzione di compensazione CUBIC sciogliendo 3,85 g di urea e 3,85 g N, N, N &#39;, N&#39;-tetrakis (2-idrossipropil) ethylenediamine a 5.38 ml di acqua distillata. Nota: Un agitatore caldo viene utilizzato. Aggiungere 2,31 g di polietilene glicole mono-p-isooctylphenyl etere / Triton X-100 alla soluzione una volta che è chiaro e si raffredda a temperatura ambiente. Nota: 10 g di questi tre sostanze chimiche sono per un mouse. </li><li> Tagliare il mouse midollo spinale coronale in segmenti lunghi 2-3 mm con una lametta e metterli in 5 ml di soluzione di compensazione CUBIC sopra. Posizionare su un agitatore in forno a 37 &#160; ° C per 3 giorni. </li><li> Tre giorni dopo, cambiare la soluzione di compensazione per una nuova. Nota: la suddetta soluzione viene preparata prima dell&#39;uso. </li><li> Due o tre giorni più tardi, dopo l&#39;aggiornamento della soluzione di compensazione CUBIC, controllare la trasparenza del tessuto contro una carta con carattere 8 lettere. Se è trasparente, le lettere sono visibili attraverso il tessuto eliminato. Rimuovere la soluzione di compensazione e aggiungere 4 ml di PBST (0,1% Triton-X100) per lavare il tessuto 4 volte al giorno (ogni 6 ore). </li></ol></li></ol><ol><li> Incubare i segmenti del midollo spinale nella soluzione di anticorpo primario (anti-serotonina, cresciuto a coniglio, diluito 1: 100 in PBST) per 3 giorni su un agitatore in forno a 37 °. </li><li> Rimuovere la soluzione di anticorpi e aggiungere 4 ml di PBST (0,1% Triton-X100) per lavare i segmenti del midollo spinale 4 volte al giorno (ogni 6 ore) in un forno a 37 ° C. </li><li> Rimuovere la soluzione PBST e aggiungere la soluzione di anticorpo secondario (594 coniugato capra anti-IgG di coniglio, diluito 1: 100 in PBST) incubare il tessuto per 3 giorni su un agitatore in forno a 37 °. </li><li> Rimuovere la soluzione di anticorpo secondario e aggiungere 4 ml di PBST per lavare i segmenti del midollo spinale 4 volte al giorno (ogni 6 ore) su un agitatore in forno a 37 °. Il giorno successivo il tessuto è pronto per l&#39;imaging. </li></ol> 3. Imaging <ol><li> Rimuovere la soluzione PBST e aggiungere 4 ml di 85% glicerolo per rendere l&#39;indice di rifrazione del tessuto anche. </li><li&gt; Controllare la chiarezza del tessuto. Una volta chiarito in circa 1 ora, mettere il tessuto su una grande, scivolo vetro sottile che si adatta il telaio del multi-photo fluorescente microscopio tenuta. </li><li> Mettere qualche goccia di 85% glicerolo accanto al tessuto del midollo spinale e applicare il coprioggetto con un bicchiere coprioggetto 22 x 50 mm 2. Nota: L&#39;orientamento del midollo spinale decide come l&#39;immagine assomiglia. </li><li> Mettere il vetrino sul telaio del microscopio a fluorescenza multi-photon tenuta e spostare il tessuto nel percorso della luce. </li><li> Scegliere l&#39;elio neon linea laser 594 nm e regolare l&#39;intensità del laser al livello ottimale controllando la luminosità del segnale positivo nell&#39;immagine live. </li><li> Selezionare l&#39;area di scansione del tessuto del midollo spinale con l&#39;obiettivo 20X (in acqua, NA 0,7) e preparare per creare un z-stack, e la profondità di ogni passo impostato a 3 micron. </li><li> Acquisire il tessuto dalla parte superiore alla parte inferiore del z-stack sotto la obiet 20Xtiva e quindi obiettivo 63X (in olio, NA 1.4) (dimensione del passo è stato di 1 micron), rispettivamente. Ricostruire i video 3D utilizzando un software 3D 28. </li></ol>

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The authors have nothing to disclose.

Il protocollo descritto mostra come le fibre immagine serotoninergici nel topo midollo spinale con la chiarezza combinato e tecniche di cubi. Si introduce un processo di compensazione più veloce rispetto al protocollo di compensazione passiva sviluppato da Cheung et al. 14 e Tomer et al. 15 e permette il tessuto del midollo spinale per essere ben supportato dal idrogel durante compensazione. Un passo importante durante la fissazione del mouse midollo spinale, come riportato da Cheung et al. 14 e Tomer et al. 15, è di mantenere tutte le soluzioni raffreddare su ghiaccio, che impedisce la polimerizzazione delle sostanze chimiche. L&#39;altro punto critico è al perfusione il mouse lentamente con la soluzione di idrogel per evitare gonfiore del tessuto nervoso. Per la soluzione di idrogel per diventare un gel, è necessario degasaggio. In alternativa a questo, abbiamo usato parafilm per coprire la parte superiore del tubo e lasciato bolle d&#39;aria tra la hydsoluzione Roghel e il parafilm. Questo ha funzionato molto bene e la soluzione idrogel diventato un gel dopo poche ore. Sebbene alcune bolle sono stati trovati nel gel, essi non interferiscano con i seguenti esperimenti, che è stato dimostrato per l&#39;assenza di bolle nel tessuto del midollo spinale. La compensazione è la fase più critica, e richiede una combinazione di reagenti. La soluzione di compensazione, contenente ammino-alcool, rimuove i lipidi del tessuto del midollo spinale molto più veloce rispetto alla / soluzione compensazione SDS acido borico base utilizzato nel protocollo CHIAREZZA originale 16. Il vantaggio della chiarezza e CUBIC rispetto alle tecniche tradizionali di immunoistochimica è che l&#39;intero blocco tessuto può essere ripreso senza sezionare il tessuto, di conseguenza, la continuità delle fibre serotoninergici viene mantenuta, ed è possibile seguire la stessa fibra per una lunga distanza nel z-stack. Con questo vantaggio, collateralizzazione può essere tranquillamente identificato con le immagini in 3D, unND nuove conclusioni possono quindi essere fatte sulla natura dei percorsi serotoninergici. Il protocollo descritto è uno strumento utile per ricerca di sistemi in fibra, così come nuclei semplicemente etichettatura tessuto con anticorpi specifici. Questa tecnica è anche una scelta ideale per interrogare i sistemi diversi nello stesso cervello di topo mediante etichettatura doppia o tripla. Nel nostro ambiente di laboratorio, un microscopio a fluorescenza multi-photon era disponibile con una distanza di lavoro di circa 300 micron. Questo limita la nostra immagine ad un massimo di 600 micron se sono scansionati entrambi i lati del tessuto. Tuttavia, un microscopio a fluorescenza foglio leggero, può immagine tutto il cervello e il midollo spinale del mouse attraverso la loro larghezza, lunghezza e profondità. L&#39;altro vantaggio di questa tecnica è che gli anticorpi possono essere eluiti e poi un altro anticorpo o anticorpi diversi applicati nuovo al tessuto stesso. Questo riduce l&#39;utilizzo degli animali considerevolmente, così come i costi connessi con la preparazione di nuovo tessuto cerebraleper ogni esperimento. Per lo stesso motivo, questa tecnica può essere applicata ad altri tessuti. Ci sono già stati studi condotti su polmoni, reni, cuore, intestino, e il tessuto tumorale 26,27. Ci sono anche limitazioni a questa tecnica, per esempio, i nuclei del rafe in romboencefalo topo contiene molti tipi di neuroni, tra neuroni GABAergici. Sebbene CLARITY e CUBIC consentiranno lo studio della distribuzione dei diversi tipi di fibre raphespinal in un singolo topo cervello e il midollo spinale. Tali neuroni, che non sono indicate specificamente dai neurotrasmettitori o marcatori proteici, non possono essere identificati in questa impostazione. Un&#39;alternativa è quella di etichettare i neuroni specifici o sistemi in fibra con sonde di acido nucleico come nella ibridazione in situ. Questo è, invece, limitato dal sistema di etichettatura e filtri disponibili per l&#39;imaging fluorescente. Imaging tutte le fibre raphespinal resta dunque una sfida.Se queste tecniche sono stati combinati con una iniezione intracranica di un tracciante anterogrado (come Phaseolus vulgaris leucoagglutinin - PHA-L), potrebbe essere possibile vedere fibre serotoninergici, fibre GABAergici, e altri.

Supraspinal projections are responsible for the modulation of diverse behaviors such as pain perception. One of the projections carrying nociceptive information contains serotoninergic fibers, which originate from the hindbrain raphe and adjacent reticular nuclei1,2. Physiological and pharmacological studies have demonstrated an increased release of serotonin in the dorsal horn of the spinal cord after electrical stimulation of the raphe nuclei in the hindbrain3-5. In the rat and opossum, serotonergic raphespinal fibers have dense terminals, not only in the dorsal horn6-8, but also in the intermediate zone7,9,10, the ventral horn7,11, and even lamina 1012,13. There are no similar studies in the mouse. The present study aimed to map the termination pattern of serotonergic fibers arising from the hindbrain raphe nuclei and their adjacent reticular nuclei in the mouse spinal cord using the recently published CLARITY14 method and its modification - CUBIC15. 
Conventional fluorescence or peroxidase immunohistochemistry of the spinal cord clearly shows the distribution of serotonergic fibers in the gray matter of the spinal cord in 30-40 &#181;m thick cross-sections. However, this approach does not show the continuity of the serotonergic fiber tracts in the white matter and their collaterals in the gray matter. Although the 3D reconstruction of histological sections has advanced our knowledge of fiber tracts, it remains a challenge for histologists and anatomists to follow a single tract due to small distortions in the tissue caused by cutting. To circumvent this obstacle a number of researchers have developed various protocols for making the whole tissue structure transparent, and collecting an image of unaltered tissue in a single video file17-21. So far, the clear, lipid-exchanged, acrylamide-hybridized rigid, imaging/ immunostaining compatible, tissue hydrogel (CLARITY) technique, developed by Deisseroth's group14,15, as well as CUBIC, developed by Susaki et al16 are the most successful. Since the publication of the protocols, many researchers have started using these techniques to investigate various aspects of biological tissues, including, not only the brain22-25, but also the heart, kidneys, intestine, and the lungs26,27. 
By fixing the mouse spinal cord with the hydrogel solution (CLARITY) and clearing with the CUBIC reagents (which is a much faster method than that described by the original CLARITY protocol14,15), a spinal cord tissue block of 2-3 mm long was cleared within two weeks and immunofluorescence staining for serotonin completed in eight days. With just a combination of chemical agents, conventional immunohistochemistry can be used to create an image of individual fiber tracts in a 3D video file in approximately one month.

<table><tbody><tr><td>Photoinitiator VA044</td><td>Wako</td><td>va-044/225-02111</td><td>http://www.wako-chem.co.jp/specialty/waterazo/VA-044.htm</td></tr><tr><td>40% acrylamide solution</td><td>Bio Rad</td><td>161-0140</td><td>http://www.bio-rad.com/en-au/sku/161-0140-40-acrylamide-solution</td></tr><tr><td>2% Bis Solution</td><td>Bio Rad</td><td>161-0142</td><td>http://www.bio-rad.com/en-au/sku/161-0142-2-bis-solution?parentCategoryGUID=5e7a4f31-&lt;br/&gt;879c-4d63-ba0b-82556a0ccf1d</td></tr><tr><td>paraformaldehyde</td><td>Sigma</td><td>158127</td><td>http://www.sigmaaldrich.com/catalog/product/sial/158127?lang=en&amp;amp;region=AU</td></tr><tr><td>urea</td><td>Merck Millipore</td><td>66612</td><td>http://www.merckmillipore.com/AU/en/product/Urea---CAS-57-13-6---Calbiochem,EMD_BIO-66612</td></tr><tr><td>N,N,N&amp;rsquo;,N&amp;rsquo;-tetrakis (2-hydroxypropyl) ethylenediamine</td><td>Merck Millipore</td><td>821940</td><td>http://www.merckmillipore.com/AU/en/product/Ethylenediamine-N,N,N&#039;,N&#039;-tetra-2-propanol,MDA_CHEM-821940</td></tr><tr><td>Triton-X 100</td><td>Merck Millipore</td><td>648462</td><td>http://www.merckmillipore.com/AU/en/product/TRITON&amp;reg;-X-100-Detergent---CAS-9002-93-1---Calbiochem,EMD_BIO-648462</td></tr><tr><td>sucrose</td><td>Sigma</td><td>S0389</td><td>http://www.sigmaaldrich.com/catalog/product/sigma/s0389?lang=en&amp;amp;region=AU</td></tr><tr><td>serotonin antibody</td><td>Merck Millipore</td><td>AB938</td><td>http://www.merckmillipore.com/AU/en/product/Anti-Serotonin-Antibody,MM_NF-AB938</td></tr><tr><td>goat anti rabbit IgG (H+L) Secondary Antibody, Alexa Fluor&amp;reg; 594 conjugate</td><td>Life Technologies</td><td>&amp;nbsp;A-11012</td><td>https://www.lifetechnologies.com/order/genome-database/antibody/Rabbit-IgG-H-L-Secondary-Antibody-Polyclonal/A-11012</td></tr><tr><td>multi-photon microscope</td><td>Leica</td><td>Leica TCS SP5 MP STED</td><td>http://www.leica-microsystems.com/products/confocal-microscopes/details/product/leica-tcs-sp5-mp/</td></tr></tbody></table>

imaging serotonergic fibers in the mouse spinal cord using the clarity/cubic technique

fibre lunghe che scendono verso il midollo spinale sono essenziali per la locomozione, percezione del dolore, e altri comportamenti. Il modello terminazione in fibra nel midollo spinale della maggior parte di questi sistemi in fibra non sono state esaminate a fondo in qualsiasi specie. fibre serotoninergici, che proiettano al midollo spinale, sono stati studiati nei ratti e opossum su sezioni istologiche e il loro significato funzionale è stata dedotta in base al loro modello di terminazione in fibra nel midollo spinale. Con lo sviluppo della chiarezza e tecniche cubica, è possibile studiare questo sistema di fibre e la sua distribuzione nel midollo spinale, che rischia di rivelare caratteristiche precedentemente sconosciuti di percorsi supraspinal serotoninergici. Qui, forniamo un protocollo dettagliato per l&#39;imaging delle fibre serotoninergici nel topo midollo spinale utilizzando la nitidezza, associato e le tecniche cubi. Il metodo prevede la perfusione di un mouse con una soluzione idrogel e chiarificazione del tessuto con una combinzione di compensazione reagenti. tessuto del midollo spinale è stato eliminato in poco meno di due settimane, e la conseguente colorazione immunofluorescenza contro la serotonina è stata completata in meno di dieci giorni. Con un microscopio a fluorescenza multi-fotone, il tessuto è stato digitalizzato e l&#39;immagine 3D è stata ricostruita utilizzando il software OsiriX.

Questa sezione mostra i risultati di anticorpi serotonina colorazione nel topo trasparente midollo spinale utilizzando una combinazione di chiarezza e protocolli cubi. Abbiamo dimostrato che le fibre serotoninergici sono presenti in tutti lamine del midollo spinale con una predominanza nella porzione ventrale del corno ventrale (Figura 1, vedi anche Video 1). Il tessuto di controllo non ha avuto fibre positive (risultato non è stato mostrato). Nel corno ventrale, fibre serotoninergici densamente sono presenti nella porzione ventromediale del corno ventrale e si estendono verso la porzione laterale del corno ventrale (Figura 2, vedi anche Video 2). Alcune fibre immunopositive estendono anche dal corno ventrale verso il corno dorsale o il canale centrale. fibre immunopositive sono presenti in tutti lamine del corno dorsale, ma c&#39;è un piccolo spazio tra le fibre positivi in ​​lamina 2 e 4, soprattutto nel lateraleparte del corno dorsale. La maggior parte delle fibre serotoninergici del corno dorsale sono di piccolo diametro e solo un piccolo numero di essi sono di grande diametro (Video 1). In una sezione orizzontale, sono state osservate fibre serotoninergici densamente nel corno ventrale di viaggiare lungo l&#39;asse longitudinale del midollo spinale e formare un fascio di fibre. La scoperta più interessante è che questo fascio di fibre emette rami regolarmente lungo l&#39;asse longitudinale, che sono perpendicolari al fascio di fibre. Questi rami ulteriormente collateralizzare lungo i loro percorsi alla porzione laterale del corno ventrale. Rispetto a questi rami, quelli che si estendono verso la linea mediana sono irregolari e piccole (Figura 2). Sotto l&#39;obiettivo 63X, sono state osservate le fibre serotoninergici del corno dorsale di viaggiare lungo l&#39;asse del midollo spinale e terminare in vari punti lungo il percorso del tratto. Le fibre spesse sono sporadicamente distribuiti tra la F sottile<html> IBERS (Figura 3, vedi anche Video 3). <img alt="Figura 1" src="/files/ftp_upload/53673/53673fig1.jpg" /> Figura 1:. Fibre serotoninergici in una sezione coronale del cavo lombare 200X ingrandimenti sinistra è corno ventrale destro è corno dorsale, dorsale è la parte mediale del midollo spinale, e ventrale è la parte laterale del midollo spinale. La barra della scala è di 100 micron. fibre serotoninergici sono presenti in tutti lamine, soprattutto nella parte ventromediale del corno ventrale dove la densità delle fibre è alto. Queste fibre si estendono sia verso la parte laterale del corno ventrale e il canale o dorsale centrale corno. La densità di fibre in altre lamine è basso. La maggior parte delle fibre serotoninergici sono sottili. Solo un piccolo numero di fibre sono spesse e sono considerati sia nel corno dorsale e il corno ventrale (&gt; vedi anche Video 1 (Tasto destro del mouse per scaricare)). <img alt="figura 2" src="/files/ftp_upload/53673/53673fig2.jpg" /> Figura 2:. Fibre serotoninergici nella parte ventrale del corno ventrale a 200X di ingrandimento di sinistra e destra sono i lati laterali del midollo spinale, dorsale è la parte rostrale del midollo spinale, e ventrale è la parte caudale del midollo spinale. La barra della scala è di 100 micron. fibre serotoninergici sono mostrati viaggiando lungo l&#39;asse longitudinale del midollo spinale e confezionati nella porzione mediale del corno ventrale dove formano un fascio di fibre di spessore. Su questo fascio, alcuni rami sono estesi a intervalli regolari verso la porzione laterale del corno ventrale, mentre altri che si estendono verso la linea mediana sono irregolari e più brevi. Queste fibre terminano in molti punti lungo il percorso del tratto ( <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/53673/Video2.mp4">vedi anche Video 2</a>(Tasto destro del mouse per scaricare)). <img alt="Figura 3" src="/files/ftp_upload/53673/53673fig3.jpg" /> Figura 3: fibre serotoninergici nel corno dorsale sotto a 630X di ingrandimento di sinistra è la parte laterale del midollo spinale, a destra è la parte mediale del midollo spinale, dorsale e ventrale, fare riferimento alla dorsale e parte ventrale del corno dorsale, rispettivamente. . La barra della scala è di 7 micron. fibre serotoninergici viaggiano lungo l&#39;asse longitudinale del midollo spinale e terminano lungo il percorso del tratto. Queste fibre sono principalmente di piccolo diametro con un piccolo numero di fibre spesse mescolati. Queste fibre raramente si sovrappongono tra di loro ( <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/53673/Video3.mp4">vedi anche Video 3 (clic destro per il download)</a> ).

Watch this Scientific Journal Video about Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique at JoVE.com

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Imaging Fibre serotoninergici nel midollo spinale mouse Uso del CHIAREZZA / Tecnica CUBIC

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

imaging-serotonergic-fibers-mouse-spinal-cord-using-claritycubic

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

10.4K Views.  Neuroscience Research Australia. Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Video: Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

Imaging in vivo del midollo spinale di topo Utilizzo di due fotoni Microscopia

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

Ricerca

Neuroscienze

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 ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Due fotoni Imaging di Cellular Dynamics nel midollo spinale del mouse

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Clearing ottica del sistema nervoso centrale del mouse Utilizzando CHIAREZZA passivo

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...

Immunostaining of Whole-Mount Retinas with the CLARITY Tissue Clearing Method

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining and imaging of thick brain samples. However, this advanced technology has not yet been used for whole-mount retinas. Although the retina is partially transparent, its thickness of approximately 200 &#181;m (in mice) still limits the penetration of antibodies into the deep tissue as well as reducing light penetration for high-resolution imaging. Here, we adapted the CLARITY method for whole-mount mouse retinas by polymerizing them with an acrylamide monomer to form a nanoporous hydrogel and then clearing them in sodium dodecyl sulfate to minimize protein loss and avoid tissue damage. CLARITY-processed retinas were immunostained with antibodies for retinal neurons, glial cells, and synaptic proteins, mounted in a refractive index matching solution, and imaged. Our data demonstrate that CLARITY&#160;can improve the quality of standard immunohistochemical staining and imaging for retinal neurons and glial cells in whole-mount preparation. For instance, 3D resolution of fine axon-like and dendritic structures of dopaminergic amacrine cells were much improved by CLARITY. Compared to non-processed whole-mount retinas, CLARITY can reveal immunostaining for synaptic proteins such as postsynaptic density protein 95. Our results show that CLARITY renders the retina more optically transparent after the removal of lipids and preserves fine structures of retinal neurons and their proteins, which can be routinely used for obtaining high-resolution imaging of retinal neurons and their subcellular structures in whole-mount preparation.

Here we present a protocol to adapt the CLARITY method of the brain tissues for whole-mount retinas to improve the quality of standard immunohistochemical staining and high-resolution imaging of retinal neurons and their subcellular structures.

Immunostaining delle retine a montaggio intero con il metodo clarity tissue clearing

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining ...

Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the morphology of processes and substructures within these neurons. As such, much of what is known about brain function has arisen alongside developments in imaging technologies that allow further insight into how neurons are organized and connected in the brain. Improvements in tissue clearing have allowed for high-resolution imaging of thick brain slices, facilitating morphological reconstruction and analyses of neuronal substructures, such as dendritic arbors and spines. In tandem, advances in image processing software provide methods of quickly analyzing large imaging datasets. This work presents a relatively rapid method of processing, visualizing, and analyzing thick slices of labeled neural tissue at high-resolution using CLARITY tissue clearing, confocal microscopy, and image analysis. This protocol will facilitate efforts toward understanding the connectivity patterns and neuronal morphologies that characterize healthy brains, and the changes in these characteristics that arise in diseased brain states.

Neuronal dendritic morphology often underlies function. Indeed, many disease processes that affect the development of neurons manifest with a morphological phenotype. This protocol describes a simple and powerful method for analyzing intact dendritic arbors and their associated spines.

Imaging e quantificazione di dendriti neuronali intatti tramite CLARITY Tissue Clearing

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the ...

A Tissue Clearing Method for Neuronal Imaging from Mesoscopic to Microscopic Scales

A detailed protocol is provided here to visualize neuronal structures from mesoscopic to microscopic levels in brain tissues. Neuronal structures ranging from neural circuits to subcellular neuronal structures are visualized in mouse brain slices optically cleared with ScaleSF. This clearing method is a modified version of ScaleS and is a hydrophilic tissue clearing method for tissue slices that achieves potent clearing capability as well as a high-level of preservation of fluorescence signals and structural integrity. A customizable three dimensional (3D)-printed imaging chamber is designed for reliable mounting of cleared brain tissues. Mouse brains injected with an adeno-associated virus vector carrying enhanced green fluorescent protein gene were fixed with 4% paraformaldehyde and cut into slices of 1-mm thickness with a vibrating tissue slicer. The brain slices were cleared by following the clearing protocol, which include sequential incubations in three solutions, namely, ScaleS0 solution, phosphate buffer saline (&#8211;), and ScaleS4 solution, for a total of 10.5&#8211;14.5 h. The cleared brain slices were mounted on the imaging chamber and embedded in 1.5% agarose gel dissolved in ScaleS4D25(0) solution. The 3D image acquisition of the slices was carried out using a confocal laser scanning microscope equipped with a multi-immersion objective lens of a long working distance. Beginning with mesoscopic neuronal imaging, we succeeded in visualizing fine subcellular neuronal structures, such as dendritic spines and axonal boutons, in the optically cleared brain slices. This protocol would facilitate understanding of neuronal structures from circuit to subcellular component scales.

The protocol provides a detailed method of neuronal imaging in brain slice using a tissue clearing method, ScaleSF. The protocol includes brain tissue preparation, tissue clarification, handling of cleared slices and confocal laser scanning microscopy imaging of neuronal structures from mesoscopic to microscopic levels.

Un metodo di compensazione dei tessuti per l'imaging neuronale dalle scale mesoscopiche a quelle microscopiche

A detailed protocol is provided here to visualize neuronal structures from mesoscopic to microscopic levels in brain tissues. Neuronal structures ranging ...

Establishing a Mouse Contusion Spinal Cord Injury Model Based on a Minimally Invasive Technique

Using minimally invasive methods to model spinal cord injury (SCI) can minimize behavioral and histological differences between experimental animals, thereby improving the reproducibility of the experiments.
These methods need two requirements to be fulfilled: clarity of the surgical anatomical pathway and simplicity and convenience of the laboratory device. Crucially for the operator, a clear anatomical pathway provides minimally invasive exposure, which avoids additional damage to the experimental animal during the surgical procedures and allows the animal to maintain a consistent and stable anatomical morphology during the experiment.
In this study, the use of a novel integrated platform called the SCI coaxial platform for spinal cord injury in small animals to expose the T9 level spinal cord in a minimally invasive way and stabilize and immobilize the vertebra of mice using a vertebral stabilizer is researched, and, finally, a coaxial gravity impactor is used to contuse the spinal cord of mice to approach different degrees of T9 spinal cord injury. Finally, histological results are provided as a reference for the readers.

Minimally invasive techniques and a simple laboratory device improve the reproducibility of the spinal cord injury model by reducing operative damage to the experimental animals and allowing anatomical morphology maintenance. The method is worthwhile because the reliable results and reproducible procedure facilitate investigations of the mechanisms of disease reparation.

Stabilire un modello di lesione del midollo spinale per contusione murina basato su una tecnica minimamente invasiva

Using minimally invasive methods to model spinal cord injury (SCI) can minimize behavioral and histological differences between experimental animals, ...

Establishment of Central Cord Syndrome Model in C57BL/6J Mouse

Animal models of central cord syndrome (CCS) could substantially benefit preclinical research. Identifiable anatomical pathways can give minimally invasive exposure approaches and reduce extra injury to experimental animals during operation, enabling the maintenance of consistent and stable anatomical morphology during experiments to minimize behavioral and histological differences between individuals to improve the reproducibility of experiments. In this study, the C6 level spinal cord was exposed using a spinal cord injury coaxial platform (SCICP) and combination with a minimally invasive technique. With the assistance of a vertebral stabilizator, we fixed the vertebrae and compressed the spinal cords of C57BL/6J mice with 5 g/mm2 and 10 g/mm2 weights with SCICP to induce different degrees of C6 spinal cord injury. In line with the previous description of CCS, the results reveal that the lesion in this model is concentrated in the gray matter around the central cord, enabling further research into CCS. Finally, histological results are provided as a reference for the readers.

The present protocol simulating central cord syndrome (CCS) in mice has improved repeatability and minimized operation damage to the experimental animals, avoiding disrupting the anatomical structure excessively. The strategy in this study is advantageous because it allows for research into injury mechanisms by producing consistent results.

Definizione del modello di sindrome del midollo centrale nel topo C57BL/6J

Animal models of central cord syndrome (CCS) could substantially benefit preclinical research. Identifiable anatomical pathways can give minimally ...

Imaging the Gut with "CLARITY"

CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging compatible Tissue hYdrogel) has recently evolved as a valuable technique involving acrylamide embedding to delipidate tissue (without sectioning) and to preserve the 3-D tissue structure for immunostaining. The technique is highly relevant in imaging the dynamic gut environment where different cell types interact during homeostasis and disease states. This method optimized for the mouse gut is described here, which helps to trace cell types like epithelia, enteroendocrine, neurons, glia, and the neuronal projections into the epithelia or enteroendocrine cells that mediate microbial sensing or nutrient chemo sensing respectively. The gut tissue (1-1.5 cm) is fixed in 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS) at 4 &#176;C overnight on day 1. On day 2, PFA is discarded, and the tissue is washed thrice with PBS. The tissue is hydrogel embedded to preserve its integrity by incubation in 4% hydrogel (acrylamide) solution in PBS (diluted from 30% ProtoGel) overnight at 4 &#176;C. On day 3, the tissue-hydrogel solution is incubated at 37&#160;&#176;C for 1 h to allow hydrogel polymerization. Tissue is then washed thrice gently with PBS to remove excess hydrogel. The subsequent step of delipidation (clearing) involves tissue incubation in sodium dodecyl sulfate (8% SDS in PBS) at 37 &#176;C for 2 days (days 4 &#38; 5) on a shaker at room temperature (RT). On day 6, the cleared tissue is thoroughly washed with PBS to remove SDS. Tissue can be immunostained by incubation in primary antibodies (diluted in 0.5% normal donkey serum in PBS containing 0.3% Triton X-100), overnight at 4&#176;C, and subsequent incubation in appropriate secondary Alexa Fluor antibodies for 1.5 h at RT, and nuclear staining with DAPI (1: 10000). The tissue is transferred onto a clean glass slide and mounted using VectaShield for confocal imaging.

We describe the protocol for passive CLARITY (PACT) staining of mouse intestine to enable visualization of subepithelial tissues, including neurons, glia and enteroendocrine cells (EEC) without tissue sectioning. The protocol involves hydrogel embedding of formaldehyde-fixed tissue, and subsequent delipidation using an anionic detergent to "clear" the tissue.

CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging compatible Tissue hYdrogel) has recently evolved as a valuable technique involving ...