Questo lavoro è stato reso possibile grazie al sostegno finanziario della Fondation pour la Recherche Médicale (FRM), la Compagnia Milton Safenowitz post-dottorato di ricerca per la SLA (ALS Association), NIH Grants NINDS NS05462 e NS034382, e ANR di Grant HyperMND.

1. Fase uno Pre-anestetico farmaci: 10-15 minuti prima dell&#39;induzione dell&#39;anestesia, iniettare atropina (0,20 mg / kg) e methylprenidsolone (0,05 mg) sottocute per prevenire ipersalivazione ed edema, rispettivamente. 2. Fase due Induzione di anestesia: iniettare sodio pentobarbital (70 mg / kg) o di una miscela di chetamina / xilazina (100 mg / kg e 10 mg / kg, rispettivamente) intra-peritoneale. Lasciate il mouse andare sotto fino a quando non reflex pizzico punta del piede può essere ottenuto. Se l&#39;anestesia sembra troppo leggero, integrare con 1/4 della dose. 3. Fase tre * Nota: questo tipo di chirurgia è una procedura terminale. Quando un aereo chirurgico di anestesia è stato raggiunto, trasferire il mouse su una coperta calda sollevata in posizione prona. <ol><li> Coprire il muso del topo con una maschera fornire puro O 2 ad un flusso di 100 ml / min. </li><li&gt; Utilizzo di un tagliatore di capelli, la barba la regione dorsale dalla nuca del collo alla base della coda. </li><li> Shave anche il diritto di arti posteriori. </li><li> Capovolgere il mouse in posizione supina, ma fare attenzione a lasciare la maschera per l&#39;ossigeno sul posto. </li></ol> 4. Fase quattro Fissare il mouse in posizione con suture occhiello attorno al arti e fissata ai quattro angoli della superficie di lavoro. 5. Fase cinque Inserire una sonda di temperatura per monitorare la temperatura interna del mouse. Regolare coperta termica / potenza della lampada per mantenere la temperatura interna tra 36 ° C e 38 ° C. 6. Tracheotomia e la ventilazione artificiale <ol><li> Utilizzando le forbici spuntate, fare un taglio sopra la trachea e tirare la pelle di distanza su entrambi i lati. </li><li> Utilizzando un forcipe smussato, lacerare la ghiandola salivare modo da esporre due muscoli sottili (sterno-ioideo) coprono la trachea. </li><li> Utilizzando pinze smussato, separare i due muscles lungo il loro separazione mediale per rivelare la trachea. </li><li> Utilizzando un 7 forcipe Dumont, far scorrere due lunghezze di 4,0 sutura di seta sotto la trachea. </li><li> Praticare un taglio trasversale nella trachea tra due anelli cartilaginei, ma fate attenzione a non completamente la sezione della trachea. </li><li> Inserire il tubo tracheale lungo la trachea, quindi fissarla su entrambi i lati dell&#39;apertura di introduzione legando le suture sopra. Il tubo tracheale è collegata ad un ventilatore mouse (SAR-830/AP, CWE Inc.) e un capnografo (μcapstar, CWE Inc.). Il ventilatore è collegato, attraverso un sacchetto rispetto, ad una sorgente di pura O 2. Regolare i parametri del ventilatore (respirazione tasso di 100-150 bpm, il volume di fine espirazione 170-310 pl) in modo che il mouse non sta combattendo contro la ventilazione artificiale e di fine espirazione pCO 2 è stabile tra il 4 e il 5%. </li></ol> 7. Posizionamento delle linee per via endovenosa <ol><li> Usando le tecniche di dissezione smussa, esporre la giugularevena su un lato del collo. A questo livello, la vena giugulare si divide in due tronchi principali, le vene anteriori e posteriori per il viso. </li><li> Effettuare le seguenti operazioni per due volte, una serie per ciascuna di queste linee: <ol><li> Utilizzando Dumont 4 o 5 pinze, separare delicatamente la vena dal suo tessuto circostante congiuntivo. </li><li> Utilizzando Dumont 7 pinze, far scorrere due lunghezze di 6,0 punti di sutura in seta sotto la vena. separarli per quanto possibile lungo la lunghezza della vena. </li><li> Posizionare una clip piccola imbarcazione sul lato prossimale della vena (il lato più vicino al cuore), e legare il lato distale della vena. </li><li> Utilizzo molto fine forbici iris, fare una piccola incisione trasversale nella vena, avendo cura di non grande sezione della vena completamente. </li><li> Inserire preriempita catetere 1FR (premicath, Vygon) nell&#39;apertura, fino al clip nave. </li><li> Tenendo la vena e il catetere insieme in un 4 forcipe Dumont, rimuovere con attenzione la clip nave, quindi spingere il catetere una mor pochimillimetri posta nella vena. </li><li> Fissare il catetere legandola con entrambe le suture su entrambi i lati l&#39;inserimento. </li></ol></li><li> Uno dei cateteri è collegato ad una siringa o una pompa a siringa per iniettare dosi supplementari di anestetici se necessario (solitamente ogni 10-30 min). La dose IV è o 6 mg / kg di sodio pentobarbital o 1250 40 mcg / kg / min di ketamina / xilazina 12. L&#39;altro catetere è collegato ad una pompa a siringa per infusione endovenosa lenta (50 microlitri / h) di una soluzione contenente glucosio 4% NaHCO 3 (1%) e plasmion (14%). </li></ol> 8. Chiudere la pelle del collo con ago e sutura, e torna il mouse alla posizione prona 9. La dissezione del muscolo degli arti posteriori e Nervi <ol><li> Con forbici, praticare un&#39;incisione dalla cima della coscia al tendine di Achille. Separare la pelle dai muscoli sottostanti, facendo attenzione a non danneggiare i vasi sanguigni. Cauterizzare come necessario. </li><li&gt; Identifica il bordo anteriore del bicipite femorale, che appare come una linea bianca che corre lungo il coscia. Con forbici, aprire delicatamente lungo la linea dal ginocchio fino all&#39;osso dell&#39;anca. Separare i muscoli. Il nervo sciatico è ora visibile sotto il bicipite femorale. </li><li> Attenzione sezionare il bicipite femorale. Cauterize / legatura come necessario per prevenire le emorragie. Il bicipite femorale può essere completamente rimosso o semplicemente reclinata per esporre il nervo sciatico e il tricipite surale muscoli. </li><li> Staccare il nervo surale. </li><li> Utilizzo di 8/0 filo di seta, legare la parte distale del nervo peroneo comune e tagliare distale al nodo. Staccare il nervo tutta la strada fino, il più vicino al fianco il più possibile. </li><li> Identificare il nervo tibiale tra i nervi peroneo comune e Sural. Tra i diversi rami del nervo tibiale, individuare i rami che innervano il tricipite surale dai rami che vanno più profondo (d&#39;ora in poi indicato come nervo tibiale). </li><li> Utilizzando 8/0 setafilo, legare insieme tutti i rami del nervo tibiale, mantenendo intatti i rami innervano il tricipite surale. Tagliare il nervo tibiale distalmente al nodo e sezionare il nervo up per quanto possibile. </li><li> Coprire l&#39;intera hindlimb con una garza imbevuta con soluzione salina per evitare essiccazione mentre si procede con il passo successivo. </li></ol> 10. Laminectomia <ol><li> Utilizzando le forbici, fare un&#39;incisione lungo la spina dorsale. Separare la pelle dai muscoli sottostanti. </li><li> Fare due incisioni su ciascun lato delle vertebre per separare i muscoli sottocutanei. Quindi tagli ogni tendine dei muscoli che si attaccano sul lato delle vertebre su ogni lato. </li><li> Utilizzando pinze smussati e una curette fine, rimuovere il resto del muscolo sul lato dorsale delle vertebre intorno ai processi spinosi di identificare chiaramente ogni singola vertebra. </li><li> Identificare vertebre T13 e L1. T13 è l&#39;ultima vertebra di avere nervature allegato. La colonna vertebrale will essere immobilizzato con i Spinal Cord morsetti vertebrali Cunningham (Stoelting Co.). Posizionare i morsetti spinali su ciascun lato del T13 e L1. Fare attenzione a non comprimere il midollo spinale, ma mettere un po &#39;di tensione lungo l&#39;asse longitudinale. Verificare che la colonna vertebrale è ben protetto premendo delicatamente con una pinza. </li><li> Utilizzando rongeurs sottili, rimuovere i processi spinali poi le lamine sopra T13 e L1, esponendo così il midollo spinale. </li><li> Coprire il midollo spinale esposto con piccoli pezzi di cotone o spongel assorbito con soluzione fisiologica per prevenire l&#39;essiccamento. </li><li> Inserire un costume da bagno in plastica sulla parte superiore della schiena, che circonda il midollo spinale. Fissare il bagno in posizione con 4/0 filo di seta. Assicurarsi che il bagno è impermeabile, sigillando con Kwik-Cast sigillante (WPI). </li><li> Una volta che il Kwik-Cast è asciugato, rimuovere il cotone che copre il midollo spinale, e riempire la vasca con olio minerale. </li><li> Utilizzando Dumont 5 pinze e molto sottili forbici iris, delicatamente tirare il compagno durar circonda il midollo spinale, e aprirlo quanto possibile in entrambe le direzioni. Piegare la dura su ciascun lato del midollo spinale. </li><li> Abbassare la piattaforma sollevata su cui il mouse viene disteso in modo da tenerla sospesa con il morsetto vertebrale. </li><li> Utilizzare un altro morsetto vertebrale per serrare un processo spinoso della regione sacrale di sostenere la regione posteriore dell&#39;animale. </li><li> Un terzo morsetto viene usato per immobilizzare l&#39;arti posteriori destro e caviglia, flesso a un angolo di 90 ° al ginocchio. </li></ol> 11. Del tendine di Achille Dissection <ol><li> Con l&#39;arto piegato a 90 ° al ginocchio e alla caviglia, rimuovere la garza che copre la zona degli arti posteriori, e sezionare il tendine di Achille libera dei tessuti circostanti. Sezionare il più possibile il tricipite surale dal tessuto circostante pure. </li><li> Tagliare il tendine del muscolo plantare vicino al calcagno, quindi tagliare nuovamente per rimuovere completamente il tendine. </li><li> Utilizzando un ago filettata, collegare 6/0 di seta filo attraverso °e tendine di Achille e fare un nodo triplo intorno al tendine. </li><li> Posizionare il trasduttore di forza vicino al nodo, tagliare la parte distale del tendine, e collegarlo al trasduttore di forza con il filo di seta. </li><li> Inserire due lunghezze di filo di acciaio inossidabile sotto la fascia dei muscoli tricipite surale. Questi fili sono collegati ad un amplificatore extracellulare AC per la registrazione EMG. </li><li> Posizionare il nervo peroneo comune e il nervo tibiale su due elettrodi bipolari gancio. </li><li> Posizionare il tricipite surale nervo sul catodo di un elettrodo gancio, con l&#39;anodo toccando un muscolo vicina. </li><li> Collegare tutti gli elettrodi stimolanti ad una unità di isolamento. </li><li> Posizionare un elettrodo palla sulla superficie dorsale del midollo spinale, collegato ad un amplificatore extracellulare CA per registrare potenziali cordone dorso. Inserire un Ag / AgCl elettrodo di riferimento in contatto con un muscolo posteriore. </li></ol> 12. Passo Dodici Stimolare il tricipite surale nervoutilizzando un impulso di 50 psec quadrato di crescente intensità a bassa frequenza (&lt;1 Hz) fino a quando l&#39;ampiezza massima contrazione si osserva. Spostare lentamente il trasduttore di forza di allungare il muscolo monitorando l&#39;ampiezza della risposta neuromuscolare finché l&#39;ampiezza contrazione raggiunge un massimo. 13. Registrazioni intracellulari dei motoneuroni Da questo punto in poi, le tecniche standard elettrofisiologiche sono usati per preparare un elettrodo intracellulare, penetrare un neurone nel midollo spinale e identificare come motoneuroni. <ol><li> Tirare una micropipetta di vetro a una punta ~ 1 micron utilizzando un estrattore pipetta (P-97 Micropipetta Puller, Strumenti Sutter). Riempire l&#39;elettrodo con una soluzione di KCl 3M (resistenza dell&#39;elettrodo 10-20 MW). </li><li> Utilizzando un micropositioner, guidare la micropipetta, collegato ad un amplificatore intracellulare (Axoclamp 2B, Axon Instruments) nel midollo spinale. Monitorare i potenziali di campo locali suscitate dalla stimolazione di each coraggio di individuare il parco macchine del tricipite surale. </li><li> Avvicinarsi con cautela motoneuroni putativi monitorando la resistenza del microelettrodo. Quando spinge contro una membrana, la resistenza aumenta. La penetrazione può a volte essere facilitata utilizzando la funzione &quot;buzz&quot; dell&#39;amplificatore intracellulare. </li></ol> 14. Eutanasia Procedura Alla fine dell&#39;esperimento, l&#39;animale è eutanasia da una overdose di pentobarbital (210 mg / kg IV), seguita da decapitazione.

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Nessun conflitto di interessi dichiarati.

Il preparato qui descritto è il primo che permette, nel topo adulto, registrazione simultanea intracellulare di una motoneuron lombare e la misura della forza prodotta dalle fibre muscolari innervate dal suo assone. A causa della piccola dimensione dell&#39;animale, le capacità chirurgici necessari per questa preparazione può essere difficile da acquisire. Tuttavia, una volta che tali capacità sono padronanza, tutta la chirurgia può essere eseguita in tre ore, e gli animali possono sopr...

A correction was made to <a href="http://www.jove.com/video/4312">Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo</a>. There was an error in the name of one author, Marin Manuel. The author's name has been corrected to:
Marin Manuel 
 
instead of: 
 
Manuel Marin

A correction to author list has been made for article <a href="http://www.jove.com/video/4312">Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo</a>.

Erratum: Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In Vivo

<table border="1"><tbody><tr><td> Nome del reagente </td><td> Azienda </td><td> Numero di catalogo </td><td> Commenti (opzionale) </td></tr><tr><td> Atropina solfato </td><td> Aguettant </td><td></td><td></td></tr><tr><td> Methylprenidsolone </td><td> Pfizer </td><td> Solu-Medrol </td><td></td></tr><tr><td> Sodio pentobarbitone </td><td> Sanofi-Aventis </td><td> Pentobarbital </td><td></td></tr><tr><td> Ketamina </td><td></td><td></td><td></td></tr><tr><td> Xylazina </td><td></td><td></td><td></td></tr><tr><td> Glucosio </td><td></td><td></td><td></td></tr><tr><td> Plasma Expander </td><td> Roger Bellon </td><td> Plasmagel </td><td></td></tr><tr><td> Forbici Blunt </td><td> FST </td><td> 14079-10 </td><td></td></tr><tr><td> Blunt forbici sottili </td><td> FST </td><td> 15025-10 </td><td></td></tr><tr><td> Vannas Primavera Forbici </td><td> FST </td><td> 15002-08 </td><td></td></tr><tr><td> Belle pinza dentata </td><td> FST </td><td> 11370-32 </td><td></td></tr><tr><td> Belle pinza dentata </td><td> FST </td><td> 11370-31 </td><td></td></tr><tr><td> Cunningham Adattatore spinale </td><td> Stoelting Co. </td><td></td><td></td></tr><tr><td> Kwik-Cast sigillante </td><td> WPI </td><td> # KWIK-CAST </td><td></td></tr><tr><td> Ventilatore </td><td> CWE Inc </td><td> SAR-830/AP </td><td></td></tr><tr><td> Capnografo </td><td> CWE Inc </td><td> μcapstar </td><td></td></tr><tr><td> Riscaldamento coperta </td><td> Harvard Apparatus </td><td> 507221F </td><td></td></tr><tr><td> Amplificatore intracellulare </td><td> Axon Instrumenti </td><td> Axoclamp 2B </td><td></td></tr><tr><td> Pipettare estrattore </td><td> Sutter Instruments </td><td> P-97 </td><td></td></tr><tr><td> KCl </td><td> Sigma-Aldrich </td><td> P9333-500G </td><td></td></tr></tbody></table>

simultaneous intracellular recording of a lumbar motoneuron and the force produced by its motor unit in the adult mouse in vivo

Il motoneurone spinale è da tempo un sistema di buon modello per studiare la funzione neurale perché è un neurone del sistema nervoso centrale con le proprietà uniche di (1) avente obiettivi facilmente identificabili (le fibre muscolari) e quindi con una funzione molto noto (per controllare contrazione muscolare), (2) essendo il bersaglio convergente di molte reti spinali e discendente, da cui il nome di &quot;definitiva percorso comune&quot;, e (3) avente una grande soma che rende possibile penetrare con taglienti elettrodi intracellulari . Inoltre, quando studiata in vivo, è possibile registrare simultaneamente l&#39;attività elettrica dei motoneuroni e la forza sviluppata dal loro obiettivi muscolari. Esecuzione di registrazioni intracellulari di motoneuroni in vivo quindi mettere il sperimentatore nella posizione unica di poter studiare, allo stesso tempo, tutti i comparti del &quot;gruppo motore&quot; (il nome dato al motoneurone, il suo assone, ele fibre muscolari che innerva 1): gli ingressi che influiscono sul motoneurone, le proprietà elettrofisiologiche del motoneurone, e l&#39;impatto di queste proprietà sulla funzione fisiologica dei motoneuroni, cioè la forza prodotta dal suo gruppo motore. Tuttavia, questo approccio è molto impegnativo, perché la preparazione non può essere paralizzato e quindi la stabilità meccanica per la registrazione intracellulare è ridotta. Pertanto, questo tipo di esperimenti è stato conseguito solo nei gatti e nei ratti. Tuttavia, lo studio dei sistemi spinali motore potrebbe fare un salto formidabile se era possibile effettuare esperimenti simili in topi normali e geneticamente modificati. Per motivi tecnici, lo studio delle reti spinali nei topi è stata perlopiù limitata a neonatale in preparati in vitro, in cui i motoneuroni spinali e le reti sono immaturi, i neuroni motori sono separati dai loro obiettivi, e quando ha studiato a fette, il motoneurons sono separati dalla maggior parte dei loro ingressi. Fino a poco tempo, solo pochi gruppi erano riusciti a effettuare registrazioni intracellulari di motoneuroni in vivo 2-4, compresa la nostra squadra che ha pubblicato un nuovo preparato che ci ha permesso di ottenere le registrazioni molto stabili di motoneuroni in vivo in topi adulti 5,6. Tuttavia, queste registrazioni sono state ottenute in animali paralizzati, cioè senza la possibilità di registrare la forza di uscita di questi motoneuroni. Qui vi presentiamo una estensione di questa preparazione originale in cui siamo stati in grado di ottenere le registrazioni simultanee delle proprietà elettrofisiologiche dei motoneuroni e della forza sviluppata dal loro gruppo motore. Questo è un risultato importante, in quanto consente di identificare i diversi tipi di motoneuroni base al loro profilo di forza, e rivelando così la loro funzione. Accoppiato con modelli genetici disturbare circuito spinale segmentale 7-9, o reproducting umana DiseÃse 10,11, ci aspettiamo che questa tecnica sia uno strumento essenziale per lo studio del sistema motorio spinale.

La figura 1 mostra come identificare un motoneuron dal gruppo tricipite surale dopo la penetrazione. A bassa intensità di stimolazione, solo una EPSP monosinaptico può osservare (Figura 1A). A maggiore intensità, l&#39;EPSP potrebbe essere sufficiente per attivare un &quot;ortodromica&quot; spike (Figura 1B). A intensità di stimolazione ancora più in alto, un tutto-o-niente antidromica spike appare, con una latenza inferiore al EPSP monosinaptico (Figura 1C...

Watch this Scientific Journal Video about Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo at JoVE.com

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

Questo nuovo metodo permette la registrazione simultanea intracellulare di una motoneuron singolo topo adulto e la misura della forza prodotta dalle sue fibre muscolari. L&#39;indagine combinato delle proprietà elettriche e meccaniche delle unità motorie negli animali normali e geneticamente modificato è una svolta per lo studio del sistema neuromuscolare.

Registrazione simultanea intracellulare di un motoneuroni lombare e la forza prodotta dal suo gruppo motore nel topo adulto<em&gt; In vivo</em

The  spinal motoneuron has long been a good model system for studying neural  function because it is a neuron of the central nervous system with the ...

simultaneous-intracellular-recording-lumbar-motoneuron-force-produced

Research

JoVE Journal

Neuroscience

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

14.6K Views.  Northwestern University Feinberg School of Medicine. Questo nuovo metodo permette la registrazione simultanea intracellulare di una motoneuron singolo topo adulto e la misura della forza prodotta dalle sue fibre muscolari. L'indagine combinato delle proprietà elettriche e meccaniche delle unità motorie negli animali normali e geneticamente modificato è una svolta per lo studio del sistema neuromuscolare.

Video: Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

Trasfezione delle cellule gangliari della retina di topo In vivo Elettroporazione

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Ricerca

Neuroscienze

A Functional Motor Unit in the Culture Dish: Co-culture of Spinal Cord Explants and Muscle Cells

Human primary muscle cells cultured aneurally in monolayer rarely contract spontaneously because, in the absence of a nerve component, cell differentiation is limited and motor neuron stimulation is missing1. These limitations hamper the in vitro study of many neuromuscular diseases in cultured muscle cells. Importantly, the experimental constraints of monolayered, cultured muscle cells can be overcome by functional innervation of myofibers with spinal cord explants in co-cultures.
Here, we show the different steps required to achieve an efficient, proper innervation of human primary muscle cells, leading to complete differentiation and fiber contraction according to the method developed by Askanas2. To do so, muscle cells are co-cultured with spinal cord explants of rat embryos at ED 13.5, with the dorsal root ganglia still attached to the spinal cord slices. After a few days, the muscle fibers start to contract and eventually become cross-striated through innervation by functional neurites projecting from the spinal cord explants that connecting to the muscle cells. This structure can be maintained for many months, simply by regular exchange of the culture medium.
The applications of this invaluable tool are numerous, as it represents a functional model for multidisciplinary analyses of human muscle development and innervation. In fact, a complete de novo neuromuscular junction installation occurs in a culture dish, allowing an easy measurement of many parameters at each step, in a fundamental and physiological context.
Just to cite a few examples, genomic and/or proteomic studies can be performed directly on the co-cultures. Furthermore, pre- and post-synaptic effects can be specifically and separately assessed at the neuromuscular junction, because both components come from different species, rat and human, respectively. The nerve-muscle co-culture can also be performed with human muscle cells isolated from patients suffering from muscle or neuromuscular diseases3, and thus can be used as a screening tool for candidate drugs. Finally, no special equipment but a regular BSL2 facility is needed to reproduce a functional motor unit in a culture dish. This method thus is valuable for both the muscle as well as the neuromuscular research communities for physiological and mechanistic studies of neuromuscular function, in a normal and disease context.

Cultured muscle cells are an inadequate model to recapitulate innervated muscle in vivo. A functional motor unit can be reproduced in vitro by innervation of differentiated human primary muscle cells using rat embryo spinal cord explants. This article describes how co-cultures of spinal cord explants and muscle cells are established.

Un gruppo motore funzionale nella piastra di coltura: Co-coltura degli espianti midollo spinale e le cellule muscolari

Human primary muscle cells cultured  aneurally in monolayer rarely contract spontaneously because, in the absence of  a nerve component, cell ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

Retrograda Etichettatura fluorescente Consente mirata extracellulare singola unità di registrazione da parte dei neuroni identificati<em&gt; In vivo</em

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

Time-lapse imaging delle Migrazioni neuroblasti in fette acuta del prosencefalo topo adulto

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Multi-unit Metodi di registrazione per caratterizzare l&#39;attività neurale nel Locust (<em&gt; Schistocerca Americana</em&gt;) Circuiti olfattive

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

Una guida per<em&gt; In vivo</em&gt; Singola unità di registrazione da Optogenetically Identificato corticale inibitorio interneuroni

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

Simultaneo<em&gt; Ex vivo</em&gt; Test funzionale di due retine per<em&gt; In vivo</em&gt; Sistema Elettroretinogramma

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

Simultanea a due fotoni<em&gt; In Vivo</em&gt; Imaging di Synaptic ingressi e degli obiettivi postsinaptici nella corteccia mouse retrosplenial

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Metodo elettrofisiologiche per la registrazione intracellulare di tensione Risposte di<em&gt; Drosophila</em&gt; Fotorecettori e Interneuroni a luce stimoli<em&gt; In Vivo</em

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.