The authors sincerely thank the Canadian Institutes of Health Research MOP 130258 and the Star Foundation for Children&#8217;s Health Research, along with the Molly Towell Foundation, for the provision of the research facility and financial support. The authors also sincerely thank Dr. Kinkead Richard for manuscript proofreading and advice.

Questo metodo richiede l&#39;utilizzo di soggetti animali, consentite dalla Laval University comitato etico degli animali (protocollo # 2012-170). 1. Impostazione e preparazione <ol><li> Soluzioni <ol><li> Preparare soluzioni aCSF secondo le seguenti ricette 7,17. Altre ricette con variazioni di concentrazione sono disponibili in letteratura. Soluzioni memoria Stock a 4 ° C per un massimo di un mese. <ol><li> Soluzione salina: aggiungere 75,39 g di NaCl (129 mm finale); 2,5 g di KCl (3,35 mM finale); 0,81 g di NaH2PO4 (0,58 mM finale); 2,33 g di MgCl2 (1,15 mM finale); 1,85 g di CaCl2 (1,26 mM). Sciogliere in 800 ml di acqua distillata poi riempire a 1 L con acqua distillata. </li><li> Soluzione di bicarbonato: aggiungere 17,65 g di NaHCO 3 (21 mm finale). Sciogliere in 900 ml di acqua distillata poi riempire a 1 L con acqua distillata. Variazioni della concentrazione di bicarbonato provocheranno variazioni di pH. </li><li> Soluzione di glucosio: aggiungi 54.06g di glucosio (30 mM finale). Sciogliere in 400 ml di acqua distillata poi riempire a 500 ml con acqua distillata. </li></ol></li><li> Preparare aCSF diluendo 100 ml di soluzione salina, 100 ml di soluzione di bicarbonato, e 50 ml di soluzione di glucosio in 1 L di acqua distillata. Conservare a 4 ° C fino al momento dell&#39;uso. </li><li> Preparare un elettrodo di vetro riscaldando e si estende un tubo di vetro fino a rompersi. Sand giù la punta prima dell&#39;uso. L&#39;elettrodo può essere riutilizzato molte volte finché rimane pulito. </li></ol></li><li> Setup sperimentale Nota: I dati di set-up sono presentati nella figura 2. <ol><li> Accendere l&#39;amplificatore, averager, sistema di acquisizione dati, controllo della temperatura e della pompa in movimento. Controllare l&#39;amplificazione del segnale (guadagno = 10.000), la filtrazione (soglia bassa, 10 Hz; soglia alta, 5 kHz), e la frequenza di campionamento della conversione da analogico a digitale del segnale grezzo (2,5 kHz). </li><li> Riempire una bottiglia con aCSF e bolla con CarboGen (95% O 2 </sub&gt;, 5% di CO 2). Inducono una bubbled- e riscaldato aCSF flusso nella camera di registrazione (volume 5 ml) a 4 ml / min. Mantenere la temperatura nella camera di registrazione a 26 (± 1) ° C. pH deve essere 7,4 in queste condizioni standard. </li><li> Conservare 50 ml di RT e CarboGen-bolle aCSF in una siringa da 50 ml in prossimità della camera di dissezione. </li><li> Accendere il computer e avviare il software di registrazione. </li></ol></li></ol> 2. Dissection <ol><li> Pesare e determinare visivamente il sesso dell&#39;animale (i maschi hanno un ano-genitale distanza più lunga, mentre le femmine hanno un breve tratto ano-genitale, i maschi genitali sono spesso oscuro mentre genitali femminili sono di colore rosa). </li><li> Anestetizzare i roditori neonati da una delle seguenti opzioni: cryoanesthesia (immergere completamente l&#39;animale in ghiaccio per 4 - 5 min 18), iniezione (equitensine a 4 ml / kg) 19 o inalazione di anestetici volatili (isoflurano 20 o etere 21). Confirm un piano adeguata di anestetici dall&#39;assenza di un ritiro riflesso zampa. </li><li> Posto l&#39;animale in panchina, faccia ventrale giù. Sezione della parte rostrale della testa coronale con un bisturi a livello di bregma (visibile attraverso la pelle e il cranio). Eseguire questo passaggio subito dopo che l&#39;animale è anestetizzato. </li><li> Sezione coronale il corpo con un bisturi sotto i membri anteriori. </li><li> Togliere la pelle, i muscoli, tessuti grassi e viscere con le forbici chirurgiche e sottili e pinze. Dal momento che le ossa sono morbidi a questa età, essere prudenti per non danneggiare il sistema nervoso. Eseguire questo passaggio in panchina. </li><li> Posizionare la preparazione nella camera dissezione. Utilizzare il aCSF memorizzato nella siringa per ossigenare preparazione. Da questo punto alla fine della registrazione, utilizzare un microscopio. </li><li> Sulla faccia dorsale del preparato, tagliare il cranio e vertebre dal rostrale alla parte caudale lungo l&#39;asse medio con forbici chirurgiche e sottili e pinze. Aprire il cranio taglioe vertebre, al fine di esporre il tessuto nervoso. Utilizzare il aCSF strored nella siringa per ossigenare la preparazione. </li><li> Con le pinze, rimuovere la membrana aracnoidea, un sottile tessuto che ricopre la superficie del tessuto nervoso. Conservare la pia madre e vasi sanguigni contro il tessuto nervoso. Utilizzare il aCSF memorizzato nella siringa per ossigenare preparazione. </li><li> Posizionare la preparazione con il dorso rivolto verso il basso, e con attenzione abbassare il tronco encefalico e del midollo spinale tagliando i nervi e tessuto connettivo con le forbici mentre si tiene la preparazione sul posto. Un approccio dorsale è anche possibile. Mantenere le radici e nervi più a lungo possibile. Togliere le ossa per isolare il tronco encefalico e del midollo spinale. Utilizzare il aCSF memorizzato nella siringa per ossigenare preparazione. </li><li> Rimuovere il cervelletto e rimanendo strutture rostrali da loro sezionamento con il bisturi. Utilizzare il aCSF memorizzato nel ago per ossigenare preparazione. </li><li> Opzionalmente seconda sperimenprogettazione tal, rimuovere il ponte con il bisturi da una sezione anteriore coronale alla inferiore arteria cerebellare 22. Notare che mantenendo tutta ponte rallenterà il ritmo in ratti e completamente inibirla in topi. Preparazioni che includono solo la parte caudale del ponte mostrano una attività respiratoria simile con una frequenza stabile alla radice C4. </li></ol> 3. Registrazione <ol><li> Mettere il preparato in camera di registrazione, ventrale a faccia in su. Fissare la preparazione con perni nella parte più bassa del midollo spinale e rostrale-maggior parte del tronco cerebrale. </li><li> Utilizzando una siringa collegata all&#39;elettrodo dal suo ago, indurre una depressione nell&#39;elettrodo (diametro punta di vetro: 150 - 225 micron) tirando il pistone della siringa, per riempire parzialmente l&#39;elettrodo con aCSF. </li><li> Utilizzando un micromanipolatore, posizionare accuratamente l&#39;elettrodo vicino ad una delle quarte radici ventrali. Altre radici ventrali potrebbero anche presentare una delle vie respiratorie come l&#39;attività (e.g., XII nervo cranico, C1 radice ventrale). </li><li> Inducono una depressione nel serbatoio dell&#39;elettrodo tramite la siringa estraendo lo stantuffo per aspirare delicatamente il rootlet nervo. Quindi spostare accuratamente l&#39;elettrodo di applicare contro il midollo spinale. Nota: Idealmente la dimensione del rootlet corrisponde alla dimensione dell&#39;apertura elettrodo e crea una tenuta tra i compartimenti interni ed esterni dell&#39;elettrodo così. Poiché gli amplificatori differenziali sono comunemente utilizzati per tali registrazioni, ogni apertura tra l&#39;interno dell&#39;elettrodo e la camera di registrazione riduce la qualità del segnale e rende difficile per eliminare il rumore di fondo. </li><li> Avviare la registrazione. </li><li> Registra il ritmo prodotto dalla preparazione in condizioni di normossia (cioè, aCSF gorgogliare con CarboGen: 95% O 2 e 5% CO 2) per almeno 20 minuti per determinare i parametri di base della preparazione. </li><li> Passare la perfusione da CarboGen-bolle aCSF astimolo aCSF (cioè, con il 95% gorgogliare N 2 e 5% di CO 2 per ipossia stimolo) per 15 min. Alter durata dell&#39;esposizione a seconda del protocollo sperimentale. Registrare la durata dell&#39;esposizione sulla registrazione degli animali e scheda tecnica. Utilizzare tubi in acciaio inossidabile tra il flacone e la camera aCSF quando possibile per evitare la diffusione di gas verso l&#39;esterno del tubo. </li><li> Accendere la perfusione torna a aCSF CarboGen-bolle standard per almeno 15 minuti per una registrazione di recupero. Registrare questo sulla registrazione degli animali e scheda tecnica. </li><li> Terminare la registrazione. </li></ol> 4. Analisi statistica <ol><li> Dal segnale integrato, calcolare la frequenza del numero di burst per minuto (espressa in burst / min), l&#39;ampiezza come differenza tra la linea di base e il picco del burst (espressa in mV), la durata di scoppio della durata da l&#39;inizio e la fine del burst (espresso in s), l&#39;area di scoppio come l&#39;area sotto la curve del burst sul segnale integrato (espressa in mV · s) (Figura 3). </li><li> Calcolare la frequenza, ampiezza, durata scoppiare e zona scoppiare sotto normossia (baseline), ipossia (stimolo) e post-ipossia condizioni (post-stimolo) di recupero come la media degli ultimi 5 minuti delle registrazioni per ogni condizione. Non analizzare la prima porzione di ciascuna condizione perché c&#39;è un ritardo tra l&#39;accensione e perfusione aCSF omogeneizzazione nella camera di registrazione. </li><li> Express poi stimolo (cioè, ipossia) e post-stimolo (cioè, dopo ipossia) valori di recupero come percentuale dei valori di base per la registrazione corrispondente. Esprimere i risultati come media ± SD </li></ol>

<ol>
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The authors have no competing financial interests or conflicts of interests to disclose.

Quantificazione accurata dell&#39;attività respiratoria può essere impegnativo. Infatti, la respirazione è una funzione che può essere sia automatica e volontaria, e che è modulata in base all&#39;ambiente, le esigenze del corpo, lo stato emotivo ed il comportamento. Il vantaggio di questa tecnica è l&#39;isolamento degli elementi neurali responsabili della produzione del comando respiratoria. Così, registrazioni elettrofisiologiche di preparati midollo spinale e tronco-pletismografia sono tecniche complementari ...

La respirazione è un complesso e vitale attività controllata dal cervello, permettendo diossigeno (O 2) e l&#39;assorbimento di anidride carbonica (CO 2) eliminazione. Il drive respiratorio centrale è generato da una complessa rete situata nel tronco cerebrale in entrambi i mammiferi 1, 2 anfibi, rettili, uccelli 3 4 5 e pesci. Anche se lo studio di respirazione può essere elaborata in vivo, precise indagini meccanicistiche richiedono l&#39;accesso diretto alla rete di controllo respiratorio. A tal fine, Adrian e Buytendijk sviluppato una ridotta preparazione pesci rossi, in cui elettrodi posti sulla superficie del tronco cerebrale record ritmo generato associato ventilazione gill 5. Questo approccio è stato in seguito adattato da Suzue nel 1984 6 per l&#39;impiego nei roditori appena nati. L&#39;avvento di questa preparazione ha portato a significativi progressi nella neurobiologia respiratoria. Poiché è relativamente semplice, la tecnica presentato here è suscettibile di una vasta gamma di ricerche di base di comportamenti motori ritmici e le loro origini nei roditori appena nati. L&#39;obiettivo generale di questo metodo è quello di registrare il correlato neurale di attività inspiratoria, un ritmo respiratorio simile chiamato respirazione fittizia, prodotta dalla rete respiratoria. Questo metodo può essere impiegato in una vasta gamma di obiettivi di ricerca, puntando risposte inspiratori alle variazioni respiratorie o farmacologia in entrambi i selvatici tipo 7 e 8 animali transgenici. Dato che gli esperimenti vengono eseguiti a bassa temperatura, senza afferenti sensoriali, e in condizioni in cui le concentrazioni di glucosio e O 2 ai aCSF sono alti, questioni sono state sollevate per quanto riguarda la rilevanza fisiologica del segnale registrato. Mentre vi sono chiare differenze tra in vivo e in vitro condizioni (ad es., La frequenza di raffiche inspiratori) resta il fatto che la presenza digli elementi fondamentali della rete respiratoria 6 permettono di studiare un ritmo robusto associato ad una funzione omeostatica vitale 9,10. La logica dietro lo sviluppo e l&#39;utilizzo di questa tecnica è quello di facilitare l&#39;accesso diretto agli elementi del tronco della rete respiratoria, che sono difficilmente accessibili in vivo, soprattutto nei neonati. Il tronco encefalico è posto in condizioni rigorosamente controllate: il ritmo registrato non è modulato da input afferenti periferici dai polmoni o gli organismi carotidei, permettendo lo studio di concentrarsi sul drive respiratorio centrale stessa 11. Così, questo accesso è utilizzato per applicare stimoli e registrare il segnale di uscita. In contrasto con pletismografia registrazioni, il ritmo respiratorio è modulata da tutti i suoi componenti in tutto il corpo (ad es., Distensione polmonare, chemosensori periferiche), rendendo difficile applicare stimoli precisi. In unratto ewborn, il protocollo consiste nel registrare il quarto segnale ventrale root su un tronco isolato e midollo spinale troncato, mantenuto in fluido cerebrospinale artificiale (aCSF). Il ritmo generato dai preparativi del midollo spinale tronco-è composto da singole raffiche lenti che sono collegati al segnale inspiratorio 9. Isolato preparativi del midollo spinale tronco-sono facilmente registrabili in ratti da post-natale giorno 0-4 (P0 - P4) 7. Questo approccio è comunemente utilizzato per valutare la risposta ipossica della rete respiratoria, e anche la risposta ai ipercapnia, acidosi o droghe. Un protocollo acuta all&#39;ipossia è presentato qui. Questa stimolazione è ottenuto dal prelievo di O 2 nel aCSF; questo approccio è comunemente utilizzato per valutare la tolleranza e la risposta agli insulti ipossici. Il protocollo induce una depressione ritmo dal primo minuto fino alla fine dell&#39;esposizione ipossia (Figura 1) 12. Questa depressione è invertitadurante la post-ipossica 12 recupero. Per quanto riguarda il disegno sperimentale, è importante notare che il ponte, si trovano nella parte rostrale del tronco cerebrale, ha un&#39;azione inibitoria sul generatore ritmo 8. Così, preparazioni di piena del tronco encefalico e del midollo spinale rostrale mostrano un ritmo inferiore. L&#39;inclusione del ponte del campione isolata per la registrazione è determinato secondo il fine dell&#39;esperimento 13; studio dell&#39;influenza pontino sulla rete midollo allungato richiederebbe registrazioni con e senza il ponte di confrontare i risultati 14. Inoltre, uno dei vantaggi di questa tecnica è la possibilità di estendere la parte rostrale della preparazione per includere mesencefalico e / o regioni diencefaliche 15,16, che consenta di valutare l&#39;effetto di queste regioni della rete respiratoria ponto-midollare.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Sylgard</td>
			<td>Sigma Aldrich</td>
			<td>761036-5EA</td>
			<td>Use under hood</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Bioshop</td>
			<td>SOD002</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Bioshop</td>
			<td>POC888</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Bioshop</td>
			<td>CCL444</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Bioshop</td>
			<td>MAG510</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Bioshop</td>
			<td>SOB999</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Bioshop</td>
			<td>SPM306</td>
			<td></td>
		</tr>
		<tr>
			<td>D-glucose</td>
			<td>Bioshop</td>
			<td>GLU501</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbogen</td>
			<td>Linde</td>
			<td>343-02-0006&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature Controller</td>
			<td>Warner Instruments, Hamden, CT, USA</td>
			<td>TC-324B</td>
			<td></td>
		</tr>
		<tr>
			<td>Suction electrode</td>
			<td>A-M Systems, Everett, WA, USA</td>
			<td>model 573000</td>
			<td></td>
		</tr>
		<tr>
			<td>Differential AC amplifier</td>
			<td>A-M Systems, Everett, WA, USA</td>
			<td>model 1700</td>
			<td></td>
		</tr>
		<tr>
			<td>Moving averager</td>
			<td>CWE, Ardmore, PA, USA</td>
			<td>model MA-821</td>
			<td></td>
		</tr>
		<tr>
			<td>Data acquisition system</td>
			<td>Dataq Instruments, Akron, OH, USA</td>
			<td>model DI-720</td>
			<td></td>
		</tr>
		<tr>
			<td>LabChart software</td>
			<td>ADInstruments, Colorado Springs, CO, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Prism sofware</td>
			<td>Graphpad, La Jolla, CA, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection chamber</td>
			<td>Plastic box (e.g. petri box) will do</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Recording chamber</td>
			<td>Home made</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Base</td>
			<td>Kanetec, Bensenville, IL, USA</td>
			<td>MB</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>World Precision Instrument Inc, Sarasota, FL, USA</td>
			<td>KITE-R</td>
			<td></td>
		</tr>
		<tr>
			<td>Base</td>
			<td>Kanetec, Bensenville, IL, USA</td>
			<td>MB</td>
			<td></td>
		</tr>
		<tr>
			<td>Peristaltic pump</td>
			<td>Gilson, Middleton, WI, USA</td>
			<td>MINIPULS 3</td>
			<td></td>
		</tr>
		<tr>
			<td>Faraday Cage</td>
			<td>Home made</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Computer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

electrophysiology on isolated brainstem-spinal cord preparations from newborn rodents allows neural respiratory network output recording

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to stimuli remain to be fully understood. To overcome the difficulty of accessing the brainstem in the whole animal, isolation of the brainstem and part of the spinal cord is performed. This preparation is maintained in artificial cerebro-spinal fluid where gases, concentrations, and temperature are controlled and monitored. The output signal from the respiratory network is recorded by a suction electrode placed on the fourth ventral root. In this manner, stimuli can be directly applied onto the brainstem, and the effect can be recorded directly. The signal recorded is linked to the inspiratory signal sent to the diaphragm via the phrenic nerve, and can be described as bursts (around 8 bursts per minute). Analysis of these bursts (frequency, amplitude, length, and area under the curve) allows precise characterization of the stimulus effect on the respiratory network. The main limitation of this method is the viability of the preparation beyond the early post-natal stages. Thus, this method greatly focuses on the study of the whole network without the peripheral inputs in the newborn rat.

Come accennato nell&#39;introduzione, uno dei vantaggi più importanti di questa tecnica è l&#39;accesso diretto al tronco cerebrale applicare vari stimoli. Come esempio, ipossia stato applicato qui. Figura 1. AB mostra una registrazione completa protocollo, con entrambe le condizioni normossia e ipossia. Figura 1.CE visualizza il ritmo registrata in condizioni di normossia (cioè, aCSF gorgogliare con 95% O 2 e 5% CO 2 a 26 ° C). Come già illustrato in...

Watch this Scientific Journal Video about Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording at JoVE.com

Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording

The central respiratory drive is located in the brainstem. Spontaneous respiratory motor output from an isolated brainstem-spinal cord is recorded by placing an electrode on the fourth ventral root. This experimental approach is valuable for pharmacological investigations or the assessment of respiratory challenges and genetic manipulations on rhythmic motor behavior.

Elettrofisiologia su isolati tronco cerebrale, midollo spinale Ammassi di Newborn Roditore Consente Neural Network respiratoria registrazione Uscita

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to ...

electrophysiology-on-isolated-brainstem-spinal-cord-preparations-from

Research

JoVE Journal

Neuroscience

8.4K Views.  Laval University. The central respiratory drive is located in the brainstem. Spontaneous respiratory motor output from an isolated brainstem-spinal cord is recorded by placing an electrode on the fourth ventral root. This experimental approach is valuable for pharmacological investigations or the assessment of respiratory challenges and genetic manipulations on rhythmic motor behavior.

Video: Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Dissezione e la cultura di neuroni embrionali commissurali da midollo spinale

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Ricerca

Neuroscienze

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Spinal Cord Elettrofisiologia II: extracellulare fabbricazione di elettrodi di aspirazione

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

Examining Local Network Processing using Multi-contact Laminar Electrode Recording

Cortical layers are ubiquitous structures throughout neocortex1-4 that consist of highly recurrent local networks. In recent years, significant progress has been made in our understanding of differences in response properties of neurons in different cortical layers5-8, yet there is still a great deal left to learn about whether and how neuronal populations encode information in a laminar-specific manner.
Existing multi-electrode array techniques, although informative for measuring responses across many millimeters of cortical space along the cortical surface, are unsuitable to approach the issue of laminar cortical circuits. Here, we present our method for setting up and recording individual neurons and local field potentials (LFPs) across cortical layers of primary visual cortex (V1) utilizing multi-contact laminar electrodes (Figure 1; Plextrode U-Probe, Plexon Inc).
The methods included are recording device construction, identification of cortical layers, and identification of receptive fields of individual neurons. To identify cortical layers, we measure the evoked response potentials (ERPs) of the LFP time-series using full-field flashed stimuli. We then perform current-source density (CSD) analysis to identify the polarity inversion accompanied by the sink-source configuration at the base of layer 4 (the sink is inside layer 4, subsequently referred to as granular layer9-12). Current-source density is useful because it provides an index of the location, direction, and density of transmembrane current flow, allowing us to accurately position electrodes to record from all layers in a single penetration6, 11, 12.

A fundamental issue in our understanding of cortical circuitry is how networks in different cortical layers encode sensory information. Here we describe electrophysiological techniques utilizing multi-contact laminar electrodes to record single-units and local field potentials and present analyses to identify cortical layers.

Esaminando l&#39;elaborazione locale di rete che utilizza multi-contatto laminare elettrodi di registrazione

Cortical layers are ubiquitous structures throughout neocortex1-4 that consist of highly recurrent local networks. In recent years, significant progress ...

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 ...

Neural Circuit Recording from an Intact Cockroach Nervous System

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological effects of insecticides. This article describes the scope of cockroach sensory modalities that can be used to assay how an insect nervous system responds to environmental perturbations. Emphasis here is on the escape behavior mediated by cerci to giant fiber transmission in Periplaneta americana. This in situ preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of neuronal activity. Peptides or other chemical reagents can then be applied directly to the nervous system in solution with the physiological saline. Insecticides could also be administered prior to dissection and the escape circuit can serve as a proxy for the excitable state of the central nervous system. In this context the assays described herein would also be useful to researchers interested in limb regeneration and the evolution of nervous system development for which P. americana is an established model organism.

This article describes the cockroach ventral nerve cord dissection and extracellular recordings from the cercal nerve and connectives. Evoked responses are generated by electrical stimulation of the cercal nerve or direct mechanical stimulation of the cerci.

Neural Circuit Registrazione da un intatto scarafaggio Sistema Nervoso

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Aumento del tempo della risposta alla rete alla stimolazione delle colture cellulari neuronali sulle matrici di microelettrodi

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...

Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a signal driving the rhythmic contraction of inspiratory muscles. Rhythmic neural activity generated in the pBC and carried to other neuronal pools to drive the musculature of breathing may be studied using various approaches, including en bloc nerve recordings and transverse slice recordings. However, previously published methods have not extensively described the brainstem-spinal cord dissection process in a transparent and reproducible manner for future studies. Here, we present a comprehensive overview of a method used to reproducibly cut rhythmically-active brainstem slices containing the necessary and sufficient neuronal circuitry for generating and transmitting inspiratory drive. This work builds upon previous brainstem-spinal cord electrophysiology protocols to enhance the likelihood of reliably obtaining viable and rhythmically-active slices for recording neuronal output from the pBC, hypoglossal premotor neurons (XII pMN), and hypoglossal motor neurons (XII MN). The work presented expands upon previous published methods by providing detailed, step-by-step illustrations of the dissection, from whole rat pup, to in vitro slice containing the XII rootlets.

This protocol both visually communicates the brainstem-spinal cord preparation and clarifies the preparation of brainstem transverse slices in a comprehensive step-by-step manner. It was designed to increase reproducibility and enhance the likelihood of obtaining viable, long lasting, rhythmically-active slices for recording neural output from the respiratory regions of the brainstem.

Preparazione di ritmicamente-attivo In Vitro neonatale del roditore del tronco cerebrale-spinale e fetta sottile

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Registrazione ottica a singolo fotone a campo largo nelle sezioni cerebrali utilizzando Colorazione sensibile alla tensione

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an increasingly detailed view of the circuits underpinning spinal pain processing. However, the effects of these connections for broader network activity in the DH remain less well understood because most studies focus on the activity of single neurons and small microcircuits. Alternatively, the use of microelectrode arrays (MEAs), which can monitor electrical activity across many cells, provides high spatial and temporal resolution of neural activity. Here, the use of MEAs with mouse spinal cord slices to study DH activity induced by chemically stimulating DH circuits with 4-aminopyridine (4-AP) is described. The resulting rhythmic activity is restricted to the superficial DH, stable over time, blocked by tetrodotoxin, and can be investigated in different slice orientations. Together, this preparation provides a platform to investigate DH circuit activity in tissue from na&#239;ve animals, animal models of chronic pain, and mice with genetically altered nociceptive function. Furthermore, MEA recordings in 4-AP-stimulated spinal cord slices can be used as a rapid screening tool to assess the capacity of novel antinociceptive compounds to disrupt activity in the spinal cord DH.

The combined use of microelectrode array technology and 4-aminopyridine-induced chemical stimulation for investigating network-level nociceptive activity in the spinal cord dorsal horn is outlined.

Registrazione dell'attività di rete nei circuiti nocicettivi spinali utilizzando array di microelettrodi

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an ...

Comprendere le risposte dei motoneuroni alla stimolazione del midollo spinale

The Ex vivo Preparation of Spinal Cord Slice for the Whole-Cell Patch-Clamp Recording in Motor Neurons During Spinal Cord Stimulation

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to execute sensorimotor behaviors, directly studying the electrical responses of motor neurons with SCS can help us understand the underlying logic of spinal motor modulation. To simultaneously record diverse stimulus characteristics and cellular responses, a patch-clamp is a good method to study the electrophysiological characteristics at a single-cell scale. However, there are still some complex difficulties in achieving this goal, including maintaining cell viability, quickly separating the spinal cord from the bony structure, and using the SCS to successfully induce action potentials. Here, we present a detailed protocol using patch-clamp to study the electrical responses of motor neurons to SCS with high spatiotemporal resolution, which can help researcher improve their skills in separating the spinal cord and maintaining the cell viability at the same time to smoothly study the electrical mechanism of SCS on motor neuron and avoid unnecessary trial and mistake.

Autore in primo piano: Avanzamento della stimolazione del midollo spinale - Esplorazione delle risposte cellulari dei motoneuroni attraverso l'elettrofisiologia del patch-clamp 

This protocol describes a method using a patch-clamp to study the electrical responses of motor neurons to spinal cord stimulation (SCS) with high spatiotemporal resolution, which can help researchers improve their skills in separating the spinal cord and maintaining cell viability simultaneously.

La preparazione ex vivo di una fetta di midollo spinale per la registrazione del patch-clamp a cellula intera nei motoneuroni durante la stimolazione del midollo spinale

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to ...