Questo lavoro è stato sostenuto dal National Institutes of Health (K22NS050137 a JCB).

1. Coltura cellulare <ol><li> Neuroni corticali sono isolate e coltivate utilizzando tecniche precedentemente descritte e placcato in capsule di Petri con fondo di vetro (MatTek Corporation, Ashland, MA) rivestito con poli-D-lisina e laminina 1. </li></ol> 2. Preparare le soluzioni per il magazzino sonde fluorescenti TMRM e H 2 DCF-DA <ol><li> Preparare un 10-mM soluzione madre di TMRM sciogliendo 5,0 mg di TMRM in 1 ml di dimetilsulfossido anidro. Vortex è per 1 min. Poi, fare aliquote e conservarle a -20 ° C, proteggere dalla luce, e utilizzare entro un mese. </li><li> Successivamente, preparare un 10-mM soluzione madre di H 2 DCF-DA, sciogliendo 4,87 mg di H 2 DCF-DA in 1 ml di DMSO anidro. Allo stesso modo, vortice che per 1 min. Poi, fare aliquote e conservarle a -20 ° C, proteggere dalla luce, e utilizzare entro una settimana. </li></ol> 3. Neuroni corticali di ratto di carico con TMRM e H 2 DCF-DA TMRM è un potenziometro, cellula-permeabile indicatore fluorescente che si accumula al suo interno una forte carica negativa dei mitocondri. E &#39;importante utilizzare le concentrazioni basse (10-50 nm) di TMRM per evitare l&#39;auto-spegnimento del TMRM mitocondriale. Poi, il segnale di fluorescenza di TMRM possono essere direttamente legati alla co-ΔΨm attraverso la membrana mitocondriale interna. Una perdita di ΔΨm cause TMRM a fuoriuscire dai mitocondri con conseguente perdita di intensità di fluorescenza. H 2 DCF-DA è la cellula-permeabile sonda convertito in DCF-DA dalle esterasi intracellulari, ei suoi risultati ossidazione DCF fluorescente. La concentrazione finale di H 2 DCF-DA varia tra 2-10 micron e dovrebbe essere testato empiricamente nei neuroni derivati ​​da differenti regioni del cervello in quanto le concentrazioni di carico elevato potrebbe provocare la saturazione del DCF fluorescenza anche in assenza di H 2 O 2. La presenza di ossidanti endogeni o esogeni (ad esempio, di ossido nitrico, perossido di idrogeno) aumenterà DFC intensità di fluorescenza. Di seguito, forniamo un protocollo per il caricamento di neuroni corticali di ratto con TMRM e H 2 DCF-DA. <ol><li> Per caricare i neuroni di ratto corticali con TMRM, in primo luogo, lavare i neuroni coltivati ​​3 volte con tampone Tyrode di (Overlay Testo: TB: 145 mM NaCl, 5 mM KCl, 10 glucosio mm, 1,5 mm CaCl 2, 1 mM MgCl 2, e 10 HEPES mM; aggiustare il pH a 7,4 con NaOH). Poi, preparare 20 Nm di TMRM diluendo i 10 magazzino mM TMRM 1 / 1000 volte in TB e poi aggiungere 2 ml di TMRM diluito per 1 ml di TB. Incubare i neuroni con TMRM per 45 minuti al buio a temperatura ambiente. Dopo 45 minuti, montare il piatto cultura sul palcoscenico del microscopio e iniziare imaging. </li><li> Per caricare i neuroni corticali di ratto con H 2 DCF-DA, lavare i neuroni in coltura 3 volte con TB. Quindi, preparare 2 mM di H 2 DCF-DA diluendo i 10 mm H 2 DCF-DA magazzino 1 / 10 volte in TB e poi aggiungere 2 ml di diluito H 2 DCF-DA per 1 ml di TB. Poi, incubare i neuroni con H 2 DCF-DA per 45 minuti al buio a temperatura ambiente. Dopo 45 minuti, lavare i neuroni 4 volte con TB per rimuovere l&#39;eccesso indicatore fluorescente prima di ottenere le immagini. </li></ol> 4. Vivere l&#39;imaging di neuroni incubate con TMRM per determinare ΔΨm <ol><li> Per eseguire l&#39;imaging dal vivo di neuroni incubate con TMRM, microscopia confocale a scansione laser (Overlay Testo: LSM 510, Carl Zeiss Inc.), con l&#39;applicazione di vivere serie temporali programma, viene utilizzato. Applicare a bassa risoluzione e attenuato laser di potenza (Overlay Testo: bassa risoluzione: 256 x 256, potenza laser: 1%) per ridurre al minimo il tempo necessario per ottenere immagini ed evitare photobleaching. </li><li> . Successivamente, regolare la messa a fuoco dei neuroni montato caricato con TMRM usando la luce riflessa. Esaminare la fluorescenza TMRM di illuminazione a 514 nm e la rilevazione a 570 nm. Impostare il guadagno di rilevamento di una macchina fotografica appena sotto il livello di saturazione. </li><li> Una volta che tutti i parametri che includono la risoluzione, potenza del laser, ottenere il rilevamento di una macchina fotografica, e time-lapse intervallo di ottenere immagini sono serie: non modificare queste impostazioni tra esperimenti. Successivamente, modificare il campo. Iniziare a raccogliere le immagini. </li><li> Per verificare i cambiamenti in ΔΨm, stimoli come 1 microM di FCCP o 2 mg / ml di oligomicina, può essere applicato, causando notevoli depolarizzare o hyperpolarize il potenziale della membrana mitocondriale, rispettivamente. Questi cambiamenti si rifletteranno da una diminuzione dell&#39;intensità TMRM fluorescenza rispetto con l&#39;intensità di fluorescenza di base in caso di FCCP, o un aumento di intensità TMRM fluorescenza in caso di oligomicina. </li></ol> 5. Immagini dal vivo di neuroni incubate con H 2 DCF-DA per determinare ROS <ol><li> Per eseguire l&#39;imaging dal vivo di neuroni incubate con H 2 DCF-DA, in primo luogo, montare il piatto cultura sul palcoscenico di un microscopio. Regolare il fuoco delle cellule noiING luce riflessa. Esaminare DCF fluorescenza da eccitazione a 488 nm ed emissione a 515 nm. </li><li> Quindi, regolare la potenza del laser al 5-7%, il guadagno del rivelatore e risoluzione di 256 x 256. Non modificare queste impostazioni tra esperimenti. Quindi, impostare la frequenza per ottenere le immagini in diretta con il programma di serie temporali. </li><li> Selezionare un campo nuovo e iniziare l&#39;acquisizione di immagini. Per rilevare i cambiamenti nei livelli di ROS, il trattamento di cellule con 100-200 mM di H 2 O 2. Questo sarà tradursi in un aumento di intensità DCF fluorescenza rispetto al livello di base. </li></ol> 6. Analisi dei dati <ol><li> Usa regione di interesse (Overlay Testo: ROI) strumento dal programma LSM per selezionare le aree. Poi, misurare la TMRM o ROS intensità di fluorescenza. Seleziona ROI dalle regioni mitocondriale o ROI dal corpo cellulare nelle cellule intero fotografato per misurare l&#39;intensità di fluorescenza di TMRM o ROS, rispettivamente. </li><li> Calcolare l&#39;intensità media di fluorescenza da tutte le ROI di ogni cella per TMRM o da corpi cellulari intero per tutte le cellule ripreso per ROS per ogni punto del tempo. Selezionare le regioni accanto alle cellule per calcolare l&#39;intensità di fluorescenza di fondo. Prendere le misure diversi e calcolare l&#39;intensità media di fondo. </li><li> Sottrarre la media intensità di fluorescenza di fondo dalla media intensità di fluorescenza di ROI in ciascuna cella per ogni punto di tempo utilizzando Microsoft Excel. Dopo aver sottratto l&#39;intensità di fondo, normalizzare l&#39;intensità di fluorescenza TMRM o DCF per la fluorescenza di base utilizzando questa formula (Overlay Testo: ΔF = FF o / o F x 100, dove F = intensità della fluorescenza in un punto qualsiasi momento, Per = baseline fluorescenza). Quindi, utilizzare il programma Sigma Plot per generare la trama che mostra le variazioni di intensità di fluorescenza nel tempo. </li></ol> 7. Rappresentante Risultati La figura 1A mostra una immagine di fluorescenza di neuroni corticali di ratto incubate con TMRM. L&#39;aggiunta di FCCP, un disaccoppiatore mitocondriale, porta alla depolarizzazione mitocondriale e una perdita di intensità di fluorescenza TMRM (Fig. 1B). La linea di base a livello di fluorescenza TMRM rimane stabile prima dell&#39;aggiunta FCCP (i primi 350 secondi;. Fig. 1C). Analisi quantitativa di fluorescenza TMRM cambiamenti nel tempo mostra una significativa diminuzione della fluorescenza TMRM dopo l&#39;aggiunta di FCCP (Fig. 1C). 1D figura mostra l&#39;immagine di fluorescenza di topo neuroni corticali caricato con DCF. L&#39;aggiunta di H 2 O 2 si traduce in un aumento dell&#39;intensità della fluorescenza DCF in corpi cellulari (Fig. 1E). Il livello di base DCF fluorescenza è invariato (i primi 120 secondi) prima dell&#39;applicazione di H 2 O 2. Time-lapse misure di fluorescenza DCF mostrare i suoi livelli costanti, che aumentano dopo H 2 O 2 terapia (Fig. 1F). <img src="/files/ftp_upload/2704/2704fig1.jpg" alt="Figura 1"/> Figura 1. Valutazione del potenziale della membrana mitocondriale e dei livelli di ROS in neuroni corticali di ratto dal vivo. (A) Rappresentante immagine di fluorescenza di neuroni corticali caricato con TMRM. Dopo la scansione della linea di base TMRM fluorescenza, i neuroni sono stati trattati con il FCCP protonophore (1 mM). A destra è una barra pseudocolori intensità di fluorescenza TMRM con brillante massimo giallo e nero che rappresentano e l&#39;intensità minima, rispettivamente. La perdita di fluorescenza TMRM dalle regioni mitocondriale indica il crollo del ΔΨm su FCCP trattamento (pannello B). La rappresentazione quantitativa del cambiamento di intensità TMRM fluorescenza in diversi momenti, prima e dopo il trattamento FCCP è mostrato nel pannello C. (D) l&#39;immagine di fluorescenza di neuroni corticali di ratto caricato con H2DCF-DA. Dopo aver determinato la linea di base DCF fluorescenza, le cellule sono state trattate con 200 mM H 2 O 2, e la variazione del DCF fluorescenza è stata valutata. Un aumento della fluorescenza DCF riflette l&#39;aumento dei livelli di ROS su H 2 O 2 trattamento (E). L&#39;analisi quantitativa del cambiamento in DCF fluorescenza, prima e dopo H 2 O 2 trattamento, viene mostrato nella barra pannello F. Scala = 10 micron Video.7.1 - labmedia 2704_Joshi.avi Vivere l&#39;imaging cellulare di TMRM nei neuroni corticali, prima e dopo l&#39;aggiunta FCCP con obiettivo 40X. L&#39;intensità pseudocolori mostra un massimo (giallo brillante, prima aggiunta FCCP) e diminuito (colore rosso, dopo l&#39;aggiunta FCCP) intensità di fluorescenza TMRM dopo l&#39;aggiunta FCCP. <a href="http://www.jove.com/files/ftp_upload/2704/supplemental_video1.avi">Clicca qui per visualizzare il video</a> Video. 7.5 - labmedia 2704_Joshi.avi Vivere l&#39;imaging cellulare di DCF in neuroni corticali prima e dopo H 2 O 2 oltre utilizzando obiettivo 40X. La linea di base DCF fluorescenza ha colore verde chiaro in corpi cellulari e H2O2 Inoltre aumenta la DCF intensità della fluorescenza di colore verde brillante. <a href="http://www.jove.com/files/ftp_upload/2704/supplemental_video2.avi">Clicca qui per visualizzare il video</a>

<ol>
	<li>Hilgenberg, L. G., Smith, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+dissociated+mouse+cortical+neuron+cultures.">Preparation of dissociated mouse cortical neuron cultures.</a> J. Vis. Exp. , (2007).</li><li>Scaduto, R. C., Grotyohann, L. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+mitochondrial+membrane+potential+using+fluorescent+rhodamine+derivatives.">Measurement of mitochondrial membrane potential using fluorescent rhodamine derivatives.</a> Biophys. J. 76, 469-477 (1999).</li><li>Ward, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+amyloid+precursor+protein+intracellular+domain+(AICD)+disrupts+actin+dynamics+and+mitochondrial+bioenergetics.">The amyloid precursor protein intracellular domain (AICD) disrupts actin dynamics and mitochondrial bioenergetics.</a> J. Neurochem. 113, 275-284 (2010).</li><li>Gunasekar, P. G., Kanthasamy, A. G., Borowitz, J. L., Isom, G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NMDA+receptor+activation+produces+concurrent+generation+of+nitric+oxide+and+reactive+oxygen+species:+implication+for+cell+death.">NMDA receptor activation produces concurrent generation of nitric oxide and reactive oxygen species: implication for cell death.</a> J. Neurochem. 65, 2016-2021 (1995).</li></ol>

Nessun conflitto di interessi dichiarati.

Abbiamo presentato un passo-passo procedura che descrive come determinare ΔΨm e ROS in neuroni corticali di ratto utilizzando la fluorescenza indicatori TMRM e H 2 DCF-DA, rispettivamente. Per altri tipi di cellule, è importante per determinare empiricamente la concentrazione finale e il tempo di caricamento per TMRM o H 2 DCF-DA. In generale, la gamma TMRM concentrazioni di 20-200 nM, e il tempo di incubazione delle cellule con TMRM varia da 20 a 60 min. La concentrazione finale di H 2 DCF-DA range 2-10 mM, e l&#39;incubazione di cellule in soluzione tampone contenente tale indicatore varia da 30-45 min. E &#39;importante per ottimizzare la potenza del laser e velocità di scansione di prendere le immagini per evitare sia foto-tossicità per le cellule e le variazioni dell&#39;intensità di fluorescenza (ad esempio lo sfarfallio di TMRM fluorescenza), in assenza di qualsiasi stimolo. Le impostazioni ottimizzate ottico dovrebbe sfociare in un segnale di fluorescenza che non è sopra o sotto saturi (soglia) in assenza di stimolo. Le condizioni ottimali per raccogliere le immagini da un campo selezionato in una potenza del laser particolare e una velocità di scansione si ottengono quando non ci sono cambiamenti nella intensità di fluorescenza della sonda, in assenza di qualsiasi stimolo per 10-15 minuti di immagini dal vivo. Altre sonde a fluorescenza per determinare ΔΨm comprendono rodamina 123 e tetra metilico rodamina etile (TMRE). Tuttavia, essi sono stati trovati per inibire i processi respiratori nella isolata 2 mitocondri. Importante, TMRM non ha alcun effetto sulla respirazione mitocondriale a basse concentrazioni 2 e ha fototossicità bassa e photobleaching 3 rispetto con altre sonde. H 2 DCF-DA è un buon indicatore di ROS, come è ben conservato nelle cellule e riconosce diverse specie ossidanti, come i perossidi, ossidi super, e l&#39;ossido nitrico 4.

<table border="1"><tbody><tr><th> Nome del reagente </th><th> Azienda </th><th> Numero di catalogo </th></tr><tr><td> Con fondo di vetro piatto cultura </td><td> MatTek Corporation </td><td> P35G-1.5-14-C </td></tr><tr><td> NbActive4 </td><td> BrainBits </td><td> NbActive4 </td></tr><tr><td> TMRM </td><td> Invitrogen </td><td> T668 </td></tr><tr><td> H 2 DCF-DA </td><td> Invitrogen </td><td> C400 </td></tr><tr><td> NaCl </td><td> Sigma </td><td> S6191 </td></tr><tr><td> KCl </td><td> Sigma </td><td> P3911 </td></tr><tr><td> CaCl 2 • 2H 2 O </td><td> Sigma </td><td> C3306 </td></tr><tr><td> MgCl 2 • 6H 2 O </td><td> Sigma </td><td> M2670 </td></tr><tr><td> D-glucosio </td><td> Sigma </td><td> G6152 </td></tr><tr><td> HEPES </td><td> Invitrogen </td><td> 15630 </td></tr></tbody></table>

determination of mitochondrial membrane potential and reactive oxygen species in live rat cortical neurons

Potenziale di membrana mitocondriale (ΔΨm) è fondamentale per il mantenimento della funzione fisiologica della catena respiratoria per generare ATP. Una perdita significativa di cellule ΔΨm rende impoverito di energia con conseguente morte. Specie reattive dell&#39;ossigeno (ROS) sono importanti molecole di segnalazione, ma il loro accumulo in condizioni patologiche porta allo stress ossidativo. Le due principali fonti di ROS nelle cellule sono tossine ambientali e il processo di fosforilazione ossidativa. Disfunzione mitocondriale e lo stress ossidativo sono stati coinvolti nella fisiopatologia di molte malattie, quindi, la possibilità di determinare ΔΨm ROS e può fornire indizi importanti circa lo stato fisiologico della cellula e la funzione dei mitocondri. Diverse sonde fluorescenti (rodamina 123, TMRM, TMRE, JC-1) può essere utilizzato per determinare Δψm in una varietà di tipi di cellule, e molti indicatori di fluorescenza (Dihydroethidium, diidrorodamina 123, H 2 DCF-DA) può essere usato per determinare ROS . Quasi tutte le sonde a fluorescenza a disposizione per valutare ΔΨm o ROS sono singola lunghezza d&#39;onda indicatori, che aumentano o diminuiscono la loro intensità di fluorescenza proporzionale a uno stimolo che aumenta o diminuisce i livelli di ΔΨm o ROS. Quindi, è indispensabile per misurare l&#39;intensità di fluorescenza di queste sonde a livello basale e dopo l&#39;applicazione di uno stimolo specifico. Questo permette di determinare la percentuale di variazione di intensità di fluorescenza tra il livello base e uno stimolo. Questo cambiamento di intensità di fluorescenza riflette la variazione dei livelli relativi di ΔΨm o ROS. In questo video ci dimostra come applicare l&#39;indicatore di fluorescenza, TMRM, in neuroni corticali di ratto per determinare la variazione percentuale TMRM intensità di fluorescenza tra il livello basale e dopo l&#39;applicazione FCCP, un disaccoppiatore mitocondriale. I livelli più bassi di TMRM fluorescenza risultante dal trattamento FCCP riflettono la depolarizzazione del potenziale della membrana mitocondriale. Mostriamo anche come applicare la fluorescenza della sonda H 2 DCF-DA per valutare il livello di ROS in neuroni corticali, prima in condizioni basali e dopo l&#39;applicazione di H 2 O 2. Questo protocollo (con piccole modifiche) può essere utilizzato anche per determinare le variazioni ΔΨm e ROS in diversi tipi di cellule e nei neuroni isolati da altre regioni del cervello.

Watch this Scientific Journal Video about Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons at JoVE.com

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Dimostriamo applicazione del indicatore di fluorescenza, TMRM, nei neuroni corticali per determinare le variazioni relative a TMRM intensità di fluorescenza, prima e dopo l&#39;applicazione di uno stimolo specifico. Mostriamo anche l&#39;applicazione della sonda di fluorescenza H<sub&gt; 2</sub&gt; DCF-DA per valutare il livello relativo di specie reattive dell&#39;ossigeno nei neuroni corticali.

Determinazione della mitocondriale Oxygen Species potenziale di membrana e reattiva in neuroni corticali di ratto dal vivo

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

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Neuroscience

64.8K Views.  Loyola University Chicago. Dimostriamo applicazione del indicatore di fluorescenza, TMRM, nei neuroni corticali per determinare le variazioni relative a TMRM intensità di fluorescenza, prima e dopo l'applicazione di uno stimolo specifico. Mostriamo anche l'applicazione della sonda di fluorescenza H 2 DCF-DA per valutare il livello relativo di specie reattive dell'ossigeno nei neuroni corticali.

Video: Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

Neuromodulazione e mitocondriale di trasporto: Imaging Live in neuroni dell&#39;ippocampo oltre durate a lungo

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Ricerca

Neuroscienze

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

Analisi quantitativa di Replenishment pool delle vescicole sinaptiche in colture di neuroni granuli cerebellari utilizzando coloranti FM

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or co-culture model. This system is suitable for a variety of experimental needs requiring either a glass or plastic growth substrate and can also be used for culture of other types of neurons.
Rat cortical neurons obtained from the late embryonic stage (E17) are plated on glass coverslips or tissue culture dishes facing a feeder layer of glia grown on dishes or plastic coverslips (known as Thermanox), respectively. The choice between the two configurations depends on the specific experimental technique used, which may require, or not, that neurons are grown on glass (e.g. calcium imaging versus Western blot). The glial feeder layer, an astroglia-enriched secondary culture of mixed glia, is separately prepared from the cortices of newborn rat pups (P2-4) prior to the neuronal dissection.
A major advantage of this culture system as compared to a culture of neurons only is the support of neuronal growth, survival, and differentiation provided by trophic factors secreted from the glial feeder layer, which more accurately resembles the brain environment in vivo. Furthermore, the co-culture can be used to study neuronal-glial interactions1.
At the same time, glia contamination in the neuronal layer is prevented by different means (low density culture, addition of mitotic inhibitors, lack of serum and use of optimized culture medium) leading to a virtually pure neuronal layer, comparable to other established methods1-3. Neurons can be easily separated from the glial layer at any time during culture and used for different experimental applications ranging from electrophysiology4, cellular and molecular biology5-8, biochemistry5, imaging and microscopy4,6,7,9,10. The primary neurons extend axons and dendrites to form functional synapses11, a process which is not observed in neuronal cell lines, although some cell lines do extend processes.
A detailed protocol of culturing rat hippocampal neurons using this co-culture system has been described previously4,12,13. Here we detail a modified protocol suited for cortical neurons. As approximately 20x106 cells are recovered from each rat embryo, this method is particularly useful for experiments requiring large numbers of neurons (but not concerned about a highly homogenous neuronal population). The preparation of neurons and glia needs to be planned in a time-specific manner. We will provide the step-by-step protocol for culturing rat cortical neurons as well as culturing glial cells to support the neurons.

Here we provide a protocol for culturing rat cortical neurons in the presence of a glial feeder layer. The cultured neurons establish polarity and create synapses, and can be separated from the glia for use in various applications, such as electrophysiology, calcium imaging, cell survival assays, immunocytochemistry, and RNA/DNA/protein isolation.

Bilaminar co-coltura di neuroni corticali primari di ratto e Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

In Vivo a due fotoni di imaging dei cambi molecolari esperienza-dipendenti nei neuroni corticali

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Potenziale di membrana Dye Imaging di ventromediale dell&#39;ipotalamo neuroni da topi adulti per studiare glucosio Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

La quantificazione di filamentosa Actina (F-actina) Puncta nel ratto neuroni corticali

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

In Vivo Due-fotone Imaging dei neuroni corticali in topi neonatali

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using 128-channel high-density electroencephalography (EEG). The specific aim of the described stimuli and analyses is to examine whether it is feasible to replicate previously reported CVEP morphological patterns elicited by an apparent motion stimulus, designed to simultaneously stimulate both ventral and dorsal central visual networks, using object and motion stimuli designed to separately stimulate ventral and dorsal visual cortical networks.&#160; Four visual paradigms are presented: 1. Randomized visual objects with consistent temporal presentation. 2. Randomized visual objects with inconsistent temporal presentation (or jitter).&#160; 3. Visual motion via a radial field of coherent central dot motion without jitter.&#160; 4. Visual motion via a radial field of coherent central dot motion with jitter.&#160; These four paradigms are presented in a pseudo-randomized order for each participant.&#160; Jitter is introduced in order to view how possible anticipatory-related effects may affect the morphology of the object-onset and motion-onset CVEP response.&#160; EEG data analyses are described in detail, including steps of data exportation from and importation to signal processing platforms, bad channel identification&#160;and removal, artifact rejection, averaging, and categorization of average CVEP morphological pattern type based upon latency ranges of component peaks. Representative data show that the methodological approach is indeed sensitive in eliciting differential object-onset and motion-onset CVEP morphological patterns and may, therefore, be useful in addressing the larger research aim. Given the high temporal resolution of EEG and the possible application of high-density EEG in source localization analyses, this protocol is ideal for the investigation of distinct CVEP morphological patterns and the underlying neural mechanisms which generate these differential responses.&#160; &#160; &#160;&#160;

In this paper, we present a protocol to investigate differential cortical visual evoked potential morphological patterns through stimulation of ventral and dorsal networks using high-density EEG. Visual object and motion stimulus paradigms, with and without temporal jitter, are described. Visual evoked potential morphological analyses are also outlined.&#160;&#160;

Modelli morfologici potenziali evocati di tempo evocati da corticali specifici dello stimolo

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using ...

Generation of Neurospheres from Mixed Primary Hippocampal and Cortical Neurons Isolated from E14-E16 Sprague Dawley Rat Embryo

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have a robust in vitro model that can be exploited for various neurobiology studies. Though primary neuron cultures (i.e., long-term hippocampal cultures) have provided scientists with models, it does not yet represent the complexity of brain network completely. In the wake of these limitations, a new model has emerged using neurospheres, which bears a closer resemblance to the brain tissue. The present protocol describes the plating of high and low densities of mixed cortical and hippocampal neurons isolated from the embryo of embryonic day 14-16 Sprague Dawley rats. This allows for the generation of neurospheres and long-term primary neuron culture as two independent platforms to conduct further studies. This process is extremely simple and cost-effective, as it minimizes several steps and reagents previously deemed essential for neuron culture. This is a robust protocol with minimal requirements that can be performed with achievable results and further used for a diversity of studies related to neuroscience.

Presented here is a protocol for the spontaneous generation of neurospheres enriched in neural progenitor cells from high density plated neurons. During the same experiment, when neurons are plated at a lower density, the protocol also results in prolonged primary rat neuron cultures.

Generazione di neurosfere da neuroni ippocampali primari misti e corticali isolati da E14-E16 Sprague Dawley Rat Embryo

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have ...

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. In addition, photoreceptors also exhibit high aerobic glycolysis like cancer cells. Precise measurements of these metabolic activities can provide valuable insights into cellular homeostasis under physiological conditions and in disease states. High throughput microplate-based assays have been developed to measure mitochondrial respiration and various metabolic activities in live cells. However, a vast majority of these are developed for cultured cells and have not been optimized for intact tissue samples and for application ex vivo. Described here is a detailed step-by-step protocol, using microplate-based fluorescence technology, to directly measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration, as well as extracellular acidification rate (ECAR) as an indicator of glycolysis, in intact ex vivo retinal tissue. This method has been used to successfully assess metabolic activities in adult mouse retina and demonstrate its application in investigating cellular mechanisms of aging and disease.

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Determinazione della respirazione mitocondriale e della glicolisi in campioni di tessuto retinico Ex Vivo

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. ...