Ce travail a été soutenu par le National Institutes of Health (K22NS050137 à JCB).

1. La culture cellulaire <ol><li> Les neurones corticaux sont isolées et cultivées en utilisant des techniques décrites précédemment et étalées sur des boîtes de culture avec un fond de verre (MatTek Corporation, Ashland, MA) recouvertes de poly-D-lysine et de la laminine 1. </li></ol> 2. Préparer les solutions mères pour les sondes fluorescentes TMRM et H 2 DCF-DA <ol><li> Préparer une solution stock de 10 mM de TMRM en dissolvant 5,0 mg de TMRM dans 1 ml de diméthylsulfoxyde anhydre. Vortex c&#39;est pendant 1 min. Ensuite, faire des aliquots et de les stocker à -20 ° C, protéger de la lumière, et l&#39;utiliser dans un mois. </li><li> Ensuite, préparer une solution stock de 10 mM de H 2 DCF-DA en dissolvant 4,87 mg de H 2 DCF-DA dans 1 ml de DMSO anhydre. De même, il vortex pendant 1 min. Ensuite, faire des aliquots et de les stocker à -20 ° C, protéger de la lumière, et l&#39;utiliser dans une semaine. </li></ol> 3. Chargement des neurones de rat corticale avec TMRM et H 2 DCF-DA TMRM est un potentiométrique, les cellules perméables indicateur fluorescent qui s&#39;accumule dans l&#39;intérieur, fortement chargées négativement des mitochondries. Il est important d&#39;utiliser les faibles concentrations (10-50 nm) des TMRM pour éviter l&#39;auto-extinction de TMRM mitochondriale. Ensuite, le signal de fluorescence de TMRM peuvent être directement liés à la co-ΔΨm à travers la membrane mitochondriale interne. Une perte de ΔΨm provoque une fuite de TMRM des mitochondries entraînant une perte d&#39;intensité de fluorescence. H 2 DCF-DA est la cellule-perméable sonde converti en signal DCF-DA par des estérases intracellulaires, et ses résultats d&#39;oxydation dans DCF fluorescente. La concentration finale de H 2 DCF-DA varie entre 2-10 uM et il devrait être testé empiriquement dans les neurones issus de différentes régions du cerveau car les concentrations de chargement élevée peut entraîner la saturation du DCF de fluorescence, même en l&#39;absence de H 2 O 2. La présence de tout oxydant endogène ou exogène (par exemple, l&#39;hydrogène hypoazotique oxyde) va augmenter DFC de fluorescence d&#39;intensité. Ci-dessous, nous fournissons un protocole pour le chargement de neurones corticaux de rat avec TMRM et H 2 DCF-DA. <ol><li> Pour charger les neurones corticaux de rat avec TMRM, d&#39;abord, lavez les neurones cultivés à 3 reprises avec du tampon de Tyrode (Superposition de texte: la tuberculose: 145 mM NaCl, 5 mM de KCl, 10 mM de glucose, 1,5 mM de CaCl2, MgCl2 1 mM, et 10 mM HEPES; ajuster le pH à 7,4 avec NaOH). Ensuite, préparez 20 nM de TMRM en diluant le stock de 10 mM TMRM 1 / 1000 fois dans la tuberculose, puis ajouter 2 pi de TMRM dilué par 1 ml de la tuberculose. Incuber les neurones avec TMRM pendant 45 min dans le noir à température ambiante. Après 45 minutes, monter la boîte de culture sur la scène du microscope et de commencer l&#39;imagerie. </li><li> Pour charger les neurones corticaux de rat avec du H 2 DCF-DA, laver les neurones cultivés 3 fois avec la tuberculose. Ensuite, préparez 2 uM de H 2 DCF-DA en diluant le 10 mM H 2 DCF-DA stock de 1 / 10 fois dans la tuberculose, puis ajouter 2 pi de dilution de H 2 DCF-DA pour 1 ml de la tuberculose. Puis, incuber les neurones avec du H 2 DCF-DA pendant 45 min dans le noir à température ambiante. Après 45 minutes, laver les neurones 4 fois avec la tuberculose pour enlever l&#39;excès indicateur fluorescent, avant d&#39;obtenir des images. </li></ol> 4. En direct d&#39;imagerie des neurones incubées avec TMRM pour déterminer ΔΨm <ol><li> Pour effectuer l&#39;imagerie en direct de neurones incubées avec TMRM, microscopie confocale à balayage laser (Superposition de texte: LSM 510, objectif Carl Zeiss Inc), avec l&#39;application de vivre des séries chronologiques de programme, est utilisé. Appliquer à faible résolution et atténué la puissance laser (Superposition de texte: basse résolution: 256 x 256; puissance du laser: 1%) pour minimiser le temps nécessaire pour obtenir des images et d&#39;éviter de photoblanchiment. </li><li> . Ensuite, régler la netteté de l&#39;neurones montés chargé avec TMRM utilisant la lumière réfléchie. Examiner la fluorescence TMRM par illumination à 514 nm et la détection à 570 nm. Réglez le gain de détection d&#39;une caméra juste en dessous du niveau de saturation. </li><li> Une fois tous les paramètres qui incluent la résolution, la puissance du laser, le gain de détection d&#39;une caméra, et time-lapse d&#39;intervalle pour obtenir des images sont définies, ne pas modifier ces paramètres entre les expériences. Ensuite, modifiez le champ. Commencer à recueillir des images. </li><li> Pour tester des changements dans ΔΨm, des stimuli tels que 1 uM de FCCP ou 2 ug / ml de oligomycine, peut être appliquée, ce qui va considérablement dépolariser ou hyperpolariser le potentiel de membrane mitochondriale, respectivement. Ces changements seront reflétés par une diminution de l&#39;intensité de fluorescence TMRM par rapport à l&#39;intensité de la fluorescence de base dans le cas du FCCP, ou une augmentation de TMRM intensité de fluorescence dans le cas de oligomycine. </li></ol> 5. Imagerie en direct de neurones incubées avec H 2 DCF-DA ROS pour déterminer <ol><li> Pour effectuer l&#39;imagerie en direct de neurones incubées avec H 2 DCF-DA, d&#39;abord, monter la boîte de culture sur la scène d&#39;un microscope. Ajustez la mise au point des cellules-nousING lumière réfléchie. Examinez DCF de fluorescence par excitation à 488 nm et émission à 515 nm. </li><li> Ensuite, régler la puissance du laser à 5-7%, le gain du détecteur, et la résolution de 256 x 256. Ne pas modifier ces paramètres entre les expériences. Ensuite, définissez la fréquence pour obtenir des images en direct en utilisant le programme de séries chronologiques. </li><li> Sélectionner un domaine nouveau et commencer à acquérir des images. Pour détecter les changements dans les niveaux de ROS, traiter avec des cellules 100-200 pm de H 2 O 2. Cela sera reflété par une augmentation de l&#39;intensité de fluorescence DCF par rapport au niveau de base. </li></ol> 6. L&#39;analyse des données <ol><li> Utilisez région d&#39;intérêt (Superposition de texte: RCI) l&#39;outil à partir du programme LSM pour sélectionner les domaines. Ensuite, mesurer la TMRM ou ROS intensités de fluorescence. Sélectionnez ROIs de régions mitochondrial ou ROIs du corps cellulaire dans les cellules de toute imagée pour mesurer les intensités de fluorescence de TMRM ou ROS, respectivement. </li><li> Calculer les intensités moyennes de fluorescence de tous les ROIs de chaque cellule pour TMRM ou d&#39;organismes à cellules entières pour toutes les cellules imagé pour ROS pour chaque point de temps. Sélectionnez les régions à côté de la cellules pour calculer l&#39;intensité de fluorescence de fond. Prenez plusieurs mesures et de calculer l&#39;intensité de fond moyenne. </li><li> Soustraire l&#39;intensité de fluorescence de fond moyen de la moyenne des intensités de fluorescence de ROIs dans chaque cellule de chaque point de temps en utilisant Microsoft Excel. Après soustraction de l&#39;intensité de fond, de normaliser l&#39;intensité de fluorescence TMRM ou DCF à la fluorescence de base en utilisant cette formule (Superposition de texte: AF = FF o / o F x 100, où F = intensité de fluorescence à aucun moment, Fo = base de fluorescence). Ensuite, utilisez le programme tracé Sigma pour générer le graphique montrant les changements d&#39;intensité de fluorescence au cours du temps. </li></ol> 7. Les résultats représentatifs La figure 1A montre une image de fluorescence des neurones corticaux de rat incubées avec TMRM. Ajout de FCCP, un découplant mitochondrial, conduit à une dépolarisation mitochondriale et une perte d&#39;intensité de fluorescence TMRM (Fig. 1B). Le niveau de fluorescence de base TMRM reste stable avant l&#39;addition du FCCP (le premier 350 sec;. Fig 1C). L&#39;analyse quantitative des changements de fluorescence TMRM au fil du temps montre une diminution significative de la fluorescence après ajout de TMRM FCCP (Fig. 1C). La figure 1D montre l&#39;image de fluorescence des neurones corticaux de rats chargé avec DCF. L&#39;addition de H 2 O 2 résultats dans DCF augmenté l&#39;intensité de fluorescence dans les corps cellulaires (figure 1E). La base de fluorescence DCF niveau est inchangé (le premier 120 sec) avant l&#39;application de H 2 O 2. Time-lapse mesures de fluorescence montrent DCF ses niveaux stables, ce qui augmente après H 2 O 2 de traitement (Fig. 1F). <img src="/files/ftp_upload/2704/2704fig1.jpg" alt="Figure 1"/> Figure 1. Évaluation du potentiel de membrane mitochondriale et les niveaux de ROS dans des neurones corticaux rat vivant. (A) représentant l&#39;image de fluorescence des neurones corticaux chargés avec TMRM. Après la numérisation de la base TMRM de fluorescence, les neurones ont été traités avec le FCCP protonophore (1 uM). A droite se trouve une barre d&#39;intensité de fluorescence pseudo TMRM vif maximal représentant jaune et noir et une intensité minimale, respectivement. La perte de fluorescence TMRM des régions mitochondrial indique l&#39;effondrement du FCCP ΔΨm sur le traitement (tableau B). La représentation quantitative des changements dans l&#39;intensité de fluorescence TMRM à différents moments avant et après traitement FCCP est montré dans le panneau C (D) image de fluorescence des neurones corticaux de rat chargé avec H2DCF-DA. Après détermination de la base DCF de fluorescence, les cellules ont été traitées avec 200 uM H 2 O 2, et le changement de DCF de fluorescence a été évaluée. Une augmentation de la fluorescence DCF reflète l&#39;augmentation des niveaux de ROS lors de H 2 O 2 de traitement (E). L&#39;analyse quantitative des changements dans la fluorescence DCF, avant et après H 2 O 2 de traitement, est montré dans le panneau de F. Échelle bar = 10 um Video.7.1 - labmedia 2704_Joshi.avi Direct imagerie cellulaire des neurones corticaux dans TMRM avant et après addition FCCP utilisant l&#39;objectif 40X. L&#39;intensité pseudo montre un maximum (jaune vif, avant l&#39;addition FCCP) et a diminué (de couleur rouge, après addition FCCP) TMRM intensité de fluorescence après ajout FCCP. <a href="http://www.jove.com/files/ftp_upload/2704/supplemental_video1.avi">Cliquez ici pour voir la vidéo</a> Vidéo. 7.5 - labmedia 2704_Joshi.avi Direct imagerie cellulaire du DCF dans les neurones corticaux, avant et après H 2 O 2 en utilisant plus objectif 40X. La base de fluorescence a DCF lumière de couleur verte dans les corps cellulaires et H2O2 ajout augmente le DCF l&#39;intensité de fluorescence à la couleur vert vif. <a href="http://www.jove.com/files/ftp_upload/2704/supplemental_video2.avi">Cliquez ici pour voir la vidéo</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>

Nous avons présenté une procédure étape par étape décrivant comment déterminer ΔΨm et ROS dans les neurones corticaux de rat en utilisant les indicateurs fluorescents TMRM et H 2 DCF-DA, respectivement. Pour d&#39;autres types cellulaires, il est important de déterminer empiriquement la concentration finale et le temps de chargement pour TMRM ou H 2 DCF-DA. En général, la gamme des concentrations de TMRM 20-200 nm, et le temps d&#39;incubation des cellules avec des TMRM varie de 20 à 60 min. La concentration finale de H 2 DCF-DA varie de 2 à 10 uM, et l&#39;incubation des cellules dans une solution de chargement contenant cet indicateur varie de 30-45 min. Il est important pour optimiser la puissance du laser et la vitesse de balayage de prendre les images pour éviter à la fois photo-toxicité pour les cellules et les changements dans l&#39;intensité de fluorescence (par exemple vacillante de TMRM fluorescence) en l&#39;absence de tout stimulus. Les paramètres optimisés optiques devrait se traduire par un signal de fluorescence qui n&#39;est pas plus ou moins saturés (seuil) en l&#39;absence de stimulus. Les conditions optimales pour recueillir les images à partir d&#39;un champ sélectionné à un laser de puissance particulière et une vitesse de balayage sont obtenus lorsque il n&#39;ya pas de changements dans l&#39;intensité de fluorescence de la sonde en l&#39;absence de tout stimulus pendant 10-15 min d&#39;imagerie en temps réel. Autres sondes de fluorescence pour déterminer ΔΨm comprennent la rhodamine 123 et tétra méthyl ester éthylique de la rhodamine (TMRE). Cependant, ils ont été trouvés pour inhiber les processus respiratoires en 2 mitochondries isolées. Surtout, TMRM n&#39;a aucun effet sur ​​la respiration mitochondriale à de faibles concentrations 2 et a phototoxicité faible et photoblanchiment 3 par rapport à d&#39;autres sondes. H 2 DCF-DA est un bon indicateur de ROS comme elle est bien conservée dans les cellules et reconnaît espèces oxydantes de plusieurs, tels que les peroxydes, oxydes super, et l&#39;oxyde nitrique 4.

<table border="1"><tbody><tr><th> Nom de réactif </th><th> Société </th><th> Numéro de catalogue </th></tr><tr><td> Boîte de culture à fond de verre </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-glucose </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

Potentiel de membrane mitochondriale (ΔΨm) est essentiel pour maintenir la fonction physiologique de la chaîne respiratoire pour produire l&#39;ATP. Une perte importante de ΔΨm rend les cellules épuisées d&#39;énergie avec la mort subséquente. Espèces réactives de l&#39;oxygène (ERO) sont importantes molécules de signalisation, mais leur accumulation dans des conditions pathologiques conduit au stress oxydatif. Les deux principales sources de ROS dans les cellules sont des toxines environnementales et le processus de phosphorylation oxydative. Dysfonction mitochondriale et le stress oxydatif ont été impliqués dans la physiopathologie de nombreuses maladies et, par conséquent, la capacité de déterminer ΔΨm et ROS peuvent fournir des indices importants sur l&#39;état physiologique de la cellule et la fonction des mitochondries. Plusieurs sondes fluorescentes (Rhodamine 123, TMRM, TMRE, JC-1) peut être utilisée pour déterminer Δψm dans une variété de types de cellules, et de nombreux indicateurs de fluorescence (Dihydroethidium, dihydrorhodamine 123, H 2 DCF-DA) peut être utilisée pour déterminer les ROS . Presque toutes les sondes de fluorescence disponibles utilisés pour évaluer ΔΨm ou ROS sont une seule longueur d&#39;onde des indicateurs, qui augmentent ou diminuent leur intensité de fluorescence proportionnelle à un stimulus qui augmente ou diminue les niveaux de ΔΨm ou ROS. Ainsi, il est impératif de mesurer l&#39;intensité de fluorescence de ces sondes au niveau basal et après l&#39;application d&#39;un stimulus spécifique. Cela permet de déterminer le pourcentage de changement dans l&#39;intensité de fluorescence entre le niveau de référence et un stimulant. Ce changement dans l&#39;intensité de fluorescence reflète le changement dans les niveaux relatifs de ΔΨm ou ROS. Dans cette vidéo, nous démontrons comment appliquer l&#39;indicateur de fluorescence, TMRM, chez le rat neurones corticaux de déterminer le pourcentage de variation de l&#39;intensité de fluorescence TMRM entre le niveau de base et après l&#39;application FCCP, un découplant mitochondrial. Les niveaux inférieurs de la fluorescence TMRM résultant du traitement FCCP refléter la dépolarisation du potentiel de membrane mitochondriale. Nous montrons aussi comment appliquer la fluorescence de sonde H 2 DCF-DA pour évaluer le niveau de ROS dans des neurones corticaux, d&#39;abord au départ et puis après application de H 2 O 2. Ce protocole (avec des modifications mineures) peuvent être également utilisés pour déterminer les changements dans ΔΨm et ROS dans différents types cellulaires et dans les neurones isolés à partir d&#39;autres régions cérébrales.

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

Nous démontrons l&#39;application de l&#39;indicateur de fluorescence, TMRM, dans les neurones corticaux de déterminer les changements relatifs d&#39;intensité de fluorescence TMRM avant et après application d&#39;un stimulus spécifique. Nous montrons aussi l&#39;application de la fluorescence de la sonde H<sub&gt; 2</sub&gt; DCF-DA pour évaluer le niveau relatif des espèces réactives de l&#39;oxygène dans les neurones corticaux.

Détermination des espèces d&#39;oxygène Potentiel membrane mitochondriale et réactive dans les neurones corticaux de rat en direct

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

determination-mitochondrial-membrane-potential-reactive-oxygen

Research

JoVE Journal

Neuroscience

64.7K vues.  Loyola University Chicago. Nous démontrons l'application de l'indicateur de fluorescence, TMRM, dans les neurones corticaux de déterminer les changements relatifs d'intensité de fluorescence TMRM avant et après application d'un stimulus spécifique. Nous montrons aussi l'application de la fluorescence de la sonde H 2 DCF-DA pour évaluer le niveau relatif des espèces réactives de l'oxygène dans les neurones corticaux.

Vidéo: 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.

Neuromodulation et mitochondriale de transport: Live in Imaging neurones de l&#39;hippocampe sur des durées longues

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

Recherche

Neurosciences

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.

Analyse quantitative des vésicules reconstitution Piscine synaptique dans des cultures de neurones granulaires du cervelet utilisant des colorants 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.

Didermique co-culture de neurones de rat corticale primaire et des cellules gliales

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 Imagerie biphotonique d&#39;expérience qui dépendent des changements moléculaires dans les neurones corticaux

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.

Potentiel de membrane Dye Imaging de ventromédian hypothalamus neurones de souris adulte à étudier 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 ...

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.

Quantification des filamenteuses Actine (F-actine) punctas chez le rat neurones corticaux

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 Two-photon Imaging de neurones corticaux en souris néonatales

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;

Modèles morphologiques potentiels d'evoked Visual

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.

Génération de neurosphères à partir de neurones hippocampes et corticales primaires mixtes isolées de l'embryon de rat Sprague Dawley E14-E16

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.

Détermination de la respiration mitochondriale et de la glycolyse dans des échantillons de tissu rétinien ex vivo

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