Ringraziamo il Dr. Yong-Ling Zhu per fornire lo synaptophysin-pHluorin2x costrutto e Dr. James E. Rothman per fornire VAMP2-phluorin. Ringraziamo Dr. Susan Cheng e Virginia Crocker di NINDS microscopia elettronica Facility per loro guida e supporto tecnico. Questo lavoro è stato supportato dall'Istituto nazionale dei disordini neurologici e colpo programma di ricerca intramurale in USA e una sovvenzione dal programma iniziativa ricerca KRIBB (coreano Biomedical Scientist Fellowship Program), Korea Research Institute di Bioscienze e biotecnologie, Repubblica di Corea.

Nota: il seguente protocollo descrive i metodi di formazione immagine di pHluorin e metodi EM utilizzati in neuroni hippocampal coltivati. pHluorin monitor delle vescicole sinaptiche proteine l'assorbimento nelle cellule viventi ed EM rileva l'assorbimento delle vescicole sinaptiche e cambiamenti ultrastrutturali.
 cura degli animali e procedura seguite linee guida NIH e sono stati approvati dal comitato di uso e cura degli animali NIH.
 1. pHluorin Imaging <ol><li>cultura Neurone ippocampale <ol><li>preparare ippocampo Buffer (HB) combinando 4 NaHCO 3 e 5 mM HEPES e regolare a pH 7,3 con 5 M NaOH. Fare il terreno di coltura mescolando neurobasal medi, 2% B27, 0,5 mM L-Glutammina e 1% di penicillina-streptomicina. Inoltre, preparare una miscela di tampone HB con 20% di siero bovino fetale (FBS HB/20%). 
		Nota: Il presente protocollo coltura si basa su Sankaranarayanan, et al. 29 e Wu et al. 24</li> <li>decapitare i cuccioli del mouse tra postnatale giorno 0 al giorno 2 in terreno di coltura ed estrarre il cervello nella 4 ° C HB/20% FBS. Rimuovere il tronco cerebrale ed il talamo per esporre ippocampi. Sezionare ippocampi, dopo esporla tramite rimozione del tronco cerebrale e nel talamo e trasferire al fresco 4 ° C HB/20% FBS. 
		Nota: In genere, la resa è circa 4 x 10 5 cellule/mL per un cucciolo (due ippocampi). Pulire le membrane aderente con una pinzetta o forbici, poi il trasferimento a fresco 4 ° C HB/20% FBS. Srotolare la circonvoluzione dentata e isolare il subiculum tagliando con le forbici, poi trasferimento a fresco 4 ° C HB/20% FBS.</li>
		<li>Dividere ogni ippocampo, dal taglio dall'inizio alla fine, a circa 10 fette e trasferirli in un tubo conico in polipropilene 15ml.</li>
		<li>Dopo che permette al tessuto di stabilirsi, lavare con 10 mL di HB/20% FB e poi lavare tre volte con 10ml di HB.</li>
		<li>Preparare la soluzione di digestione di 137 mM NaCl, 5 mM KCl, 7mm Na2HPO 4 e 25 mM HEPES e regolare il pH a 7,2 con NaOH M 5. Rimuovere il surnatante da ippocampi. Aggiungere 2 mL di soluzione di digestione 10 mg di tripsina e 1 mg di DNasi e filtrare attraverso una membrana da 0,22 µm sterile direttamente sul pellet campione.</li>
		<li>Incubare ippocampi per 5 min a 37 ° C, quindi lavate due volte con 10 mL di HB/20% FB e poi lavare una volta con 10 mL di HB.</li>
		<li>Dissociare le cellule con 6 mg di MgSO 4 &#8729; 7 H 2 O e 1 mg di dnasi a 2 mL HB e filtro sterile sul ippocampi a pellet attraverso un filtro da 0,22 µm sterile. Delicatamente dissociare le cellule mediante triturazione attenzione, avendo cura di evitare di introdurre bolle d'aria. Consentire le particelle di tessuto a stabilirsi per 2 min e poi lentamente trasferire il surnatante in un'altra provetta.</li>
		<li>Aggiungere 3 mL HB/20% FBS alla sospensione cellulare e centrifuga o 10 min a 4 ° C e 1000 giri/min. Scartare il surnatante e risospendere in terreno di coltura.</li>
		<li>Piastra 60.000 cellule sospese nel 150 µ l di coltura su un vetrino coprioggetti 25 mm diametro di poli-D-lisina rivestito, senza fuoriuscita fuori il vetrino coprioggetti. Aggiungere 2 mL di terreno di coltura pre-riscaldato 2 h dopo il placcaggio. Le lamelle sono mantenute in coltura piastre da 6 pozzetti o Petri sterili.</li>
		<li>Cellule di mantenere a 37 ° C in un 5% CO 2 umidificati incubatore in terreno di coltura per 14-21 giorni prima della registrazione. Durante la crescita di cultura, cambiare la metà superiore del mezzo due volte a settimana.</li>
	</ol></li> <li>Transfezione <ol><li>6-7 giorni dopo il placcaggio, transfect synaptophysin-pHluorin 2 X (SpH) o VAMP2-pHluorin in neuroni ippocampali. Per SpH, utilizzare un promotore del citomegalovirus (CMV), inserito in un di vettore pcDNA3 30. Per VAMP2-pHluorin, utilizzare un promotore CMV inserito in un vettore di pCI 31.</li>
		<li>Transfect entrambi vector di elemento portante del lipido in cellule bersaglio, usando 1 µ g del plasmide. Utilizzare il terreno di coltura dal passaggio di protocollo 1.1.1, che manca del siero, per la transfezione. Modificare il mezzo 2 h dopo la trasfezione per ridurre la tossicità. 
		Nota: In caso di bassa espressione di SpH nel boutons, aumentare la concentrazione di DNA 2 µ g o il tempo di incubazione con il vettore del lipido, a meno che le cellule sono malsane. In genere, 4-10 celle (0,006-0,008%) dei neuroni transfected.</li>
	</ol></li> <li>Microscopia <ol><li>preparare soluzione salina normale è composto da 119 mM NaCl, 2mm CaCl 2, 2,5 mM KCl, 25mm HEPES (pH 7.4), glucosio di 30 mM, 2 mM MgCl 2, 0,01 mM (6-cyano-7-nitroquinoxaline-2,3-dione CNQX) e 0,05 mM acido DL-2-ammino-5-phosphonovaleric (AP5). Prendere un vetrino coprioggetto dalla piastra di terreno di coltura e posto su una camera di imaging che permette una stimolazione di campo, con lubrificante e sigillante per evitare perdite. Evitare di lasciare che il vetro asciutto durante il trasferimento di un vetrino coprioggetto dalla piastra alla camera immediatamente aggiungendo 750 µ l di soluzione salina normale. 
		Nota: CNQX e AP5 sono stati utilizzati per bloccare l'attività post-sinaptica, che ha il potenziale per attività ricorrente. Prima di mettere una camera su un microscopio a fluorescenza invertito, è possibile utilizzare tessuto di pulizia per confermare che la camera non ci siano perdite. Colatura lento causa i cambiamenti di messa a fuoco durante la registrazione dalla miscelazione di normale soluzione salina e immersione olio, che si traduce in cambiamenti nell'indice di rifrazione.</li>
		<li>Stimolazione e registrazione d'immersione in olio (apertura numerica 1,4) <ol><li>immagine su un microscopio invertito widefield con un 60 X lente con una lampada ad alogenuri metallici. Visualizzare pHluorin con un filtro impostato per un picco di eccitazione di 480 nm, uno specchio di passaggio lungo 490 nm, un filtro di emissione di 500-550 nm e un otturatore manuale flip. Catturare immagini ogni 100 ms, con 2 x 2 binning, utilizzando una telecamera di elettrone moltiplicando Charge Coupled dispositivo (EMCCD). 
			Nota: Molte funzionalità dovrebbe essere considerata per raggiungere le giuste condizioni per evitare il photobleaching, compreso l'intervallo di acquisizione immagine e filtrare e binning, che sono dipendenti da attrezzature. Nel caso di imaging confocale, laser potenza e tempo di esposizione dovrebbe essere considerato per evitare photobleaching. L'installazione è stata determinata dopo aver registrato almeno 3 min senza stimolo e la registrazione è stata effettuata per almeno 10 s senza stimolo per cercare photobleaching.</li>
			<li>Scegliere una zona con un'alta densità dei boutons per facilitare l'analisi in ogni esperimento. Identificare cellule trasfettate dal loro segnale di fluorescenza verde, con enfasi sui modelli di espressione rotonda o ovale nelle bouton ed espressione continua tra boutons.</li>
			<li>Induce endocitosi di stimolazione di campo utilizzando un 1 impulso ms, 20 mA potenziale d'azione (AP) fornito da uno stimolatore di impulso e trasportato attraverso un elettrodo di platino all'interno dell'unità di isolamento di stimolo. Attività fluorescenza della battuta nel corso della stimolazione e durante il recupero delle cellule. 
			Nota: Nel caso di cellule morte, espressione era più forte di vita boutons e ha mostrato un modello irregolare tra boutons.</li>
		</ol></li></ol></li> <li>Analisi dell'immagine <ol><li>Per analizzare l'intensità di fluorescenza in una singola bouton, impostato su una regione di interesse (ROI) come un 1,5 x 1,5 µm quadrati; la dimensione di una bouton è all'interno di circa 1,5 µm. uso le tracce prima della stimolazione per controllare photobleaching e sottrarre come sfondo.</li>
		<li>Normalizza fluorescenza cambiare (ΔF) con l'equazione: <img alt="Equation" height="11" src="/files/ftp_upload/55862/55862eq1.jpg" width="172" /> dove F max e F 0 si riferiscono al massimo incremento dopo stimolazione e fluorescenza della linea di base, rispettivamente. Misurare endocitosi tassi come il tasso di decadimento durante i primi 4-10 s dopo il punto massimo di fluorescenza di pHluorin. Ottenere la costante di tempo (τ) dell'endocitosi inserendo il deperimento di fluorescenza di pHluorin dall'aumento del picco alla linea di base con una funzione mono-esponenziale.</li>
	</ol></li></ol> 2. Microscopia elettronica <ol><li>preparare una poli-D-lisina 6-pozzetti piatto rivestito di applicazione 1,5 mL di soluzione di poli-D-lisina filtrato sterile 0,01% ad ogni pozzetto per 1 h a temperatura ambiente, poi lavare tre volte con acqua sterilizzata. Sezionare, cultura e mantenere i neuroni hippocampal come punti 1.1.1, 1.1.2 e 1.1.3, rispettivamente.</li>
	<li>Preparare una soluzione di stimolazione elevata K + con HRP come 31,5 mM NaCl, 2mm CaCl 2, 90 mM KCl, 25mm HEPES (pH 7.4), glucosio di 30 mM, 2 mM MgCl 2, 0,01 mM CNQX, 0,05 mM AP5 e 5 mg/mL HRP, quindi regolare il pH 7.4 con 5 M NaOH.
	Soluzione di stimolazione K + <ol><li>stimolare la cultura del neurone ippocampale con 1,5 mL alto a temperatura ambiente (denominato K +) tramite l'aggiunta di 1,5 mL per ciascun pozzetto per 90 s. In condizione di riposa (denominata R), applicare la stessa concentrazione di 5 mg/mL HRP per 90 s, ma con normale soluzione salina. Per l'esempio di recupero, applicare la soluzione di stimolazione elevata K + come con l'esempio K +, poi rapidamente lavare e sostituire con normale soluzione salina e incubare per 10 min.</li>
	</ol></li> <li>Fissazione e colorazione <ol><li>preparare tampone cacodilato Na di 0.1 M utilizzando 21,4 g/L Na cacodylate a pH 7,4. Fissare le cellule con glutaraldeide al 4% in tampone cacodilato di 0.1 M Na per almeno 1 h a temperatura ambiente. Lavare tre volte con tampone cacodilato Na di 0.1 M per 7 minuti ciascuno,.</li>
		<li>Soluzione di preparare diaminobenzidina (DAB), composto da 0,5 mg/mL di tamponare con 0,3% H 2 O 2 ddH 2 O, ed il filtro con un filtro di 0,22 µm. Applicare 1,5 mL di soluzione DAB per 30 min a 37 ° C. lavare tre volte con tampone cacodilato Na di 0.1 M per 7 minuti ciascuno,. 
		Attenzione: DAB è un tossico e sospetto cancerogeno. Si prega di utilizzare guanti e camici da laboratorio. 
		Nota: Etichettatura con DAB si verifica a causa della sua ossidazione di H 2 O 2, come catalizzata da HRP. Piccoli aumenti nel risultato di componenti in un segnale aumentato nel campione. Aumento della concentrazione di HRP accelera l'effetto del catalizzatore. Sufficientemente grandi concentrazioni di H 2 O 2 consentono debilitante reazioni secondarie con HRP, inibendo l'effetto dell'etichettatura 32. In questo lavoro, le concentrazioni di questo sistema di etichettatura sono stati scelti in base alla ricerca attualmente disponibili 33 , 34.</li>
		<li>Incubare i neuroni con 1,5 mL di 1% OsO 4 in tampone cacodilato Na di 0.1 M per 1 h a 4 ° C, come la fissazione della posta. Lavare tre volte con 1,5 mL di tampone cacodilato Na di 0.1 M per 7 minuti ciascuno,. 
		Attenzione: A causa della tossicità e la reattività di OsO 4, mantenendo il campione sul ghiaccio in una cappa chimica è preferibile in molti casi utilizzando un frigo per l'incubazione.</li>
		<li>Preparare 0.1 M tampone acetato di sodio con 13,61 g/L sodio acetato e 11,43 mL/L acido acetico glaciale a pH 5.0. Lavare tre volte con tampone di acetato 1,5 mL 0,1 M a pH 5.0 per 7 minuti ciascuno e incubare con acetato di uranile 1,5 mL 1% in tampone acetato 0,1 M a pH 5.0 per 1 h a 4 ° C. Wash tre volte con tampone di acetato 0,1 M 1,5 mL per 7 minuti ciascuno,.</li>
	</ol></li> <li>Epossidica embedding <ol><li>disidratare la cultura del neurone con lavaggi di singolo 1,5 mL di etanolo 50%, 70% e 90%, 7 min in ciascuno e poi 3 lavaggi di 1,5 mL di etanolo al 100% per 7 minuti ciascuno, in una cappa aspirante.</li>
		<li>Mix 485 mL/L bisphenol-A-(epichlorhydrin), resina di anidride di 160 mL/L dodecenyl succinico (DDSA), 340 mL/L metil-5-Norbornene-2,3-dicarbossilico anidride acetica (NMA) e 15 mL/L 2, 4,6-tris (dimethylaminomethyl) fenolo (DMP-30) per creare la resina epossidica resina. Mix accuratamente, quindi memorizzare sotto vuoto per eliminare le bolle d'aria. 
		Nota: È fondamentale per rimuovere le bolle d'aria nella resina, soprattutto quelli inferiori visibili ad occhio nudo, perché possono causare cavità nella resina durante il sezionamento.</li>
		<li>Si infiltra nell'esempio sostituendo l'etanolo con 50% resina epossidica resina in etanolo per 30 min a temperatura ambiente in un agitatore, poi 70% resina epossidica in etanolo per 30 min a temperatura ambiente in un agitatore.</li>
		<li>Interruttore la resina epossidica resina soluzione con resina epossidica di 100% e incubare per 10 min a 50 scambi di eseguire due ° C. dell'epossiresina 100% fresco con incubazioni per 1 h a 50 ° C. Aggiungi fresco 100% resina epossidica resina e lasciar per indurire a 50 ° C durante la notte e poi a 60 & n. 17 6; C per oltre 36 h a indurire.</li>
	</ol></li> <li>Rimuovere ogni campione da muti-pozzetti con un gioielliere &#39; sega a mano s. Selezionare le regioni di interesse, dense concentrazioni di cellule, utilizzando un microscopio ottico invertito e poi tagliare blocchi nm 70-80 per il sezionamento di microtomo. Montare la regione di taglio nel mandrino microtomo e caricare il microtomo. Posizionare il mandrino nel microtomo e montare una lama di diamante con il bordo parallelo alla superficie del blocco. Raccogliere le sezioni di spessore nm 70-80 direttamente sulle griglie individuali.</li>
	<li>Sciogliere acetato di uranile in acqua per una soluzione di 1% in peso e sciogliere separatamente citrato di piombo in acqua per una soluzione al 3% in peso. Colorante di contrasto le sezioni da immersione con acetato di uranile acquosa 1% per 15 min e poi 3% acquosa piombo citrato per 5 min migliorare il contrasto dei campioni.</li>
	<li>Formazione immagine EM <ol><li>esaminare le sezioni con un microscopio elettronico a trasmissione e registrare le immagini con una fotocamera digitale CCD ad un ingrandimento primario di 10.000-20, 000 X 3.</li>
	</ol></li> <li>Statistiche <ol><li>eseguire un t-test per identificare differenze significative confrontando la media e l'errore standard di misura (s.e.m.) tra il controllo e i campioni sperimentali.</li>
	</ol>
	</li>
</ol>

<ol>
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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+for+recycling+of+synaptic+vesicle+membrane+during+transmitter+release+at+the+frog+neuromuscular+junction.">Evidence for recycling of synaptic vesicle membrane during transmitter release at the frog neuromuscular junction.</a> J Cell Biol. 57 (2), 315-344 (1973).</li><li>Ceccarelli, B., Hurlbut, W. P., Mauro, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Turnover+of+transmitter+and+synaptic+vesicles+at+the+frog+neuromuscular+junction.">Turnover of transmitter and synaptic vesicles at the frog neuromuscular junction.</a> J Cell Biol. 57 (2), 499-524 (1973).</li><li>Watanabe, S., Rost, B. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrafast+endocytosis+at+mouse+hippocampal+synapses.">Ultrafast endocytosis at mouse hippocampal synapses.</a> Nature. 504 (7479), 242-247 (2013).</li><li>Watanabe, S., Trimbuch, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clathrin+regenerates+synaptic+vesicles+from+endosomes.">Clathrin regenerates synaptic vesicles from endosomes.</a> Nature. 515 (7526), 228-233 (2014).</li><li>Sankaranarayanan, S., Atluri, P. P., Ryan, T. 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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+secretion+and+synaptic+transmission+with+pH-sensitive+green+fluorescent+proteins.">Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins.</a> Nature. 394 (6689), 192-195 (1998).</li><li>Arnao, M. B. B., Acosta, M., del Rio, J. A. A., Garc&#237;a-C&#225;novas, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inactivation+of+peroxidase+by+hydrogen+peroxide+and+its+protection+by+a+reductant+agent.">Inactivation of peroxidase by hydrogen peroxide and its protection by a reductant agent.</a> Biochim Biophys Acta. 1038 (1), 85-89 (1990).</li><li>De&#225;k, F., Schoch, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptobrevin+is+essential+for+fast+synaptic-vesicle+endocytosis.">Synaptobrevin is essential for fast synaptic-vesicle endocytosis.</a> Nat Cell Biol. 6 (11), 1102-1108 (2004).</li><li>Clayton, E. L., Evans, G. J. O., Cousin, 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=Bulk+synaptic+vesicle+endocytosis+is+rapidly+triggered+during+strong+stimulation.">Bulk synaptic vesicle endocytosis is rapidly triggered during strong stimulation.</a> J Neurosci. 28 (26), 6627-6632 (2008).</li><li>Kavalali, E. T., Jorgensen, E. 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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+vesicles+interchange+their+membrane+proteins+with+a+large+surface+reservoir+during+recycling.">Synaptic vesicles interchange their membrane proteins with a large surface reservoir during recycling.</a> Neuron. 51 (2), 179-186 (2006).</li><li>Gimber, N., Tadeus, G., Maritzen, T., Schmoranzer, J., Haucke, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diffusional+spread+and+confinement+of+newly+exocytosed+synaptic+vesicle+proteins.">Diffusional spread and confinement of newly exocytosed synaptic vesicle proteins.</a> Nat Commun. 6, 8392 (2015).</li><li>Nicholson-Fish, J. C., Smillie, K. J., Cousin, M. 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Qui dimostriamo due metodi per il monitoraggio di endocitosi delle vescicole sinaptiche. Nel primo metodo, abbiamo monitorato pHluorin fuse con una proteina della vescicola sinaptica nei neuroni trasfettati e successivamente elettricamente stimolata. In secondo luogo, abbiamo usato la formazione immagine EM dell'assorbimento HRP come indotto da KCl. Abbiamo usato diversi stimoli per due motivi. In primo luogo, applicazione di potassio induce la depolarizzazione di tutti i neuroni nella cultura. Questo facilita l'esame EM...

Neurotrasmettitori sono memorizzati in vescicole sinaptiche e distribuiti dalla esocitosi. La membrana della vescicola sinaptica e la proteina sono quindi interiorizzati dal endocytosis e riutilizzati nel prossimo turno di esocitosi. Endocitosi delle vescicole sinaptiche sono importante per il mantenimento piscine delle vescicole sinaptiche e rimuove vescicole sporgente dalla membrana plasmatica. PHluorin la proteina fluorescente verde sensibili al pH, che è spenta in circostanze acide e dequenched a pH neutro, è stato utilizzato per misurare il tempo di endocitosi corsi in diretta le cellule1,2,3. La proteina di pHluorin è in genere associata al lato lumenal di proteine delle vescicole sinaptiche, come lo synaptophysin o VAMP2/sinaptobrevina. A riposo, pHluorin è spenta in lumen pH 5,5 delle vescicole sinaptiche. Fusione della vescicola alla membrana plasmatica espone il lume vescicolare alla soluzione extracellulare dove il pH è ~ 7.3, conseguente aumento nella fluorescenza pHluorin. Dopo l'esocitosi, l'aumento della fluorescenza decade, a causa di endocitosi delle proteine delle vescicole sinaptiche, seguita da ri-acidificazione vescicola all'interno di tali vescicole recuperate. Anche se il decadimento riflette sia endocitosi e ri-acidificazione vescicolare, essa riflette principalmente endocitosi, perché ri-acidificazione è più veloce di endocitosi nella maggior parte delle condizioni1,4. La costante di tempo di ri-acidificazione è 3-4 s o meno5,6, che è generalmente più veloce di 10 s o superiore richiesto per endocitosi delle vescicole4,5. Se sono necessari esperimenti per distinguere endocitosi da ri-acidificazione, acidi dissetante esperimenti utilizzando la soluzione di (MES) acida 4-Morpholineethanesulfonic (25 mM) con un pH di 5.5 possono essere utilizzati per determinare se vengono recuperate proteine delle vescicole sinaptiche dalla membrana del plasma tramite endocitosi1,3,4. Così, l'aumento di intensità di fluorescenza pHluorin riflette un equilibrio di exo - ed endocitosi e la diminuzione dopo stimolazione del nervo in particolare riflette endocitosi.
pHluorin formazione immagine può essere usata non solo per misurare il corso di tempo dell'endocitosi, ma anche la dimensione della vescicola sinaptica piscine7,8, e la probabilità di rilascio evocato e spontaneo versione9. Molti fattori e proteine coinvolte nella regolazione di endocitosi, quali calcio, le proteine solubili NSF-attachment proteina recettore (rullante), factor(BDNF) neurotrophic cervello-derivato e della calcineurina sono stati identificati usando pHluorin imaging1 , 2 , 10 , 11 , 12 , 13 , 14 , 15 , 16. Inoltre, il rilascio di neurotrasmettitore potrebbe essere rilevata nei neuroni non solo primari ma in cellule di neuroblastoma con TIRFM17. Recentemente, pHTomato, dsRed, mOrange e varianti di pHluorin sono stati sviluppati per il monitoraggio di registrazioni simultanee di fattori multipli in una singola sinapsi18,19. Ad esempio, pHTomato è stato fuso con lo synaptophysin e utilizzato con un indicatore di calcio codificato geneticamente (GCaMP5K) per monitorare la fusione della vescicola presinaptica e afflusso di Ca2 + nel compartimento postsinaptico20. Di conseguenza, pHluorin associata a proteine sinaptiche fornisce un metodo utile per analizzare la relazione fra endocitosi ed esocitosi.
EM è un altro metodo comunemente usato per lo studio di endocitosi, grazie all'elevata risoluzione spaziale che mostra i cambiamenti ultrastrutturali durante endocitosi. Due aree generali sono la capacità di visualizzare cambiamenti patologici all'interno di cellule neuronali21 e tenere traccia della vescicola proteine22. In particolare, l'osservazione dell'assorbimento delle vescicole sinaptiche, curvatura di membrana rivestite di clatrina nella zona periattiva ed endosomal strutture sono possibili con EM3,23,24,25 ,26,27,28. Mentre EM comporta potenziali artefatti, come fissativo-indotto malformazioni, che possono pregiudicare la endocitosi, e analisi dei dati è laboriosa, che la risoluzione fornisce un'opportunità interessante per visualizzare la struttura cellulare. Potenziali problemi di fissativo e la limitazione a risoluzione temporale EM può essere superati da alta pressione congelamento, fornendo un metodo veloce e non chimici di stabilizzare le delicate strutture presenti durante endocitosi27.

<table>
	<tbody>
		<tr>
			<td>Lipofectamine LTX with Plus</td>
			<td>Thermo Fisher</td>
			<td>15338-100</td>
			<td>Transfection of plasmid DNA including synaptophysin or VAMP2-pHluorin</td>
		</tr>
		<tr>
			<td>neurobasal medium</td>
			<td>Thermo Fisher</td>
			<td>21103-049</td>
			<td>Growth medium for neuron, Warm up to 37&#176;C before use</td>
		</tr>
		<tr>
			<td>B27</td>
			<td>Thermo Fisher</td>
			<td>17504-044</td>
			<td>Gradient for neuronal differentiation</td>
		</tr>
		<tr>
			<td>Glutamax</td>
			<td>Thermo Fisher</td>
			<td>35050-061</td>
			<td>Gradient for neuronal culture</td>
		</tr>
		<tr>
			<td>Poly-D-Lysine coated coverslip</td>
			<td>Neuvitro</td>
			<td>GG-25-pdl</td>
			<td>Substrate for neuronal growth and imaging of pHluorin</td>
		</tr>
		<tr>
			<td>Trypsin XI from bovine pancrease</td>
			<td>Sigma</td>
			<td>T1005</td>
			<td>Neuronal culture-digest hippocampal tissues</td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I from bovine pancreas</td>
			<td>Sigma</td>
			<td>D5025</td>
			<td>Neuronal culture-inhibits viscous cell suspension</td>
		</tr>
		<tr>
			<td>pulse stimulator</td>
			<td>A-M systems</td>
			<td>model 2100</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>Slotted bath with field stimulation</td>
			<td>Warner Instruments</td>
			<td>RC-21BRFS</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>stimulus isolation unit</td>
			<td>Warner Instruments</td>
			<td>SIU102</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>lubricant</td>
			<td>Dow corning</td>
			<td>111</td>
			<td>pHluorin imaging-seal with coverslip and imaging chamber, avoid leak from chamber</td>
		</tr>
		<tr>
			<td>AP5</td>
			<td>Tocris</td>
			<td>3693</td>
			<td>Gradient for normal saline, selective NMDA receptor antagonist, inhibit postsynaptic activity which have potential for recurrent activity</td>
		</tr>
		<tr>
			<td>CNQX</td>
			<td>Tocris</td>
			<td>190</td>
			<td>Gradient for normal saline, competitive AMPA/kainate receptor antagonist, inhibit postsynaptic activity which have potential for recurrent activity</td>
		</tr>
		<tr>
			<td>Illuminator</td>
			<td>Nikon</td>
			<td>C-HGFI</td>
			<td>Metal halide light source for pHluorin</td>
		</tr>
		<tr>
			<td>EMCCD camera</td>
			<td>Andor</td>
			<td>iXon3</td>
			<td>pHluorin imaging, detect pHluorin fluorescence intensity</td>
		</tr>
		<tr>
			<td>Inverted microscopy</td>
			<td>Nikon</td>
			<td>Ti-E</td>
			<td>Imaging for synaptophysin or VAMP2 pHluorin transfected cells</td>
		</tr>
		<tr>
			<td>NIS-Elements AR</td>
			<td>Nikon</td>
			<td>NIS-Elements Advanced Research</td>
			<td>Software for imaging acquisition and analysis</td>
		</tr>
		<tr>
			<td>Igor Pro</td>
			<td>WaveMetrics</td>
			<td>Igor pro</td>
			<td>Software for imaging analysis and data presentation</td>
		</tr>
		<tr>
			<td>imaging chamber</td>
			<td>Warner Instruments</td>
			<td>RC21B</td>
			<td>pHluorin imaging, apply field stimulation on living cells</td>
		</tr>
		<tr>
			<td>poly-l-lysine</td>
			<td>Sigma</td>
			<td>P4832</td>
			<td>Electron microscopy, substrate for neuronal growth, apply on multiwell plate for 1 h at room temperature then wash with sterilized water 3 times</td>
		</tr>
		<tr>
			<td>Horseradish peroxidase(HRP)</td>
			<td>Sigma</td>
			<td>P6782</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles by catalyzing DAB in presence hydrogen peroxide, final concentration is 5 mg/mL in normal saline, make fresh before use</td>
		</tr>
		<tr>
			<td>Na cacodylate</td>
			<td>Electron Microscopy Sciences</td>
			<td>12300</td>
			<td>Electron microscopy, buffer for fixatives and washing, final concentration is 0.1 N</td>
		</tr>
		<tr>
			<td>3,3&#8242;-Diaminobenzidine(DAB)</td>
			<td>Sigma</td>
			<td>D8001</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles, substrate for HRP, final concentration is 0.5 mg/mL in DDW and filtered, make fresh before use</td>
		</tr>
		<tr>
			<td>Hydrogen peroxide solution</td>
			<td>Sigma</td>
			<td>H1009</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles by inducing HRP-DAB reaction, final concentration is 0.3% in DDW, make fresh before use</td>
		</tr>
		<tr>
			<td>glutaraldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>16365</td>
			<td>Electron microscopy, fixatives, final concentration is 4% in Na-cacodylate buffer, make fresh before use, shake well before to use</td>
		</tr>
		<tr>
			<td>TEM</td>
			<td>JEOL</td>
			<td>200CX</td>
			<td>Electron microscopy, imaging of endocytosed vesicles and ultrastructural changes</td>
		</tr>
		<tr>
			<td>CCD digital camera</td>
			<td>AMT</td>
			<td>XR-100</td>
			<td>Electron microscopy, capturing images</td>
		</tr>
		<tr>
			<td>Lead citrate</td>
			<td>Leica microsystems</td>
			<td>16707235</td>
			<td>Electron microscopy, grid staining</td>
		</tr>
	</tbody>
</table>

measuring synaptic vesicle endocytosis in cultured hippocampal neurons

Durante endocitosi, fusione delle vescicole sinaptiche sono Estratto alle terminazioni nervose, permettendo per riciclo delle vescicole e quindi il mantenimento della trasmissione sinaptica durante la cottura ripetuto del nervo. Alterata endocitosi in condizioni patologiche conduce alle diminuzioni in funzioni sinaptiche di forza e cervello. Qui, descriviamo i metodi utilizzati per misurare la endocitosi delle vescicole sinaptiche alla sinapsi ippocampo mammifera in coltura neuronale. Abbiamo monitorato endocitosi proteine delle vescicole sinaptiche fondendo una proteina di membrana vescicolare sinaptica, compreso lo synaptophysin e VAMP2/sinaptobrevina, sul lato di lumenal vescicolare, con pHluorin, una proteina fluorescente verde di pH sensibili che aumenta la sua aumenta l'intensità di fluorescenza come il pH. Durante l'esocitosi, vescicolare lumen pH aumenta, mentre durante il lume vescicolare endocitosi pH è ri-acidificato. Così, un aumento dell'intensità di fluorescenza pHluorin indica fusione, mentre una diminuzione indica endocitosi della proteina marcata delle vescicole sinaptiche. Oltre a utilizzare il metodo di imaging pHluorin per registrare endocitosi, abbiamo monitorato endocitosi membrana vescicolare da misure di microscopia elettronica (EM) del rafano perossidasi (HRP) assorbimento dalle vescicole. Infine, abbiamo monitorato la formazione dei pozzi di membrana terminale del nervo in tempi diversi dopo la depolarizzazione indotta da potassio alta. Il corso di formazione di pozzo di assorbimento e membrana HRP tempo indica il corso di tempo dell'endocitosi.

Utilizzando il metodo di elemento portante del lipido, SpH è stato espresso nei neuroni dell'ippocampo, che consenta l'identificazione dei boutons (Figura 1a). La stimolazione elettrica delle cellule indotto esocitosi e un corrispondente aumento in intensità di fluorescenza. L'aumento della fluorescenza (ΔF) è stato interrotto terminando lo stimolo (Figura 1b). L'aumento della fluorescenza è stata seguita da una lenta diminu...

Watch this Scientific Journal Video about Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons at JoVE.com

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

Endocitosi delle vescicole sinaptiche sono rilevato da microscopia chiara di pHluorin fuso con proteine delle vescicole sinaptiche e da microscopia elettronica dell'assorbimento della vescicola.

Endocitosi delle vescicole sinaptiche in colture di neuroni ippocampali di misura

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic ...

measuring-synaptic-vesicle-endocytosis-in-cultured-hippocampal-neurons

Research

JoVE Journal

Neuroscience

9.8K Views.  National Institute of Neurological Disorders and Stroke. Endocitosi delle vescicole sinaptiche sono rilevato da microscopia chiara di pHluorin fuso con proteine delle vescicole sinaptiche e da microscopia elettronica dell'assorbimento della vescicola.

Video: Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

Studiare Piscine vescicole sinaptiche con fotoconversione di coloranti Styryl

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Ricerca

Neuroscienze

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

Recupero della fluorescenza dopo photobleaching (FRAP) di fluorescenza proteine ​​Tagged in spine dendritiche di colture di neuroni ippocampali

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Analisi di Morfologia spine dendritiche in colture di neuroni del SNC

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

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 ...

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

Indurre crescita dendritica in colture di neuroni simpatici

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Diffusione laterale e esocitosi delle proteine ​​di membrana in neuroni in coltura valutata utilizzando Recovery fluorescenza e fluorescenza-loss photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

Rilevamento di palmitoilazione proteina in colture di neuroni ippocampali di immunoprecipitazione e di acil-Biotina Exchange (ABE)

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been extensively evaluated. Traditional methods are time consuming, expensive, resource intensive and require replicating cells. In contrast, the comet assay, a single cell gel electrophoresis assay, is a faster, non-invasive, inexpensive, direct and sensitive measure of DNA damage and repair. All forms of DNA damage as well as DNA repair can be visualized at the single cell level using this powerful technique.
	The principle underlying the comet assay is that intact DNA is highly ordered whereas DNA damage disrupts this organization. The damaged DNA seeps into the agarose matrix and when subjected to an electric field, the negatively charged DNA migrates towards the cathode which is positively charged. The large undamaged DNA strands are not able to migrate far from the nucleus. DNA damage creates smaller DNA fragments which travel farther than the intact DNA. Comet Assay, an image analysis software, measures and compares the overall fluorescent intensity of the DNA in the nucleus with DNA that has migrated out of the nucleus. Fluorescent signal from the migrated DNA is proportional to DNA damage. Longer brighter DNA tail signifies increased DNA damage. Some of the parameters that are measured are tail moment which is a measure of both the amount of DNA and distribution of DNA in the tail, tail length and percentage of DNA in the tail. This assay allows to measure DNA repair as well since resolution of DNA damage signifies repair has taken place. The limit of sensitivity is approximately 50 strand breaks per diploid mammalian cell 1,2. Cells treated with any DNA damaging agents, such as etoposide, may be used as a positive control. Thus the comet assay is a quick and effective procedure to measure DNA damage.

The comet assay is an efficient way of detecting single- and double-strand breaks, including alkali-labile sites and DNA-DNA/DNA-protein cross-links on the DNA in all cells including hippocampal neurons. The method takes advantage of the differential migration of DNA in an electric field due to differences in amount of DNA damage.

Il saggio danneggiare il DNA in neuroni ippocampali Utilizzo del Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Post-embedding Etichettatura immuno di proteine ​​sinaptiche in Culture Slice ippocampali

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using styryl FM dyes, which reveal membrane turnover. Conventional protocols for the use of FM dyes were designed for analyzing neurons following stimulated (evoked) synaptic activity. Recently, protocols have become available for analyzing the FM signals that accompany weaker synaptic activities, such as spontaneous or miniature synaptic events. Analysis of these small changes in FM signals requires that the imaging system is sufficiently sensitive to detect small changes in intensity, yet that artifactual changes of large amplitude are suppressed. Here we describe a protocol that can be applied to evoked, spontaneous, and miniature synaptic activities, and use cultured hippocampal neurons as an example. This protocol also incorporates a means of assessing the rate of photobleaching of FM dyes, as this is a significant source of artifacts when imaging small changes in intensity.

We describe the use of styryl FM dyes to image synaptic vesicle recycling in functional nerve terminals. This protocol can be applied not only to evoked, but also spontaneous and miniature synaptic activities. The protocol expands the variety of synaptic events that can be effectively evaluated.

Esame delle vescicole sinaptiche Riciclaggio Uso FM coloranti Durante evocati, spontaneo, e miniatura Attività Synaptic

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using ...