Ringraziamo Birgit Westernstroeer del Centro di Medicina della Riproduzione e Andrologia presso l&#39;Università di Muenster per l&#39;insegnamento della microiniezione testicolare. Inoltre, siamo grati a Ursula Antkewitz e Petra Reckling per l&#39;assistenza tecnica. Ringraziamo la Fondazione tedesca per la ricerca (DFG) per sostenere questo lavoro (WE2458 / 10-1). &#160;

Tutti gli esperimenti sugli animali effettuati sono stati approvati dal comitato etico locale (Landesamt für Landwirtschaft, Lebensmittelsicherheit und Fischerei, Mecklenburg-Vorpommern, Germania). 1 plasmidi Preparazione <ol><li> Per la preparazione plasmide, utilizzare i kit di purificazione del plasmide (vedi tabella materiali) o metodi simili con buffer di rimozione di endotossine in modo che le reazioni immunitarie dell&#39;animale possono essere evitati. Seguire le istruzioni del manuale. Impiegare DDH 2 O per diluire la soluzione plasmide. </li><li> Eliminare i detriti facendo girare la soluzione di DNA alla massima velocità (20.000 xg). Poi raccogliere il surnatante. </li><li> Determinare la concentrazione plasmide con uno spettrofotometro (vedi materiali). </li><li> Regolare la concentrazione plasmide con DDH 2 O a 1-3 mg (DNA) / ml. Nota: le concentrazioni più basse riducono l&#39;efficienza di trasfezione, mentre concentrazioni più elevate possono provocare una soluzione iniettabile troppo viscoso. Inoltre, l&#39;efficienza di trasfezione dipende dalle dimensioni del plasmide e deve essere testato singolarmente. </li><li> Prima dell&#39;iniezione preparare una miscela della soluzione con ad esempio 40 microlitri plasmide con 5 ml di PBS (10x) e 5 microlitri Verde veloce (0,5%). Veloce verde è necessaria per il monitoraggio del processo di iniezione. Preferibilmente, utilizzare parete PCR-tubi sottili o Parafilm. Nota: a seconda delle dimensioni del testicolo, sarà necessario un volume di 20-50 ml per ogni testicolo. </li></ol> 2 Preparazione di microiniezione pipetta <ol><li> Utilizzare capillari borosilicato (vedi tabella materiali) per la preparazione dei pipette microiniezione. </li><li> Tirare capillari di vetro con un capillare estrattore verticale (vedi tabella Materials). </li><li> Rompere la punta capillare con una pinza da dolcemente banding. La punta è di solito 50-80 micron di diametro e lunghezza di circa 1 cm. Cercate di evitare punte troppo lungo in quanto questi non sono sufficientemente rigido per penetrare il tessuto. </li><li> Infine, shaRPEN la punta di un 30 ° a 45 ° utilizzando un beveler micropipetta (vedi tabella Materials). </li><li> Caricare la pipetta microiniezione con il mix di soluzione iniettabile (vedere plasmidi Preparazione). Applicare la soluzione nel retro del capillare di vetro con una piccola siringa. Nota: Cercate di evitare bolle d&#39;aria durante il caricamento, altrimenti questi saranno iniettati nei tubuli seminiferi con la soluzione di plasmide e quindi ridurre la conducibilità, nonché, eventualmente, causare danni ai tessuti. </li></ol> 3 Anestesia e Chirurgia <ol><li> Eseguire l&#39;operazione in condizioni asettiche, utilizzando materiale sterile (ad esempio, siringa, ago, strumenti chirurgici, ecc). Ciò consentirà di ridurre le infezioni degli animali e garantire buoni tassi di sopravvivenza. </li><li> Utilizzare topi maschi di elettroporazione ad un&#39;età di 6-8 settimane. </li><li> Per inizializzare il trattamento analgesico post-chirurgica, forniscono il mouse con analgesici aggiunto l&#39;acqua potabile, il giorno prima dell&#39;intervento. Questo culoUres una analgesia preventiva in caso di altamente probabile hypodipsia post-chirurgica (assunzione di acqua diminuita). A tal fine, esporre acqua potabile con ad esempio una sospensione ibuprofene pediatrico (20 mg / ml). L&#39;acqua da bere medicata deve essere fornito anche il primo giorno e due di recupero in concentrazione identici. Ciò significa che una dose giornaliera di 7,5 mg / kg, valida per 5 ml / acqua potabile d e peso corporeo di 25 g per 8 settimane di età C57BL / 6J. Questo regime analgesico garantisce un sollievo dal dolore fondamentale e recupero accelerato con alto benessere 26. </li><li> Per preparare l&#39;anestesia soluzione sul giorno dell&#39;intervento di lavoro, miscelare il 10% e il 2% Ketamin Xylazin in un rapporto 1: 1 (vedi tabella Materials). </li><li> Per anestetizzare l&#39;animale, diffusa 0,25 ml / 100 mg di peso corporeo di anestetico per via sottocutanea soluzione (Ketamin = 0,125 g / kg; Xylazin = 0,025 g / kg). Utilizzare un sito di iniezione tra bacino e degli arti, al fine di evitare danni d&#39;organo e lesioni dei topi. Di solito, 10 min sono necessarifino a quando il mouse è profondamente anestetizzati. </li><li> Coprire gli occhi del mouse con vet pomata per prevenire la secchezza durante l&#39;anestesia. </li><li> Prova profonda arresto anestetico, che è evidente da una totale mancanza di risposta. A tal fine, basta pizzicare la punta dell&#39;animale. </li><li> Per avviare l&#39;operazione, rimuovere i capelli addominale con un rasoio elettrico o simile e disinfettare la zona funzionamento con sterilium o simili. </li><li> Fare una incisione ventrale direttamente sopra le ghiandole prepuziale nel centro della zona addominale. In quel luogo, prima tirare la pelle fuori e condurre un piccolo taglio trasversale cutanea di circa 8-14 mm. Quindi, proseguire lungo lo strato muscolare addominale. Nota: La lesione deve essere il più piccolo possibile per ridurre i danni all&#39;animale. </li><li> Sollevare i cuscinetti di grasso addominale con attenzione per esporre il testicolo collegato e posizionarlo su una precedente preparato impermeabile usa e getta di carta drappo proprio accanto al sito di incisione (Figura 2A). </li><li> Utilizzare un binocolo per find il condotto efferente come destinazione di iniezione. Il dotto efferente è identificato come la multa di giunzione nave tra il testicolo e dell&#39;epididimo. Si trova adiacente alla arteria testicolare prominente. Il dotto corre quasi con un angolo di 45 ° verso l&#39;arteria e visibilmente entra nel epididymidis caput. Nota: A seconda dell&#39;età del mouse, il dotto è di solito sepolto nel tessuto adiposo. </li><li> Utilizzare una pinza sottile per cancellare il canale efferente di questo tessuto adiposo. Dopo di che, posizionare il dotto rilasciato su una striscia di carta sterile per garantire la chiara visibilità del condotto senza compromettere dintorni. </li></ol> 4. Microiniezione e applicazioni del DNA <ol><li> Collegare la pipetta microiniezione, che viene caricato con la soluzione plasmide all&#39;unità micromanipolatore / iniettore. </li><li> Inserire la microiniezione ago parallelo al condotto efferente con la punta verso rete testis (Figura 2B). </li><li> Utilizzare pinzette sottili etogliere il condotto sopra la pipetta microiniezione. Assicurarsi che il capillare è mantenuta parallela al condotto mentre si tira su. Nota: Questa procedura è più conveniente che penetra la nave spostando l&#39;ago con il micromanipolatore. </li><li> Dirigere l&#39;ago con attenzione verso il testicolo e fermarsi proprio come penetra la rete testis direttamente sotto la tunica albuginea (Figura 2C). Nota: Nel caso che travalica la rete testis, la soluzione plasmide perderà nello spazio interstiziale. Questo generalmente porta al fallimento di transfezione tubuli seminiferi. </li><li> Per l&#39;iniezione della soluzione plasmide utilizzare il microinjector con le seguenti impostazioni: pi: 100 hPa, ti: 0,2 sec, e pc: 0 hPa (Figura 2D). </li><li> Monitorare l&#39;intero processo di iniezione osservando testicolo stato di riempimento con l&#39;aiuto del colore verde. Fare attenzione che il testicolo è riempito solo fino a 2/3 del suo volume con la plsoluzione asmid (Figura 2E). Nota: Se il volume di iniezione viene superato, il tessuto testicolare potrebbe essere danneggiato. </li></ol> 5. Elettroporazione di Testicolo <ol><li> Per abilitare elettroporazione efficace, immergere gli elettrodi pinzetta in PBS (1x). Questo assicura un adeguato conduttanza. </li><li> Uniformemente spremere il testicolo tra gli elettrodi bagnati (Figura 2F). Misurare la resistenza elettrica con il elettroporatore. </li><li> Applicare corrente al testicolo. Eseguire otto impulsi quadrati di 40 V a 4 diversi siti testicolo con una durata costante di tempo di impulso 50 msec e 950 msec intervallo di tempo. </li></ol> 6. ferita chiusura e post-operatorio <ol><li> Quando elettroporazione è finito, posizionare il testicolo torna nella posizione originale. </li><li> Cucire lo strato muscolare interno con sutura chirurgica (vedi materiali). </li><li> Chiudere la pelle utilizzando spezzoni di sutura (vedi tabella materiali). Tirare la pelle fino with pinzette. Assicurati di escludere lo strato muscolare. Posizionare le clips, ciascuno ad una distanza di non più di 5 mm. Nota: Poiché sutura chirurgica può essere inghiottito dal mouse, clip di sutura sono favoriti per la chiusura della lesione cutanea. </li><li> Lasciare il mouse per recuperare da anestesia su carta assorbente sterili collocati in una gabbia sterile topo vuoto, che è temperato su un 37 ° C pad caldo. Il tampone dovrebbe garantire il riscaldamento di una metà della gabbia creando un gradiente termico. Per l&#39;animale questo rende possibile effettuare la ricerca per la singola area, più comodo per la varietà temperatura nella gabbia. Inoltre, coprire il mouse con un foglio di carta assorbente sterile in modo che lo stress può essere ulteriormente ridotto. Nota: Dal momento che la superficie del corpo del mouse è insolitamente alta in relazione alla massa corporea, se confrontato con gli animali più grandi, il supporto termico è di particolare importanza per il successo il recupero dei roditori. </li><li> Quando l&#39;animale operato si è completamente risvegliato, trasferire it per un ulteriore recupero di una nuova gabbia del mouse completamente attrezzata. Non mettere l&#39;animale torna alla sua vecchia gabbia o ad un gruppo di topi. Assicurarsi che la gabbia è pulita e sterile possibile per prevenire l&#39;infezione della ferita. Monitorare l&#39;intero processo di recupero. Per trasferire il mouse torna alla struttura di cura degli animali, attendere fino a che non ha pienamente recuperato e sembra comportarsi normalmente. Nota: persona aggiuntiva e strategie post-chirurgici sono discusse da Pritchett-Corning KR et al 6. </li></ol>

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Marten Michaelis, Alexander Sobczak, e Joachim M. Weitzel impiegati presso l&#39;Istituto di Biologia della Riproduzione, Leibniz Institute for Biologia Animal Farm (FBN), Dummerstorf, Germania, dichiarano di non avere interessi finanziari in competizione.

La ricerca nel campo della biologia riproduttiva, in particolare nel settore della fertilità maschile e la spermatogenesi inevitabilmente si basa su organismi viventi. Al fine di esaminare la funzione testicolare, no / sistema di coltura cellulare adeguata in vitro è stata stabilita in grado di riflettere tutti i cambiamenti morfologici cruciali da un spermatogonio diploide ad aploide un maturo 1,2 spermatozoo. Così, la generazione di animali geneticamente modificati è spesso nec...

Spermatogenesi dei mammiferi è considerato un sofisticato processo di cellule staminali di auto-rinnovamento successivamente sottoposti a mitosi, meiosi e la differenziazione al fine di sviluppare in maturo spermatozoi aploidi. Questi cambiamenti morfologici sono orchestrati da diversi tipi di cellule e nonostante i profondi tentativi, è ancora impossibile imitare questi processi in coltura cellulare 1,2. Quindi, la ricerca sulla spermatogenesi fino ad ora si affida a organismi viventi come modelli in vivo. In generale, gli studi di funzione genica si basano generalmente su animali transgenici. Tuttavia, generare e sostenere questo tipo di modello animale è in termini di tempo, costi-intensive e molto elaborata. Questo è attribuito al processo di allevamento lungo necessario per generare e mantenere il transgene nel corso delle generazioni. Inoltre, la manipolazione genetica di tutto l&#39;organismo dall&#39;approccio transgenici o knockout è incline a causare alterazioni fisiologiche quando il targeting gEnes con funzioni essenziali in più regioni, ad esempio, al di fuori del testicolo o sistemica. Inoltre, alcuni metodi di trasfezione transiente sono associati con alcuni svantaggi cruciali. Ad esempio, inconvenienti tipici di trasferimento genico virus-mediata sono la possibile provocazione di immunoreazioni e dei regolamenti di sicurezza supplementari, mentre lipofezione 3 e 4 microparticelle bombardamento potrebbe danneggiare il tessuto e sono limitate a una certa profondità cella di una sufficiente efficienza di trasfezione. In contrasto, elettroporazione (EP) come un altro modo comune di trasfezione transiente, sembra costituire una tecnica promettente per consentire a trasfezione vivo e vivo analisi del gene coerente. In generale, EP viene indicato come un fenomeno dinamico che dipende dalla tensione transmembrana locale con pori di conseguenza mediate all&#39;interno nanosecondi. Queste lacune possono essere mantenuti per i millisecondi, sufficienti to concedere l&#39;accesso al DNA, RNA o piccole molecole 5. Quando la tensione applicata è troppo alta, il carattere di solito transitoria di EP è contrastata a causa della produzione di calore e di induzione di permeabilizzazione troppo globale con conseguente danno irreversibile della cellula 5. Qui mostriamo che elettroporazione è un sistema trasfezione efficace ed economico che possa essere utilizzato per la trasformazione genetica testicolo per chiarire caratteristiche genetiche testicolari in vivo. Questo articolo affronta la preparazione plasmide, microiniezione attraverso il dotto efferente e la successiva elettroporazione del mouse del testicolo. Questa procedura può essere il mezzo di scelta per raggiungere velocemente, specifico ed efficiente trasfezione dei tubuli seminiferi del testicolo topo in vivo al fine di indagare i processi di spermatogenesi.

<table border="0" cellpadding="0" cellspacing="3">
	<tbody>
		<tr>
			<td>
			centrifuge
			</td>
			<td>Sigma</td>
			<td>1-15 PK</td>
			<td></td>
		</tr>
		<tr>
			<td>spectrophotometer</td>
			<td>Nanodrop</td>
			<td>ND-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>micropipette puller</td>
			<td>Narishige</td>
			<td>PC-10</td>
			<td>vertical capillary puller</td>
		</tr>
		<tr>
			<td>microinjection capillaries</td>
			<td>Clark</td>
			<td>GC100-10</td>
			<td>borosilicate standard wall</td>
		</tr>
		<tr>
			<td>micropipette beveler</td>
			<td>Bachofer</td>
			<td>Typ 462</td>
			<td>rotating disk beveler</td>
		</tr>
		<tr>
			<td>electroporator</td>
			<td>Nepagene</td>
			<td>CUY 21EDIT</td>
			<td>square wave electroporator</td>
		</tr>
		<tr>
			<td>tweezer electrodes</td>
			<td>Nepagene</td>
			<td>CUY 650P5</td>
			<td>5 mm &#216; disk electrodes</td>
		</tr>
		<tr>
			<td>stereo microscope</td>
			<td>Zeiss</td>
			<td>Stemi 2000-C</td>
			<td></td>
		</tr>
		<tr>
			<td>cold light source</td>
			<td>Zeiss</td>
			<td>KL 1500LCD</td>
			<td></td>
		</tr>
		<tr>
			<td>microinjection pump</td>
			<td>Eppendorf</td>
			<td>Femtojet</td>
			<td></td>
		</tr>
		<tr>
			<td>micromanipulator</td>
			<td>homemade</td>
			<td></td>
			<td>XYZ cross table</td>
		</tr>
		<tr>
			<td>surgical instruments</td>
			<td>FST</td>
			<td></td>
			<td>scissors, forceps, 
			needle holder</td>
		</tr>
		<tr>
			<td>fine forceps</td>
			<td>FST</td>
			<td>Dumont #5, #7</td>
			<td>efferent ducts preparation</td>
		</tr>
		<tr>
			<td>Michel clip applying stapler</td>
			<td>FST</td>
			<td>Michel, 12029-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Michel suture clips</td>
			<td>FST</td>
			<td>Michel, 12040-01</td>
			<td>7.5 x 1.75 mm</td>
		</tr>
		<tr>
			<td>surgical suture</td>
			<td>Catgut 00</td>
			<td>Catgut 
			Markenkirchen</td>
			<td></td>
		</tr>
		<tr>
			<td>syringe, with 30G needle</td>
			<td>B. Braun</td>
			<td>Omnican 40</td>
			<td>to load micropipette and 
			for anesthesia</td>
		</tr>
		<tr>
			<td>plasmid isolation kit</td>
			<td>Promega</td>
			<td>Cat. # A2495</td>
			<td>plasmid Midiprep</td>
		</tr>
		<tr>
			<td>plasmid pEGFP-C1</td>
			<td>Clontech</td>
			<td>Cat. #6084-1</td>
			<td>CMV-promoter + EGFP</td>
		</tr>
		<tr>
			<td>plasmid pDsRed2-N1</td>
			<td>Clontech</td>
			<td>Cat. #6084-1</td>
			<td>CMV-promoter + DsRed2</td>
		</tr>
		<tr>
			<td>Fast Green dye</td>
			<td>Sigma</td>
			<td>F7258-25G</td>
			<td>for dilution in ddH2O</td>
		</tr>
		<tr>
			<td>10x PBS, pH 7.4</td>
			<td>Carl Roth</td>
			<td>3957.1</td>
			<td>NaCl [1.37 M]</td>
		</tr>
		<tr>
			<td>6781.1</td>
			<td>KCl [27 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>T106.2</td>
			<td>Na2HPO4+2H2O [100 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>3904.1</td>
			<td>KH2PO4 [18 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10% Ketamin</td>
			<td>Serum Werk 
			Bernburg</td>
			<td>Urotamin</td>
			<td>mix in a rate 1:1</td>
		</tr>
		<tr>
			<td>2% Xylazin</td>
			<td>Serum Werk 
			Bernburg</td>
			<td>Xylazin</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterilium</td>
			<td>Bode Chemie</td>
			<td>Sterillium</td>
			<td>disinfection</td>
		</tr>
		<tr>
			<td>vet ointment</td>
			<td>S&#38;K Pharma 
			GmbH</td>
			<td>Kerato Biciron 5% 
			Augensalbe</td>
			<td>or similar to prevent 
			dryness of eyes</td>
		</tr>
		<tr>
			<td>To-Pro-3 iodide</td>
			<td>Invitrogen</td>
			<td>T3605</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vivo microinjection and electroporation of mouse testis

Questo contributo video e articolo fornisce una descrizione completa di microiniezione ed elettroporazione di topo testicolo in vivo. Questa particolare tecnica di trasfezione di cellule testicolari del mouse permette lo studio di processi unici in spermatogenesi. Il seguente protocollo si concentra sulla trasfezione delle cellule testicolari del mouse con costrutti plasmidici. In particolare, abbiamo utilizzato il vettore giornalista pEGFP-C1, che esprime la proteina fluorescente verde potenziata (eGFP) e anche il vettore pDsRed2-N1 che esprimono la proteina fluorescente rossa (DsRed2). Entrambi i geni reporter codificati erano sotto il controllo del promotore del citomegalovirus umano immediato-precoce (CMV). Per eseguire il trasferimento genico in testicoli di topo, i costrutti reporter di plasmide vengono iniettate in testicoli di topi viventi. A tal fine, il testicolo di un animale anestetizzato è esposto e il sito di microiniezione è preparato. Il nostro posto preferito diiniezione è il condotto efferente, con la rete testis in ultima analisi, collegato come via di trasporto anatomica degli spermatozoi tra il testicolo e dell&#39;epididimo. In questo modo, il riempimento dei tubuli seminiferi dopo microiniezione è ottimamente gestito e controllato grazie all&#39;utilizzo di soluzioni di DNA macchiati. Dopo aver osservato un ripieno sufficiente del testicolo dalla sua struttura tubulo colorata, l&#39;organo è elettroporata. Questo consente il trasferimento della soluzione di DNA in cellule testicolari. Dopo 3 giorni di incubazione, il testicolo viene rimosso ed esaminato al microscopio per fluorescenza verde o rosso, illustrando il successo trasfezione. Generalmente, questo protocollo può essere impiegato per la consegna DNA o RNA-costruisce in soggiorno topo testicolo per (sopra) esprimere o abbattere i geni, facilitando l&#39;analisi della funzione genica in vivo. Inoltre, è adatto per studiare costrutti reporter o putativo regula geneElementi Tory. Così, i principali vantaggi della tecnica di elettroporazione sono prestazioni veloci in combinazione con il minimo sforzo, così come le attrezzature tecniche moderato e le competenze necessarie rispetto alle tecniche alternative.

L&#39;impostazione sperimentale per l&#39;esecuzione di microiniezione ed elettroporazione di topo testicolo in vivo in quanto è utilizzato in base al protocollo è illustrato in Figura 1. Anche se è possibile acquisire micropipette industrialmente, abbiamo preferito di generare le nostre pipette tirando (Figura 1A ) e smussatura (Figura 1B) capillari di vetro in modo che essi montati nostre esigenze. L&#39;attrezzatura per microiniezione ed elettroporazione ...

Watch this Scientific Journal Video about In Vivo Microinjection and Electroporation of Mouse Testis at JoVE.com

In Vivo Microinjection and Electroporation of Mouse Testis

Questo articolo descrive microiniezione ed elettroporazione di topo testicolo in vivo come una tecnica di trasfezione per le cellule testicolari di topo per studiare processi unici di spermatogenesi. Il protocollo presentato prevede fasi di preparazione capillare di vetro, microiniezione attraverso il dotto efferente, e trasfezione mediante elettroporazione.

In Vivo Microiniezione e Elettroporazione di mouse Testicolo

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

in-vivo-microinjection-and-electroporation-of-mouse-testis

Research

JoVE Journal

Biology

In Vivo Microinjection and Electroporation of Mouse Testis

27.6K Views.  Leibniz Institute for Farm Animal Biology (FBN).  Questo articolo descrive microiniezione ed elettroporazione di topo testicolo in vivo come una tecnica di trasfezione per le cellule testicolari di topo per studiare processi unici di spermatogenesi. Il protocollo presentato prevede fasi di preparazione capillare di vetro, microiniezione attraverso il dotto efferente, e trasfezione mediante elettroporazione. 

Video: In Vivo Microinjection and Electroporation of Mouse Testis

In Utero Intraventricular Injection and Electroporation of E15 Mouse Embryos

In-utero in-vivo injection and electroporation of the embryonic mouse neocortex provides a powerful tool for the manipulation of individual progenitors lining the walls of the lateral ventricle. This technique is now widely used to study the processes involved in corticogenesis by over-expressing or knocking down genes and observing the effects on cellular proliferation, migration, and differentiation. In comparison to traditional knockout strategies, in-utero electroporation provides a rapid means to manipulate a population of cells during a specific temporal window. In this video protocol we outline the experimental methodology for preparing mice for surgery, exposing the uterine horns through laporatomy, injecting DNA into the lateral ventricles of the developing embryo, electroporating DNA into the progenitors lining the lateral wall, and caring for animals post-surgery. Our laboratory uses this protocol for surgeries on E13-E16 mice, however, it is most commonly performed at E15, as shown in this video. 

In Utero iniezione intraventricolare e Elettroporazione di E15 embrioni di topo

In-utero in-vivo injection and electroporation of the embryonic mouse neocortex provides a powerful tool for the manipulation of individual progenitors ...

Ricerca

Biologia

Single Cell Electroporation in vivo within the Intact Developing Brain

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact tissue, including in vivo preparations. The distinct advantage of this technique is that experimental manipulations may be performed on individual cells while leaving the surrounding tissue unaltered, thereby distinguishing cell-autonomous effects from those resulting from global treatments. When combined with advanced in vivo imaging techniques, SCE of fluorescent markers permits direct visualization of cellular morphology, cell growth, and intracellular events over timescales ranging from seconds to days. While this technique is used in a variety of in vivo and ex vivo preparations, we have optimized this technique for use in Xenopus laevis tadpoles. In this video article, we detail the procedure for SCE of a fluorescent dye or plasmid DNA into neurons within the intact brain of the albino Xenopus tadpole. We also discuss methods to optimize yield, and show examples of live two-photon fluorescence imaging of neurons fluorescently labeled by SCE.

Single-cell electroporation (SCE) is a specialized technique allowing delivery of DNA or other macromolecules into individual cells within intact tissue, including in vivo preparations. Here we detail the procedure for SCE of a fluorescent dye or plasmid DNA into neurons within the intact brain of the Xenopus laevis tadpole.

Cella singola elettroporazione in vivo all&#39;interno del cervello intatto in via di sviluppo

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact ...

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell differentiation, axon guidance, retinotopic organization and synapse formation[1]. One drawback is the inability to efficiently and reliably manipulate gene expression in RGCs in vivo, especially in the otherwise accessible murine visual pathways. Transgenic mice can be used to manipulate gene expression, but this approach is often expensive, time consuming, and can produce unwanted side effects. In chick, in ovo electroporation is used to manipulate gene expression in RGCs for examining retina and RGC development. Although similar electroporation techniques have been developed in neonatal mouse pups[2], adult rats[3], and embryonic murine retinae in vitro[4], none of these strategies allow full characterization of RGC development and axon projections in vivo. To this end, we have developed two applications of electroporation, one in utero and the other ex vivo, to specifically target embryonic murine RGCs[5, 6]. 
With in utero retinal electroporation, we can misexpress or downregulate specific genes in RGCs and follow their axon projections through the visual pathways in vivo, allowing examination of guidance decisions at intermediate targets, such as the optic chiasm, or at target regions, such as the lateral geniculate nucleus. Perturbing gene expression in a subset of RGCs in an otherwise wild-type background facilitates an understanding of gene function throughout the retinal pathway. Additionally, we have developed a companion technique for analyzing RGC axon growth in vitro. We electroporate embryonic heads ex vivo, collect and incubate the whole retina, then prepare explants from these retinae several days later. Retinal explants can be used in a variety of in vitro assays in order to examine the response of electroporated RGC axons to guidance cues or other factors. In sum, this set of techniques enhances our ability to misexpress or downregulate genes in RGCs and should greatly aid studies examining RGC development and axon projections.

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

Here we present two techniques for manipulating gene expression in murine retinal ganglion cells (RGCs) by in utero and ex vivo electroporation. These techniques enable one to examine how alterations in gene expression affect RGC development, axon guidance, and functional properties.

In elettroporazione in vivo ed ex utero per l&#39;espressione genica in cellule gangliari della retina del mouse

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell ...

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo transfection of plasmids by various methods into specific tissues such as liver1, skeletal muscle2,3, and even the brain4. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis (FDB) and interosseus (IO) muscles of adult mice using an in vivo electroporation approach. The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3, calpain, FKBP12, &beta;2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and &alpha;-actinin; and membrane proteins, i.e. &alpha;1s-DHPR, RyR1, cardiac Na/Ca2+ exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats. The efficient expression of some of these proteins has been verified with biochemical3 and functional evidence5. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs. The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6-8.

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

We describe detailed procedures for the efficient transfection of plasmid DNA into the fibers of foot muscles of live mice using electroporation and the subsequent visualization of protein expression using fluorescence microscopy.

Transfection DNA dei mammiferi muscoli scheletrici con In Vivo Elettroporazione

 A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Imaging in vivo e trattamenti terapeutici in un modello di mouse ortotopico di cancro ovarico

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1. Apoptosis can be triggered when cells encounter a wide range of cytotoxic stresses. These insults initiate signaling cascades that ultimately cause the release of cytochrome c from the mitochondrial intermembrane space to the cytoplasm2. The release of cytochrome c from mitochondria is a key event that triggers the rapid activation of caspases, the key cellular proteases which ultimately execute cell death3-4.
The pathway of apoptosis is regulated at points upstream and downstream of cytochrome c release from mitochondria5. In order to study the post-mitochondrial regulation of caspase activation, many investigators have turned to direct cytoplasmic microinjection of holocytochrome c (heme-attached) protein into cells6-9. Cytochrome c is normally localized to the mitochondria where attachment of a heme group is necessary to enable it to activate apoptosis10-11. Therefore, to directly activate caspases, it is necessary to inject the holocytochrome c protein instead of its cDNA, because while the expression of cytochrome c from cDNA constructs will result in mitochondrial targeting and heme attachment, it will be sequestered from cytosolic caspases. Thus, the direct cytosolic microinjection of purified heme-attached cytochrome c protein is a useful tool to mimic mitochondrial cytochrome c release and apoptosis without the use of toxic insults which cause cellular and mitochondrial damage.
In this article, we describe a method for the microinjection of cytochrome c protein into cells, using mouse embryonic fibroblasts (MEFs) and primary sympathetic neurons as examples. While this protocol focuses on the injection of cytochrome c for investigations of apoptosis, the techniques shown here can also be easily adapted for microinjection of other proteins of interest.

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

In this protocol, we describe the direct cytoplasmic microinjection of cytochrome c protein into fibroblasts and primary sympathetic neurons. This technique allows for the introduction of cytochrome c protein into the cytoplasm of cells and mimics the release of cytochrome c from mitochondria, which occurs during apoptosis.

Attivazione di apoptosi da Microiniezione citoplasmatici del Citocromo C</em

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1.  Apoptosis can be triggered when cells encounter a ...

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Mistakes in chromosome segregation lead to aneuploid cells. In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to infertility, miscarriages or developmental disorders like Down syndrome. Haploid gametes form through species-specific developmental programs that are coupled to meiosis. The first meiotic division (MI) is unique to meiosis because sister chromatids remain attached while homologous chromosomes are segregated. For reasons not fully understood, this reductional division is prone to errors and is more commonly the source of aneuploidy than errors in meiosis II (MII) or than errors in male meiosis 1,2. 
In mammals, oocytes arrest at prophase of MI with a large, intact germinal vesicle (GV; nucleus) and only resume meiosis when they receive ovulatory cues. Once meiosis resumes, oocytes complete MI and undergo an asymmetric cell division, arresting again at metaphase of MII. Eggs will not complete MII until they are fertilized by sperm. Oocytes also can undergo meiotic maturation using established in vitro culture conditions 3. Because generation of transgenic and gene-targeted mouse mutants is costly and can take long periods of time, manipulation of female gametes in vitro is a more economical and time-saving strategy. 
Here, we describe methods to isolate prophase-arrested oocytes from mice and for microinjection. Any material of choice may be introduced into the oocyte, but because meiotically-competent oocytes are transcriptionally silent 4,5 cRNA, and not DNA, must be injected for ectopic expression studies. To assess ploidy, we describe our conditions for in vitro maturation of oocytes to MII eggs. Historically, chromosome-spreading techniques are used for counting chromosome number 6. This method is technically challenging and is limited to only identifying hyperploidies. Here, we describe a method to determine hypo-and hyperploidies using intact eggs 7-8. This method uses monastrol, a kinesin-5 inhibitor, that collapses the bipolar spindle into a monopolar spindle 9 thus separating chromosomes such that individual kinetochores can readily be detected and counted by using an anti-CREST autoimmune serum. Because this method is performed in intact eggs, chromosomes are not lost due to operator error.

Oocytes are prone to aneuploidy due to errors in chromosome segregation during meiotic maturation. Aneuploid eggs can cause infertility, miscarriages or developmental disorders like Down syndrome. Here, we describe methods to introduce materials of choice into oocytes and methods to study meiotic maturation and assess ploidy.

Microiniezione degli ovociti mouse, Maturazione e Ploidia valutazione

Mistakes in chromosome segregation lead to aneuploid cells.  In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to ...

Germ Cell Transplantation and Testis Tissue Xenografting in Mice

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies showed that microinjection of germ cells from fertile donor mice into the seminiferous tubules of infertile recipient mice results in donor-derived spermatogenesis and sperm production by the recipient animal2. The use of donor males carrying the bacterial &beta;-galactosidase gene allowed identification of donor-derived spermatogenesis and transmission of the donor haplotype to the offspring by recipient animals1. Surprisingly, after transplantation into the lumen of the seminiferous tubules, transplanted germ cells were able to move from the luminal compartment to the basement membrane where spermatogonia are located3. It is generally accepted that only SSCs are able to colonize the niche and re-establish spermatogenesis in the recipient testis. Therefore, germ cell transplantation provides a functional approach to study the stem cell niche in the testis and to characterize putative spermatogonial stem cells. To date, germ cell transplantation is used to elucidate basic stem cell biology, to produce transgenic animals through genetic manipulation of germ cells prior to transplantation4,5, to study Sertoli cell-germ cell interaction6,7, SSC homing and colonization3,8, as well as SSC self-renewal and differentiation9,10.
Germ cell transplantation is also feasible in large species11. In these, the main applications are preservation of fertility, dissemination of elite genetics in animal populations, and generation of transgenic animals as the study of spermatogenesis and SSC biology with this technique is logistically more difficult and expensive than in rodents. Transplantation of germ cells from large species into the seminiferous tubules of mice results in colonization of donor cells and spermatogonial expansion, but not in their full differentiation presumably due to incompatibility of the recipient somatic cell compartment with the germ cells from phylogenetically distant species12. An alternative approach is transplantation of germ cells from large species together with their surrounding somatic compartment. We first reported in 2002, that small fragments of testis tissue from immature males transplanted under the dorsal skin of immunodeficient mice are able to survive and undergo full development with the production of fertilization competent sperm13. Since then testis tissue xenografting has been shown to be successful in many species and emerged as a valuable alternative to study testis development and spermatogenesis of large animals in mice14.

Protocols for germ cell transplantation and testis tissue xenografting are described. Germ cell transplantation results in donor-derived spermatogenesis in recipient testes and represents a functional reconstitution assay for identification of spermatogonial stem cells (SSCs). Testis tissue xenografting reproduces testis development and spermatogenesis of various donor species in recipient mice.

Germ trapianto di cellule e di tessuto testicolare xenotrapianto in topi

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

In elettroporazione in vivo dei Morpholinos nella retina Zebrafish per adulti

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

In elettroporazione in vivo di Morpholinos nella Adult Regenerating Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...