Nous remercions Birgit Westernstroeer du Centre de médecine de la reproduction et d&#39;andrologie à l&#39;université de Münster pour l&#39;enseignement de la micro-injection des testicules. En outre, nous sommes reconnaissants à Ursula Antkewitz et Petra Reckling pour l&#39;assistance technique. Nous remercions la Fondation allemande pour la recherche (DFG) pour soutenir ce travail (WE2458 / 10-1). &#160;

Toutes les expériences pratiquées sur les animaux ont été approuvés par le comité d&#39;éthique local (Landesamt für Landwirtschaft, Lebensmittelsicherheit und Fischerei, Mecklenburg-Vorpommern, Allemagne). Préparation 1 Le plasmide <ol><li> Pour la préparation de plasmides, en utilisant des kits de purification de plasmide (voir le tableau des matériaux) ou des procédés similaires avec du tampon d&#39;élimination de l&#39;endotoxine de sorte que les réactions immunitaires de l&#39;animal peuvent être évités. Suivez les instructions du manuel. Employer le trou DDH 2 O pour diluer la solution de plasmide. </li><li> Eliminer les débris par filage de la solution d&#39;ADN à la vitesse maximale (20 000 xg). Ramassez ensuite le surnageant. </li><li> Déterminer la concentration de plasmide avec un spectrophotomètre (voir matériaux). </li><li> Ajuster la concentration de plasmide avec le trou DDH 2 O à 1-3 mg (ADN) / pl. Remarque: Des concentrations plus faibles réduisent l&#39;efficacité de la transfection, tandis que des concentrations plus élevées peuvent causer une solution d&#39;injection trop visqueux. En outre, leefficacité de la transfection dépend de la taille du plasmide et doit être testé individuellement. </li><li> Avant l&#39;injection, préparer un mélange de la solution avec 40 ul par exemple plasmide avec 5 ul de PBS (10x) et 5 ul vert Fast (0,5%). Fast Green est nécessaire pour suivre le processus d&#39;injection. De préférence, utiliser murales PCR tubes minces ou parafilm. Remarque: En fonction de la taille des testicules, un volume de 20-50 ul sera nécessaire pour chaque testicule. </li></ol> 2 Préparation de microinjection Pipette <ol><li> Utilisez capillaires de borosilicate (voir le tableau des matériaux) pour préparer les pipettes de micro-injection. </li><li> Tirez capillaires en verre avec un extracteur capillaire vertical (voir le tableau des matériaux). </li><li> Casser l&#39;embout capillaire avec une pince par baguage doucement. La pointe est habituellement de 50-80 m de diamètre et d&#39;environ 1 cm de long. Essayez d&#39;éviter les trucs trop longtemps que ceux-ci ne sont pas suffisamment rigide pour pénétrer dans le tissu. </li><li> Enfin, sharpen la pointe dans un 30 ° à 45 ° en utilisant un chanfreiner micropipette (voir le tableau des matériaux). </li><li> Charger la pipette de micro-injection avec le mélange de solution d&#39;injection (voir la préparation de plasmide). Appliquer la solution à l&#39;arrière du capillaire de verre avec une petite seringue. Remarque: Essayez d&#39;éviter les bulles d&#39;air pendant le chargement, sinon ceux-ci seront injectés dans les tubes séminifères avec la solution de plasmide et donc permettra de réduire la conductivité ainsi que, éventuellement endommager les tissus. </li></ol> 3. anesthésie et de chirurgie <ol><li> Effectuer l&#39;opération dans des conditions aseptiques en utilisant du matériel stérile (c&#39;est à dire, seringue, aiguille, instruments chirurgicaux, etc). Cela permettra de réduire l&#39;infection de l&#39;animal et assurer un bon taux de survie. </li><li> Utilisez la souris mâle pour l&#39;électroporation à l&#39;âge de 6-8 semaines. </li><li> Pour initialiser un traitement analgésique postopératoire, fournir la souris avec des analgésiques ajouté de l&#39;eau potable le jour avant la chirurgie. Ce culUres une analgésie préventive en cas de hypodipsia postopératoire hautement probable (de la consommation d&#39;eau a diminué). A cet effet, exposer à de l&#39;eau potable, par exemple une suspension d&#39;ibuprofène pédiatrique (20 mg / ml). L&#39;eau de boisson médicamenteuse doit également être fourni le jour un et deux de la reprise en concentration identique. Cela signifie une dose journalière de 7,5 mg / kg, valable pour poids de 25 g âgés de 8 semaines C57BL / 6J 5 ml / j d&#39;eau potable et le corps. Ce schéma analgésique garantit un soulagement de la douleur fondamentale et récupération accélérée avec bien-être élevé 26. </li><li> Pour la préparation de l&#39;anesthésie solution sur le jour de la chirurgie de travail, mélanger 10% kétamine et xylazine 2% dans un rapport 1: 1 (voir le tableau des matériaux). </li><li> Pour anesthésier l&#39;animal, appliquer 0,25 ml / 100 mg de poids corporel de manière sous-cutanée de la solution anesthésique (Kétamine = 0,125 g / kg; xylazine = 0,025 g / kg). Utilisez un site d&#39;injection entre pelvienne et des membres afin d&#39;éviter des dommages aux organes et les blessures de la souris. Habituellement, 10 min sont nécessairesjusqu&#39;à ce que la souris est profondément anesthésié. </li><li> Couvrir les yeux de souris avec une pommade vétérinaire pour prévenir la sécheresse pendant l&#39;anesthésie. </li><li> Testez arrestation anesthésie profonde, qui est perceptible par une absence totale de réponse. A cet effet, il suffit de pincer le bout de l&#39;animal. </li><li> Pour démarrer l&#39;opération, enlever les poils abdominale avec un rasoir électrique ou similaire et désinfecter la zone de coopération avec sterilium ou similaire. </li><li> Faire une incision ventrale directement au-dessus des glandes préputiales dans le centre de la région abdominale. A cet endroit, tirez d&#39;abord la peau loin et mener une petite incision cutanée transversale d&#39;environ 8-14 mm. Ensuite, continuer le long de la couche musculaire abdominale. Remarque: La lésion doit être aussi faible que possible pour réduire les méfaits de l&#39;animal. </li><li> Tirez vers le haut les coussinet adipeux abdominal soin d&#39;exposer les testicules ci-joint et le placer sur une feuille de papier jetable étanche drap préparé avant juste à côté du site de l&#39;incision (figure 2A). </li><li> Utilisez des jumelles à find conduit efférent en tant que cible d&#39;injection. La persistance du canal efférent est identifié comme la jonction de beau navire entre le testicule et l&#39;épididyme. Il est situé à proximité de l&#39;artère testiculaire important. La persistance du canal fonctionne presque à un angle de 45 ° par rapport à l&#39;artère et visiblement entre l&#39;épididyme habitant. Remarque: En fonction de l&#39;âge de la souris, le canal artériel est habituellement enterré dans les tissus gras. </li><li> Utiliser des pinces fines pour dégager le conduit efférent de ce tissu adipeux. Après cela, placez le canal artériel publié sur une bande de papier stérile pour assurer la visibilité de la conduite sans nuire environnement. </li></ol> La micro-injection d&#39;ADN et 4 application <ol><li> Raccorder la pipette de micro-injection, qui est chargé avec la solution de plasmide à l&#39;unité micromanipulateur / injecteur. </li><li> Placez l&#39;aiguille de microinjection parallèle au canal efférent avec la pointe dirigée vers le rete testis (figure 2B). </li><li> Utilisez des pinces fines etdépouiller le conduit sur la pipette de micro-injection. Assurez-vous que le capillaire est maintenue parallèle à la conduite tout en tirant sur. Remarque: Cette procédure est plus pratique que pénétrer dans le navire en déplaçant l&#39;aiguille avec le micromanipulateur. </li><li> Dirigez l&#39;aiguille délicatement vers le testicule et arrêter tout comme il pénètre dans le rete testis directement sous l&#39;albuginée (figure 2C). Remarque: Dans le cas de trop loin le rete testis, la solution de plasmide fuite dans l&#39;espace interstitiel. Cela conduit souvent à l&#39;échec de la transfection des tubes séminifères. </li><li> Pour l&#39;injection de la solution de plasmide d&#39;utiliser le micro-injecteur ayant les paramètres suivants: pi: 100 hPa, Ti: 0,2 sec, et pc: 0 hPa (figure 2D). </li><li> Surveiller le processus d&#39;injection ensemble en observant le testicule état de remplissage à l&#39;aide de la couleur verte. Veillez à ce que le testicule est seulement rempli jusqu&#39;au 2/3 de son volume avec le plsolution asmid (figure 2E). Remarque: Si le volume d&#39;injection est dépassée, le tissu testiculaire pourrait en être affectée. </li></ol> 5. électroporation de testicule <ol><li> Pour activer l&#39;électroporation efficace, faire tremper les électrodes de pincement dans du PBS (1x). Cela garantit conductance adéquate. </li><li> Douceur serrer les testicules entre les électrodes humides (figure 2F). Mesurer la résistance électrique avec la électroporateur. </li><li> Appliquer un courant au testicule. Effectuer huit impulsions carrées de 40 V à 4 sites différents testicules avec une durée constante de 50 temps d&#39;impulsion ms et 950 ms de temps d&#39;intervalle. </li></ol> 6. fermeture de la plaie et post-chirurgie <ol><li> Lorsque l&#39;électroporation est terminé, placez le testicule de retour dans l&#39;emplacement d&#39;origine. </li><li> Coudre la couche musculaire interne avec une suture chirurgicale (voir matériaux). </li><li> Fermez la peau en utilisant des agrafes (voir le tableau des matériaux). Tirez la peau jusqu&#39;à l&#39;esprith pincettes. Assurez-vous d&#39;exclure la couche musculaire. Placer les agrafes, chacune à une distance d&#39;au plus 5 mm. Remarque: En raison suture chirurgicale peut être avalé par la souris, des agrafes sont favorisés pour la fermeture de la lésion de la peau. </li><li> Laissez la souris pour récupérer de l&#39;anesthésie sur des serviettes en papier stériles placées dans une cage de la souris vide stérile, qui est tempéré sur un bloc chaud 37 ° C. Le tampon doit assurer le réchauffement de la moitié de la cage de créer un gradient thermique. Pour l&#39;animal ce fait possible de rechercher la zone individu, plus confortable du fait de la variété de la température dans la cage. En outre, couvrir la souris avec une feuille de papier essuie stérile afin que le stress peut être encore réduit. Remarque: Depuis la surface du corps de la souris est peu commune par rapport à la masse, par rapport à des animaux plus gros, le support thermique est d&#39;une importance particulière pour la récupération réussie de rongeurs. </li><li> Lorsque l&#39;animal opéré a pleinement éveillé, transférer it pour poursuite de la reprise à une nouvelle cage de la souris entièrement équipée. Ne pas mettre l&#39;animal à son ancienne cage ou à un groupe de souris. Assurez-vous que la cage est aussi propre et stérile que possible pour prévenir l&#39;infection de la plaie. Surveiller le processus de récupération. Pour transférer de la souris vers l&#39;établissement de soins des animaux, attendre jusqu&#39;à ce qu&#39;il ait totalement récupéré et semble se comporter normalement. Remarque: personne supplémentaire et les stratégies post-chirurgicales sont discutées par Pritchett-Corning KR et al 6. </li></ol>

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Marten Michaelis, Alexander Sobczak, et Joachim M. Weitzel employés à l&#39;Institut de biologie de la reproduction, de l&#39;Institut Leibniz pour la biologie des animaux de ferme (FBN), Dummerstorf, Allemagne, déclarent qu&#39;ils n&#39;ont aucun intérêt financier concurrents.

La recherche dans le domaine de la biologie de la reproduction, en particulier dans le domaine de la fertilité masculine et la spermatogenèse inévitablement s&#39;appuie sur les organismes vivants. Afin d&#39;examiner la fonction testiculaire, pas de système de culture cellulaire adéquate / in vitro a été mis en place capable de réfléchir tous les essentiels des changements morphologiques d&#39;un spermatogonies diploïdes à un 1,2 spermatozoïde haploïde maturité. Ainsi,...

Spermatogenèse chez les mammifères est considérée comme un processus complexe de cellules souches auto-renouvelables subissant successivement la mitose, la méiose et de la différenciation, afin de développer dans les spermatozoïdes matures haploïde. Ces changements morphologiques sont orchestrées par différents types de cellules et malgré les tentatives profondes, il est encore impossible d&#39;imiter ces procédés en culture cellulaire 1,2. Par conséquent, la recherche sur la spermatogenèse jusqu&#39;à maintenant s&#39;appuie sur les organismes vivants comme des modèles in vivo. En général, les études de la fonction des gènes sont généralement basées sur les animaux transgéniques. Toutefois, la génération et le maintien de ce type de modèle animal est long, coûteux et très élaboré. Ceci est attribué au processus de sélection de temps nécessaire pour générer et maintenir le transgène au cours des générations. En outre, la manipulation génétique de l&#39;organisme tout entier par l&#39;approche transgénique ou knock-out est sujette à provoquer des déficiences physiologiques lors du ciblage gènes avec des fonctions essentielles dans plusieurs régions, par exemple, en dehors du testicule ou systémique. En outre, certaines méthodes de transfection transitoire sont associés à des inconvénients cruciaux. Par exemple, les inconvénients typiques de transfert de gène à médiation virale sont la provocation possible de réactions immunologiques et des règlements de sécurité supplémentaires, tandis que la lipofection 3 et 4 bombardement de microparticules peut endommager les tissus et sont limités à une certaine profondeur de la cellule pour l&#39;efficacité de la transfection suffisante. En revanche, l&#39;électroporation (EP) comme une autre façon commune de transfection transitoire, semble constituer une technique prometteuse pour permettre à la transfection in vivo et in vivo une analyse cohérente de gène. En général, les PE est désigné comme un phénomène dynamique qui dépend de la tension transmembranaire locale avec des pores au sein médiées par conséquent nanosecondes. Ces écarts peuvent être maintenus pendant millisecondes, suffisant to accorder l&#39;accès à l&#39;ADN, de l&#39;ARN ou des petites molécules 5. Lorsque la tension appliquée est trop élevée, le caractère généralement transitoire de PE est compensée en raison de la production de chaleur et de perméabilisation induction trop étendu avec en conséquence des dommages irréversibles de la cellule 5. Ici, nous montrons que l&#39;électroporation est un système de transfection efficace et économique, qui est capable d&#39;être utilisé pour la transformation génétique du testicule afin d&#39;élucider les caractéristiques de gène in vivo sur les testicules. Cet article traite de la préparation de plasmides, la micro-injection par le conduit efférent et l&#39;électroporation ultérieure de testicule de souris. Cette procédure peut être le moyen de choix pour atteindre transfection rapide, spécifique et efficace des tubes séminifères des testicules de souris in vivo afin d&#39;étudier les processus de la spermatogenèse.

<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

Cette vidéo et l&#39;article contribution donne une description complète de micro-injection et l&#39;électroporation des testicules de souris in vivo. Cette technique de transfection particulier pour les cellules testiculaires de souris permet l&#39;étude des processus uniques de la spermatogenèse. Le protocole qui suit se concentre sur la transfection de cellules testiculaires de souris avec des constructions plasmidiques. Plus précisément, nous avons utilisé le vecteur rapporteur pEGFP-C1, qui exprime renforcée protéine fluorescente verte (eGFP) et aussi le vecteur pDsRed2-N1 exprimant la protéine fluorescente rouge (DsRed2). Les deux gènes rapporteurs codés étaient sous le contrôle du promoteur du cytomegalovirus humain précoce immédiat (CMV). Pour effectuer un transfert de gène dans les testicules de souris, les constructions de plasmides rapporteur sont injectés dans les testicules de souris vivantes. À cette fin, les testicules d&#39;un animal anesthésié est exposé et le site de micro-injection est préparée. Notre lieu privilégié deinjection est conduit efférent, avec le rete testis finalement connecté en tant que voie de transport anatomique des spermatozoïdes entre le testicule et l&#39;épididyme. De cette manière, le remplissage des tubes séminifères après la micro-injection est très bien gérée et contrôlée grâce à l&#39;utilisation de solutions d&#39;ADN colorées. Après avoir observé un remplissage suffisant du testicule par sa structure de tube de couleur, l&#39;organe est soumis à une électroporation. Ceci permet le transfert de la solution d&#39;ADN dans les cellules testiculaires. À la suite de 3 jours d&#39;incubation, le testicule est enlevé et étudiée au microscope à fluorescence verte ou rouge, ce qui illustre le succès de la transfection. Généralement, ce protocole peut être utilisé pour fournir l&#39;ADN ou l&#39;ARN construit dans les testicules de souris vivante pour (sur) exprimer ou abattre des gènes, ce qui facilite l&#39;analyse de la fonction des gènes in vivo. En outre, il est adapté pour l&#39;étude de constructions rapporteurs ou règlements de gène putatiféléments conservateurs. Ainsi, les principaux avantages de la technique d&#39;électroporation sont des performances rapides en combinaison avec peu d&#39;effort ainsi que l&#39;équipement technique modérée et les compétences requises par rapport à d&#39;autres techniques.

Le cadre expérimental pour réaliser une micro-injection et l&#39;électroporation des testicules de souris in vivo tel qu&#39;il est utilisé selon le protocole est illustré à la figure 1. Même si il est possible d&#39;acquérir micropipettes fabriqués industriellement, nous avons préféré pour générer nos propres pipettes en tirant (figure 1A ) et le biseautage (figure 1B) des capillaires en verre de sorte que qu&#39;ils équipés nos besoin...

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

Cet article décrit la micro-injection et l&#39;électroporation de testicule de souris in vivo en tant que technique de transfection de cellules testiculaires de souris pour étudier les procédés uniques de la spermatogenèse. Le protocole présenté implique des étapes de préparation capillaire en verre, la micro-injection par le conduit efférent, et la transfection par électroporation.

In Vivo L&#39;électroporation et microinjection de testicule de souris

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

27.7K vues.  Leibniz Institute for Farm Animal Biology (FBN).  Cet article décrit la micro-injection et l'électroporation de testicule de souris in vivo en tant que technique de transfection de cellules testiculaires de souris pour étudier les procédés uniques de la spermatogenèse. Le protocole présenté implique des étapes de préparation capillaire en verre, la micro-injection par le conduit efférent, et la transfection par électroporation. 

Vidéo: 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. 

En injection intraventriculaire Utero et l&#39;électroporation d&#39;embryons de souris E15

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

Recherche

Biologie

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.

L&#39;électroporation cellule unique in vivo dans le cerveau intact développement

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.

En électroporation in vivo et ex utero pour l&#39;expression génique dans les cellules ganglionnaires de la rétine de la souris

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 d&#39;ADN des mammifères muscles squelettiques à l&#39;aide In Vivo L&#39;électroporation

 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.

L&#39;imagerie in vivo et des traitements thérapeutiques dans un modèle de souris orthotopique de cancer des ovaires

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.

L&#39;activation de l&#39;apoptose par micro-injection cytoplasmique du cytochrome 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.

Évaluation microinjection des ovocytes de souris, la maturation et de ploïdie

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.

Transplantation des cellules germinales et de tissus testiculaire xénogreffe chez la souris

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.

En électroporation in vivo de morpholinos dans la rétine de poisson zèbre adulte

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.

En électroporation in vivo de morpholinos dans la régénération des adultes 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 ...