Gli autori ringraziano Sharmin Hossain per il time-lapse crescita film cattura di uno stato immaturo neurone e Derek Dunfield per le immagini di microscopia elettronica di SCE suggerimenti micromicropipette.

Attrezzature di set-up: <ol><li> Le apparecchiature elettriche richieste per questa tecnica sono uno stimolatore, un oscilloscopio e un supporto micropipetta dotato di un filo d&#39;argento per essere inserito nella micropipetta. Usiamo tipicamente un assone strumenti Axoporator 800A stimolatore, tuttavia altri stimolatori comuni come gli strumenti Grass SD9 stimolatori sono adatti anche per singola cellula elettroporazione. Lo stimolatore è collegato al headstage, che si interfaccia con l&#39;elettrodo filo d&#39;argento per essere collocato all&#39;interno del micropipetta di vetro contenente una soluzione interna direzione d&#39;orchestra. Il piombo altri lo stimolatore è collegata all&#39;ingresso oscilloscopio, che riceve in ingresso anche da terra girino esterno. Il terreno girino esterno è semplicemente un filo d&#39;argento che rimane in contatto con il elettrotonici girino durante elettroporazione. Il circuito è completo una volta che la micropipetta viene posto in contatto con il girino. </li><li> Elettroporazione singola cellula richiede un microscopio con una buona distanza di lavoro (almeno 5cm), dal momento che questo dà spazio di manovra e permette la micropipetta di essere portato in un angolo di circa 30-45 gradi. </li></ol> Fabbricazione di micropipette: Preparazione di micropipette appropriato è un passo fondamentale in questo protocollo. La difficoltà sta nel bilanciare una punta stretta (meno di 1μm di diametro) con uno che non sarà facile rompere quando si fora il tessuto. Abbiamo anche scoperto che le punte con un angolo di cono maggiore di 10 gradi sono preferibili. <ol><li> Preparare una micropipetta di vetro tirato con una punta di diametro inferiore a 1μm, e un angolo di punta del cono superiore a 10 gradi. <ol><li> Vetro borosilicato con un filamento interno con diametro esterno di 1,5 mm e diametro interno di 0,75 mm è adatto per questo scopo. <ol><li> Il vetro relativamente spesso fornisce un grado di rigidità alla punta. </li><li> Il filamento interno è importante per l&#39;elaborazione di back-riempiti liquido verso la punta micropipetta. </li></ol></li></ol></li><li> Fabbricazione di punte adatte spesso richiede alcune modifiche sulla base dei risultati osservati. Diversi tessuti possono richiedere una punta con una forma leggermente diversa o diametro della punta. </li></ol> Elettroporazione di singole cellule protocollo: Per i nuovi utenti, è consigliabile iniziare con tinture fluorescenti destrano poiché, a differenza geneticamente codificato proteine ​​fluorescenti, possono essere viste immediatamente in epifluorescenza. Uso di coloranti fluorescenti permette all&#39;utente di vedere direttamente la posizione della punta della micropipetta all&#39;interno del tessuto, e su elettroporazione dà un feedback immediato sul fatto che l&#39;apparecchiatura è stata impostata correttamente e se la punta della micropipetta è appropriato. <ol><li> Riempire micropipetta con una soluzione di composto da elettroporate mediante l&#39;utilizzo di una siringa di carico o di back-filling. <ol><li> Se si utilizza DNA plasmidico, affinché i campioni siano endotossina-free, e ad una concentrazione tra 1-2μg/ml, preparati in acqua. </li><li> La concentrazione di coloranti fluorescenti destrano dovrebbe essere empiricamente determinato. Spesso usiamo coloranti fluorescenti diluito in acqua a circa 300μM. </li><li> Tipicamente, i volumi di 0,5-1ml sono sufficienti. </li><li> Prima di essere caricato, centrifugare brevemente le soluzioni ad alta velocità per ridurre la quantità di residui particolati trascinati nella micropipetta, che possono portare all&#39;ostruzione. </li><li> Le bolle d&#39;aria all&#39;interno della punta spesso colpiscono il flusso di corrente e dovrebbero essere soppiantate da vigorosamente sfogliando la micropipetta prima del montaggio sul headstage. </li></ol></li><li> Inserire nel supporto micropipetta o headstage montato su un 3-asse micromanipolatore. Assicurarsi che l&#39;elettrodo filo d&#39;argento è inserito nel micropipetta, ed è in contatto elettrotonici con la soluzione interna. </li><li> Anestetizzare girini per immersione in 0,02% MS-222 (3-aminobenzoico etil estere) preparati in media allevamento girino (soluzione di Steinberg, pH 7,4). <ol><li> Normalmente usiamo stadio 44-48 albino Xenopus laevis girini nei nostri esperimenti. </li><li> Girini di solito sono completamente anestetizzati dopo circa 5 minuti. </li><li> Girini Numerosi possono essere anestetizzati contemporaneamente per ridurre il tempo di attesa, tuttavia, evitare l&#39;esposizione a MS-222 per periodi oltre 1 ora. </li></ol></li><li> Trasferire un girino solo anestetizzati alla camera di elettroporazione utilizzando una micropipetta di plastica di trasferimento. <ol><li> La camera di elettroporazione può essere facilmente fatta in casa da un girino scultura a forma di cavità in un piccolo Sylgard ® blocco di silicone, e l&#39;inserimento di un filo d&#39;argento di terra nella cavità in modo tale che sarà in contatto con il elettrotonici girino nella camera. </li></ol></li><li> Girino lato la posizione supina per usare un pennello umido. </li><li> Abbassare la micropipetta fino a quando non è in contatto con la pelle, direttamente adiacente alla regione di interesse. <ol><li> Destinata alle regioni con i corpi cellulari densamente notevolmente aumentare le probabilità di successo elettroporazione.Noi di solito bersaglio il tectum girino ottica. </li><li> Portare la micropipetta in un angolo di 30 a 45 gradi. Traiettorie più basse rendono più difficile perforare la pelle, mentre gli angoli più ripidi spesso ostacolano la propria vista. </li></ol></li><li> Continua l&#39;abbassamento della micropipetta inizialmente fossetta sulla pelle, e poi passare attraverso al tessuto sottostante. <ol><li> Cellule superficiali sono da preferire per imaging in vivo, e può essere bersaglio di modo che la punta della micropipetta si abbassa lentamente su fossette di pelle. </li></ol></li><li> Si deve monitorare costantemente la resistenza micropipetta una volta inserito all&#39;interno del tessuto. Se si utilizza un Axoporator 800A (Axon Instruments), resistenze di circa 10-40MΩ sono ottimali. La resistenza è un buon indicatore se il diametro della punta è appropriato per singola cellula elettroporazione. <ol><li> Resistenze micropipetta alta (&gt; 100M) sono indicativi di consigli intasato o micropipette con punte che sono troppo strette. </li><li> Resistenze micropipetta bassa (&lt;10MΩ) sono indicativi di un diametro di punta che è troppo grande, spesso il risultato di una punta rotta. </li><li> Se si utilizza uno stimolatore che non fornisce una misura diretta resistenza, la resistenza micropipetta può indirettamente misurati osservando l&#39;ampiezza degli impulsi di tensione misurata dal oscilloscopio (discussa più avanti). </li></ol></li><li> Applicare treno tensione. <ol><li> Per DNA plasmidico, treni di impulsi trovato per essere più efficaci consistono in negativo impulsi ad onda quadra di 1 ms di durata, consegnato a 300Hz con durata treno di impulsi di 500 ms. <ol><li> La tensione degli impulsi è determinato empiricamente in modo che l&#39;ampiezza di impulsi misurata sull&#39;oscilloscopio è compresa tra 0,75 e 1.5μA. </li></ol></li><li> Per i coloranti fluorescenti destrano, positivi impulsi ad onda quadra di durata 300μs sono consegnati ad una frequenza di 300Hz con durata treno di impulsi di 10ms. <ol><li> Come per il protocollo per il DNA, la tensione di impulso è determinato empiricamente in modo che l&#39;ampiezza di impulsi misurata sull&#39;oscilloscopio è compresa tra 0,75 e 1.5μA. </li><li> Ulteriori modifiche ai parametri di stimolo può essere fatta sulla base osservando i risultati di elettroporazione in epifluorescenza. <ol><li> Se le cellule elettroporate apparire debole, i parametri di impulso dovrebbe essere gradualmente aumentata fino a quando le cellule appaiono più luminosi. </li><li> D&#39;altra parte, se gruppi di cellule sono etichettati, i parametri di impulso deve essere ridotta fino a quando singole cellule sono etichettati. </li></ol></li></ol></li></ol></li><li> Ritrarre micropipetta, re-inserire in un sito diverso all&#39;interno del tessuto, e applicare treno tensione. <ol><li> Per migliorare la resa, di solito electroporate diversi siti ogni emisfero del tectum girino ottica con una spaziatura sufficiente a garantire che le cellule etichettate non si sovrappongano. </li></ol></li><li> Trasferire il girino elettroporate in un contenitore con la soluzione allevamento di Steinberg. <ol><li> Girini saranno rapidamente riprendersi dall&#39;anestesia in pochi minuti. </li></ol></li><li> La micropipetta stesso può essere utilizzato per numerosi girini. E &#39;importante osservare costantemente l&#39;ampiezza degli impulsi osservata sull&#39;oscilloscopio. <ol><li> Se l&#39;ampiezza dell&#39;impulso rimane bassa dopo aumentando significativamente l&#39;intensità degli impulsi questo è probabilmente il risultato della punta è intasato, visto spesso quando electroporating molti girini in una sola seduta. <ol><li> Un metodo per rimuovere uno zoccolo minore suggerimento è quello di applicare un impulso positivo solo dopo aver inserito la micropipetta nel tessuto. </li><li> Se lo zoccolo non può essere rimosso, o intasamento frequentemente ricorre, sostituire la micropipetta. </li></ol></li><li> Se l&#39;ampiezza dell&#39;impulso rimane molto grande dopo diminuendo in modo significativo i parametri degli impulsi, questo è un segno che la punta si è rotto e deve essere sostituito. </li></ol></li></ol> Screening per le cellule con successo elettroporate: <ol><li> Dopo l&#39;elettroporazione, i girini sono sottoposti a screening per le cellule etichettati utilizzando un microscopio a epifluorescenza verticale. </li><li> Nel caso di tinture fluorescenti, girini possono essere sottoposti a screening dopo appena 30 minuti dopo l&#39;elettroporazione. <ol><li> Abbiamo trovato, comunque, che intervalli più lunghi sono associati a livelli più bassi di fluorescenza di fondo. </li></ol></li><li> Per geneticamente codificato proteine ​​fluorescenti, lo screening si svolge generalmente almeno 12 ore dopo l&#39;elettroporazione. <ol><li> Cellule che esprimono proteine ​​fluorescenti continuerà ad avere più luminoso con il tempo, quindi le cellule dim possono essere nuovamente proiettati dopo un intervallo aggiuntivo di 12-24 ore. </li></ol></li><li> Girini sono anestetizzati come sopra descritto, posto in una camera di Sylgard ® e coperti. Girini individuali vengono poi esaminati al epifluorescenza. <ol><li> Si noti che l&#39;eccessiva esposizione alla luce epifluorescenza si tradurrà in fototossicità che può uccidere la cellula etichettati. Quindi, minimizzare la quantità di tempo che le cellule sono esposte alla luce fluorescente. </li></ol></li><li> Ritorna girini di contenitore con la soluzione allevamento di Steinberg. </li></ol>

<ol>
	<li>Bestman, J. E., Ewald, R. C., Chiu, S. L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+single-cell+electroporation+for+transfer+of+DNA+and+macromolecules.">In vivo single-cell electroporation for transfer of DNA and macromolecules.</a> Nature Protocols. 1, 1267-1272 (2006).</li><li>Dunfield, D., Haas, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+electroporation.">Single cell electroporation.</a> Encyclopedia of Neuroscience (4th Ed.). , .</li><li>Haas, K., Jensen, K., Sin, W. C., Foa, L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+electroporation+in+Xenopus+tadpoles+in+vivo--from+single+cells+to+the+entire+brain.">Targeted electroporation in Xenopus tadpoles in vivo--from single cells to the entire brain.</a> Differentiation. 70, 148-154 (2002).</li><li>Haas, K., Sin, W. C., Javaherian, A., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+electroporation+for+gene+transfer+in+vivo.">Single-cell electroporation for gene transfer in vivo.</a> Neuron. 29, 583-591 (2001).</li></ol>

Elettroporazione di singole cellule (SCE) è un potente strumento per determinare la funzione del gene e l&#39;esecuzione di mirati manipolazione genetica. La trasparenza del girino albino e l&#39;accessibilità del cervello fanno di questo sistema modello ideale per la visualizzazione di crescita neuronale e di eventi intracellulari all&#39;interno di un organismo vivo. SCE permette la visualizzazione della crescita di un singolo neurone, e di eseguire manipolazioni cellulari autonoma all&#39;interno di un cervello altrimenti inalterato. M...

single cell electroporation in vivo within the intact developing brain

Elettroporazione di singole cellule (SCE) è una tecnica specializzata che consentono la fornitura di DNA o di altre macromolecole in singole celle all&#39;interno del tessuto intatto, anche in preparati in vivo. Il vantaggio di questa tecnica è che le manipolazioni sperimentali può essere eseguita su singole cellule, lasciando inalterato il tessuto circostante, in modo da distinguere cellule effetti autonomi e da quelli risultanti da trattamenti globali. Se combinati con avanzate tecniche di imaging in vivo, SCE di marcatori fluorescenti permette la visualizzazione diretta della morfologia cellulare, la crescita cellulare, ed eventi intracellulari su scale temporali che variano da secondi a giorni. Mentre questa tecnica è utilizzata in una varietà di preparati in vivo ed ex vivo, abbiamo ottimizzato questa tecnica per l&#39;utilizzo in girini Xenopus laevis. In questo articolo video, abbiamo dettaglio la procedura di SCE di un colorante fluorescente o DNA plasmidico in neuroni all&#39;interno del cervello intatto l&#39;albino Xenopus girino. Abbiamo anche discutere i metodi per ottimizzare la resa, e mostrare esempi di vivere a due fotoni imaging di fluorescenza dei neuroni fluorescente da SCE.

Watch this Scientific Journal Video about Single Cell Electroporation in vivo within the Intact Developing Brain at JoVE.com

Single Cell Electroporation in vivo within the Intact Developing Brain

Elettroporazione di singole cellule (SCE) è una tecnica specializzata che consente la consegna di DNA o di altre macromolecole in singole celle all&#39;interno del tessuto intatto, anche in preparati in vivo. Qui dettaglio la procedura di SCE di un colorante fluorescente o DNA plasmidico in neuroni all&#39;interno del cervello intatto della Xenopus laevis girino.

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 ...

single-cell-electroporation-in-vivo-within-the-intact-developing-brain

Research

JoVE Journal

Biology

13.2K Views.  University of British Columbia - UBC. Elettroporazione di singole cellule (SCE) è una tecnica specializzata che consente la consegna di DNA o di altre macromolecole in singole celle all'interno del tessuto intatto, anche in preparati in vivo. Qui dettaglio la procedura di SCE di un colorante fluorescente o DNA plasmidico in neuroni all'interno del cervello intatto della Xenopus laevis girino.

Video: Single Cell Electroporation in vivo within the Intact Developing Brain

Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

Elettroporazione di micobatteri

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

Ricerca

Biologia

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 ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Unicellulari registrazioni di aspirazione da Fotorecettori Cone mouse

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Imaging in vivo di intatto Larve di Drosophila a livello sub-cellulare Risoluzione

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

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 ...

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

Imaging confocale delle singole mitocondriali superossido illumina di cuore intatto o<em&gt; In Vivo</em

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Valutazione del calcio Sparks in intatte fibre muscolari scheletriche

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

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 ...