Questo lavoro è stato sostenuto dal National Institutes of Health concede NS047726, NS072027, e AR052646.

I metodi in questo studio sono stati eseguiti in conformità con l&#39;etica Feinberg School of Medicine della Northwestern University Istituzionale Animal Care and Use Committee (IACUC) le linee guida approvate. Sono stati fatti tutti gli sforzi per ridurre al minimo le sofferenze. Procedura sperimentale 1. per elettroporazione in vivo del flessore Brevis (FDB) Muscle Bundle in mouse <ol><li> Plasmidi di progettazione per l&#39;espressione in vivo utilizzando un promotore saputo esprimere in cellule di mammifero (ad esempio, citomegalovirus, CMV) o nel muscolo (ad esempio, Muscolo Creatina chinasi, MCK) 6,7. Plasmidi più grandi possono esprimere a livelli inferiori. Il promotore CMV è stato ampiamente utilizzato 6,7. </li><li> Purificare plasmidi da E. coli utilizzando condizioni di libero endotossine le istruzioni del produttore. Sciogliere il plasmide in sterili, endotossine tampone TE libero a 2-5 mg plasmide / ml. </li><li> Diluire e un&#39;aliquota il plasmide di una concentration di 20 mg di DNA in 10 ml di endotossine sterile libero TE (2 mg / mL) tampone per iniezione in una provetta sterile. Preparare un&#39;aliquota per iniezione. </li><li> Diluire lo stock di ialuronidasi a 1x con sterile tampone fosfato senza calcio e magnesio (PBS - / -), dando una concentrazione finale di 8 unità. Aliquota 10 ml di ialuronidasi diluito per iniezione in una provetta sterile. </li><li> Anestetizzare il mouse utilizzando 2,5% isoflurano fino sedato e non risponde per la coda o pizzicare punta. Posto l&#39;animale in posizione supina su una piastra elettrica o una tabella di riscaldamento per mantenere la temperatura corporea normale. </li><li> Pulire la zampa del mouse con un programma alternato di uno scrub povidone iodio e alcol tre volte utilizzando una tecnica di preparazione proveniente dal sito di ago e irradia verso l&#39;esterno. Questa tecnica mantiene il sito ago in una condizione asettica, minimizzando introduzione patogeno. </li><li> Iniettare zampa di the mouse con 10 microlitri della soluzione 1x ialuronidasi (2 mg / ml = 8 unità) tra la pelle e il muscolo utilizzando una siringa sterile da 1 ml. Ialuronidasi digerisce componenti della matrice extracellulare per permettere l&#39;ingresso più efficiente del plasmide in miofibre. Invio alla base del tallone e far avanzare l&#39;ago verso le dita. Rilasciare lentamente il liquido mentre l&#39;ago viene retratto. Il sito di iniezione sotto la pelle si espanderà e inizia a diventare rosa. </li><li> Ripetere i punti 1.6 e 1.7 sul piede controlaterale. </li><li> Scollegare il anestesia e posizionare il mouse avanti nella gabbia. Lasciare che il mouse per riprendersi completamente dall&#39;anestesia. </li><li> Attendere 2 ore. </li><li> Ripetere le procedure di sterilizzazione e anestetizzante piede, punti 1.5 e 1.6. </li><li> Iniettare il DNA plasmidico diluita nella zampa nello stesso modo come la ialuronidasi. Assicurarsi che il volume massimo è di 20 ml o meno. Per l&#39;espressione dual plasmide iniettare 20 ug di ciascun plasmide con un volume massimo di 20 &amp;# 181; l o meno. </li><li> Ripetere l&#39;iniezione plasmide sulla zampa controlaterale o utilizzare il piede controlaterale per le iniezioni di controllo. </li><li> Sul piede che è stato inizialmente iniettato, sollevare la pelle dal muscolo sottolineare con pinza sottile e mettere un ago 27 G attraverso le sfere di pelle vicino le dita in posizione perpendicolare alla linea di guarire-punta del piede. Prendete un secondo sterili 27 ago G e posizionarlo in orizzontale attraverso la pelle sul tallone. Inserire aghi paralleli tra loro ~ 1 cm di distanza. </li><li> Utilizzando i morsetti a coccodrillo, elettrodi morsetto agli aghi paralleli assicurandosi gli aghi non siano a contatto tra loro. L&#39;uso della modellazione argilla può garantire i morsetti in posizione. </li><li> Collegare gli elettrodi di uno stimolatore elettrico. </li><li> Elettroporazione il muscolo applicando 20 impulsi, 20 msec di durata / ciascuno, a 1 Hz, 100 V / cm. Le dita possono contrarsi con la stimolazione. </li><li> Ripetere la procedura di stimolazione sul piede controlaterale, se necessario. </li><li>Rimuovere gli aghi. Pulire la zampa con iodio povidone macchia. </li><li> Spegnere l&#39;anestesia. Ritorna il mouse per la gabbia di recupero e monitorare l&#39;attività. Se la procedura è stata condotta come previsto, l&#39;animale deve riprendere il normale deambulazione entro 30 minuti e mostra alcuna differenza di deambulazione o un&#39;attività livello dal comportamento pre-iniezione. Pertanto, sono necessari analgesici post-procedura. Il recupero, rispedire l&#39;animale verso il vivaio. Nota: l&#39;espressione della proteina può essere saggiato fin da 48 ore dopo l&#39;elettroporazione. Tuttavia, per consentire il recupero più completo dopo l&#39;elettroporazione, noi preferiamo l&#39;isolamento e lo studio di miofibre 7 - 14 giorni dopo l&#39;elettroporazione 10. Espressione può essere saggiato fintanto 4 settimane dopo l&#39;elettroporazione. </li></ol> Procedura sperimentale per 2. Isolamento delle flessore Brevis (FDB) Fibre <ol><li> Collagenasi disgelo. Per ogni pacchetto FDB allestire due pozzetti in una piastra di coltura tissutale 12-bene. In un bene aggiungere 1 ml di pre-riscaldato Modified Eagles media Dulbecco più Bovine Serum Albumin (DMEM + BSA) e nell&#39;altra ben aggiungere 1 ml di pre-riscaldato soluzione Ringer. Inoltre, aggiungere freschi 10 ml di soluzione 1x Ringers ai 35 mm Sylgard piatto. </li><li> Sacrifica l&#39;animale attraverso CO 2 o inalazione di gas anestetico, seguita da dislocazione cervicale o un altro metodo per assicurare la morte. </li><li> Rimuovere le zampe posteriori sopra la caviglia con una lama di rasoio pulito e immergere in soluzione 1x Ringers nei 35 mm Sylgard piatto. </li><li> Pin un piede stretto, unico rivolto verso l&#39;alto, sul piatto Sylgard con 30 aghi da G a ciascuna delle punte 5 punta e alla caviglia. </li><li> Partendo alla caviglia, tagliare la pelle verso l&#39;alto il lato del piede lungo l&#39;attaccatura dei capelli verso le dita dei piedi, attenzione a non intaccare il muscolo sotto la pelle. Ripetere sull&#39;altro lato del piede. </li><li> Tenendo la pelle alla base della caviglia con una pinza sottile, tagliare la pelle attraverso il tallone. </li><li> Sollevamento la pellepatta con una pinza, puntare le forbici verso la pelle per non nick il muscolo. Tagliare il tessuto connettivo sottostante, rimuovere efficacemente la pelle. </li><li> Per rimuovere il fascio muscolare FDB tagliare il grande tendine del tallone per liberare il tendine dall&#39;osso. Tenendo il tendine con una pinza, sezionare il fascio FDB dal grande tendine bianco rimuovere accuratamente qualsiasi tessuto connettivo attaccato al FDB con le forbici. Se fatto correttamente il bundle sarà facile sollevare dei tendini sottostanti rivelano tendini bianchi brillanti che portano alle singole cifre. Tagliare la FDB ai tendini vicino le dita liberando il fascio dal piede. </li><li> Posizionare l&#39;intero fascio muscolare nel pozzo contenente DMEM + BSA. </li><li> Ripetere sul piede controlaterale. </li><li> Una volta che tutti i muscoli FDB sono isolate, aggiungere 100 ml di collagenasi in ciascun pozzetto contenente DMEM + BSA e muscoli. </li><li> Controllare il successo di elettroporazione con un microscopio a fluorescenza invertito prima della digestione. Se properl fattoy, verso l&#39;alto di 90% efficienza di trasfezione possono essere raggiunti. </li><li> Incubare la piastra in una umidificata, 37 ° C incubatore a 10% CO 2 per circa 1 ora. Il tempo di incubazione può variare a seconda del modello di malattia e lo sfondo e la sorte di collagenasi. </li><li> Dopo 1 ora, trasferire accuratamente il fascio FDB al pozzetto contenente unica soluzione di Ringer. </li><li> Triturare il fascio FDB nel pozzo con la soluzione Ringers con una pipetta 1 ml. Tagliare la fine della punta della pipetta con una lama di rasoio pulito in modo che il pacchetto può passare facilmente attraverso la punta della pipetta. Triturare delicatamente il muscolo (~ 15 volte) permettendo al fascio di passare su e giù parallelamente alla punta. Fibre intatte cadranno nella soluzione Ringers. Quando le fibre non facilmente cadere il fagotto, posizionare il muscolo posteriore in soluzione di collagenasi. </li><li> Piastra le fibre isolate sul piatto ~ 500 microlitri per piastra. </li><li> Lasciare il muscolo per digerire nel pozzo per 15 min. </li><li> Ripetere i punti 2.152.17 fino a fascio diminuisce dimensioni e la maggior parte delle fibre sono dissociato dal fascio (di solito un totale di 2-3 volte). Una volta dissociato il muscolo passa facilmente attraverso un puntale 1 ml. </li><li> Lasciate che le fibre si attaccano al piatto per un minimo di 15 min. Soluzione fresca Ringers può essere aggiunta alla piastra per diluire collagenasi residuo o cambiata a seconda vincoli di tempo. Nota: Le fibre sono ora pronti per l&#39;imaging. </li></ol> Procedura sperimentale 3. per la visualizzazione di laser-indotta danni della membrana utilizzando Microscopia confocale <ol><li> Fibre carico immediatamente prima di imaging con 300 ml di 2,5 micron FM 1-43 di FM 4-64 colorante in soluzione di Ringer. In generale, le fibre di immagine 15 minuti a 2 ore dopo l&#39;isolamento. Tuttavia, le fibre possono essere esposte fino a 24 ore dopo l&#39;isolamento. </li><li> Porre la capsula fondo di vetro su un microscopio confocale equipaggiato con un laser UV e un obiettivo 60X / 63X. </li><li> Individuare una fibra. Assicurarsi che la fibra è saldamente collegatoal piatto toccando la base del microscopio. Escludere le fibre che sono stati danneggiati nel processo di dissociazione. Le fibre danneggiate possono contenere blebs membrana, alti livelli di assorbimento di colorante FM, ricciolo o piegare intensamente. </li><li> Per indurre danni membrana sulla fibra, posizionare il sarcolemma al centro della finestra di esposizione con un campo libero di nuclei. Focus sul sarcolemma utilizzando il canale FM tintura in modo tale che il sarcolemma è estremamente tagliente. </li><li> Posizionare i capelli ROI croce sul sarcolemma utilizzando il canale FM 4-64. </li><li> Irradiare un punto di 1 pixel alla potenza di 80% per 3 secondi (~ 350 nm x 350 zona nm). Colorante FM entrerà nella cella nel sito di lesione. Potenza e ora possono essere modificate per aumentare o diminuire l&#39;area di insulto. </li><li> Cattura immagini della fibra prima danni, al momento del danno, ogni post-danni 2 sec, quindi ogni 10 sec per 5 min. Timing può essere regolata a seconda delle necessità. Le singole immagini possono essere convertite in filmati time-lapse in Image J. </li><li> Ripetere i passaggi3.3 attraverso 3.7 su un massimo di 3 fibre per piatto. </li></ol>

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L&#39;utilizzo di elettroporazione per studiare le proteine ​​fluorescenti tag in vivo è ideale per lo studio del muscolo in assenza di opportuni modelli di linee cellulari che riproducono fedelmente l&#39;intera struttura cellulare del muscolo. Questo protocollo descrive l&#39;utilizzo di elettroporazione e ad alta risoluzione microscopia confocale a fornire immagini dettagliate di localizzazione delle proteine ​​e la traslocazione dopo ferimento laser. Questo metodo può essere adattato per studiare ...

Miofibre individuali sono grandi, le cellule sinciziale altamente organizzati. Ci sono alcuni modelli di coltura cellulare per il muscolo; tuttavia, questi modelli hanno come principale limitazione che non fanno differenza completamente in fibre muscolari mature. Ad esempio, le linee cellulari C2C12 e L6 sono derivati ​​da topi e ratti, rispettivamente 1-4. In condizioni di deprivazione di siero, i mononucleari cellule &quot;mioblasti-like&quot; cessano proliferazione, subiscono il ritiro del ciclo cellulare, ed entrano nel programma miogenico che formano le cellule multinucleate con sarcomeri, indicati come miotubi. Con condizioni di coltura prolungata, miotubi possono esibire proprietà contrattili e &quot;contrazione&quot;, nella cultura. Linee cellulari umane hanno ormai stati istituiti 5. In aggiunta a queste linee cellulari immortalizzate, mioblasti mononucleate possono essere isolate dal muscolo e sotto le condizioni simili di deprivazione di siero formeranno miotubi. Queste linee cellulari e colture di mioblasti primari sono altamente usoFul perché possono essere trasfettate con plasmidi o trasdotte con virus e usate per studiare processi biologici cellulari di base. Tuttavia, queste cellule, anche se indotto a formare miotubi, mancano molte delle caratteristiche salienti di organizzazione muscolo maturo. In particolare, miotubi sono molto più piccoli singole fibre muscolari mature e non hanno la forma normale di miofibre. Criticamente, miotubi mancano trasversali (T-) tubuli, la rete membranosa richiesta per il rilascio del Ca 2+ efficiente in tutto il mioplasma. Un metodo alternativo per mioblasti primaria o linee cellulari miogeniche comporta usando fibre muscolari mature. Transgenesi può essere impiegato per stabilire espressione delle proteine ​​tag, ma questo metodo è costoso e richiede tempo. In vivo elettroporazione di muscoli del mouse è emerso come un metodo preferito per la sua velocità e affidabilità 6-10. Metodi per elettroporazione in vivo e l&#39;isolamento efficace di miofibre have stato ottimizzato per il mouse flessore brevis (FDB) muscolare 6. I metodi possono essere completate rapidamente e sono minimamente invasiva per indurre espressione vivo da plasmidi. Questo approccio è ora combinata con metodi di imaging ad alta risoluzione tra cui l&#39;imaging dopo interruzione del laser del 6,7 sarcolemma. La combinazione di coloranti fluorescenti e espressione di proteine ​​marcate fluorescenza può essere utilizzata per monitorare cellulari processi biologici miofibre mature.

<table>
	<tbody>
		<tr>
			<td>DMEM+A2:D32D42A2:D31AA2:D29</td>
			<td>Life Technologies</td>
			<td>11995-073</td>
			<td>Dissection Media 
			Dilution: 50ml +100g BSA</td>
		</tr>
		<tr>
			<td>Collagenase, Type II</td>
			<td>Life Technologies</td>
			<td>17101-015</td>
			<td>Fiber Digestion 
			Dilution: 160mg / 4ml DMEM (stock 40mg/ml aliquot)</td>
		</tr>
		<tr>
			<td>Falcon 12-well dishes</td>
			<td>Fisher Scientific</td>
			<td>877229</td>
			<td>Fiber Digestion</td>
		</tr>
		<tr>
			<td>Ringers Solution</td>
			<td></td>
			<td></td>
			<td>Fiber Digestion and Imaging 
			Dilution: 146 mM NaCl 
			5 mM&#160;&#160; KCl 
			2 mM&#160;&#160; CaCl2 
			1 mM&#160;&#160; MCl2 
			10mM HEPES 
			pH7.4 in H2O</td>
		</tr>
		<tr>
			<td>1ml Pipettes</td>
			<td>Axygen</td>
			<td>T-1000-C-R</td>
			<td>Digestion</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Razor Blades</td>
			<td>Personna</td>
			<td>94-120-71</td>
			<td>Digestion</td>
		</tr>
		<tr>
			<td>Precision Glide 30G needle</td>
			<td>BD Biosciences</td>
			<td>305128</td>
			<td>Dissection</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>1x PBS (-/-)</td>
			<td>Life Technologies</td>
			<td>14190-250</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Sylgard 184</td>
			<td>Fisher Scientific</td>
			<td>50-366-794</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>FST by Dumont</td>
			<td>#5/45</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Roboz</td>
			<td>RS-4913</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>World Precision Instruments</td>
			<td>500260</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A7906</td>
			<td>Dissection media 
			Dilution: 100mg / 50ml DMEM</td>
		</tr>
		<tr>
			<td>Falcon 60mm dishes</td>
			<td>Fisher Scientific</td>
			<td>08772B</td>
			<td>Dissection- used to hold sylgard</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Clay</td>
			<td></td>
			<td></td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Stimulator&#160;</td>
			<td>Grass</td>
			<td>s88x</td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Clamps</td>
			<td>Radio Shack</td>
			<td>270-356</td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Triadine</td>
			<td>Aplicare</td>
			<td>82-220</td>
			<td>Electroporation</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Precision Glide 27G needle</td>
			<td>BD Biosciences</td>
			<td>305136</td>
			<td>Electroporation</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Simulator</td>
			<td>Grass</td>
			<td>SIU-V</td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Gauze</td>
			<td>Covidien</td>
			<td>2146</td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Hyaluronidase</td>
			<td>Sigma</td>
			<td>H4272</td>
			<td>Injection 
			Dilution: Add 160ul PBS (1X) to 40&#181;l 5x stock</td>
		</tr>
		<tr>
			<td>1cc U-100 Insulin Syringe (28G1/2)&#160;</td>
			<td>BD Biosciences</td>
			<td>329420</td>
			<td>Injection</td>
		</tr>
		<tr>
			<td>Eppendorf Tubes</td>
			<td>Fisher Scientific</td>
			<td>02-682-550</td>
			<td>Injection</td>
		</tr>
		<tr>
			<td>35mm glass bottom Microwell dish</td>
			<td>MaTek</td>
			<td>P35G-1.5-14-C</td>
			<td>Microscopy</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>FM 4-64</td>
			<td>Life Technologies</td>
			<td>T-13320</td>
			<td>Microscopy 
			Dilution: Final 2.5 ug/ml</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>FM 1-43</td>
			<td>Life Technologies</td>
			<td>T-35356</td>
			<td>Microscopy 
			Dilution: Final 2.5 ug/ml</td>
		</tr>
		<tr>
			<td>Endotoxinfree Plasmid Maxi Kit</td>
			<td>Qiagen</td>
			<td>12362</td>
			<td>Plasmid Purification</td>
		</tr>
		<tr>
		</tr>
	</tbody>
</table>

dna electroporation, isolation and imaging of myofibers

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently labeled proteins in the flexor digitorum brevis (FDB) muscle. Conditions have been optimized to provide a large number of high quality myofibers expressing the electroporated plasmid while minimizing muscle damage. The method employs fluorescent tags on various proteins. Combining this expression method with high resolution confocal microscopy permits live cell imaging, including imaging after laser-induced damage. Fluorescent dyes combined with imaging of fluorescently-tagged proteins provides information regarding the basic structure of muscle and its response to stimuli.

Elettroporazione è una tecnica minimamente invasiva che introduce efficacemente purificato plasmide DNA nel flessore brevis (FDB) fascio muscolare (Figura 1A). Sette giorni dopo la transfezione fluorescently proteina tag viene visualizzato nella miofibre isolate (Figura 1B). Fibre isolate sono poi placcati e preparate per l&#39;imaging su un microscopio confocale. Le fibre possono essere sottoposti a lesione indotta da laser. Colorante FM può essere ut...

Watch this Scientific Journal Video about DNA Electroporation, Isolation and Imaging of Myofibers at JoVE.com

DNA Electroporation, Isolation and Imaging of Myofibers

This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure.

Elettroporazione DNA, isolamento e imaging di miofibre

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently ...

dna-electroporation-isolation-and-imaging-of-myofibers

Research

JoVE Journal

Biology

12.2K Views.  Northwestern University. This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure. 

Video: DNA Electroporation, Isolation and Imaging of Myofibers

Isolation of Genomic DNA from Mouse Tails

Isolamento del DNA genomico da Tails mouse

Ricerca

Biologia

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 ...

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 ...

Electroporation of Craniofacial Mesenchyme

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. For example, electroporation has been used with great success to study neural and retinal development in Xenopus, chicken and mouse 1-10. However, it is important to note that in all of these studies, investigators were not targeting soft tissues. Because we are interested in craniofacial development, we adapted a method to target facial mesenchyme.
When we searched the literature, we found, to our surprise, very few reports of successful gene transfer into cartilaginous tissue. The majority of these studies were gene therapy studies, such as siRNA or protein delivery into chondrogenic cell lines, or, animal models of arthritis 11-13. In other systems, such as chicken or mouse, electroporation of facial mesenchyme has been challenging (personal communications, Dept of Craniofacial Development, KCL). We hypothesized that electroporation into procartilaginous and cartilaginous tissues in Xenopus might work better. In our studies, we show that gene transfer into the facial cartilages occurs efficiently at early stages (28), when the facial primordium is still comprised of soft tissue prior to cartilage differentiation.
Xenopus is a very accessible vertebrate system for analysis of craniofacial development. Craniofacial structures are more readily visible in Xenopus than in any other vertebrate model, primarily because Xenopus embryos are fertilized externally, allowing analyses of the earliest stages, and facilitating live imaging at single cell resolution, as well as reuse of the mothers 14. Among vertebrate models developing externally, Xenopus is more useful for craniofacial analysis than zebrafish, as Xenopus larvae are larger and easier to dissect, and the developing facial region is more accessible to imaging than the equivalent region in fish. In addition, Xenopus is evolutionarily closer to humans than zebrafish (&tilde;100 million years closer) 15. Finally, at these stages, Xenopus tadpoles are transparent, and concurrent expression of fluorescent proteins or molecules will allow easy visualization of the developing cartilages. We anticipate that this approach will allow us to rapidly and efficiently test candidate molecules in an in vivo model system.

Craniofacial cartilages develop in close contact with other tissues and are difficult to manipulate in live animals. We are using electroporation to deliver molecular tools during growth of the craniofacial skeleton while bypassing early embryonic effects. This approach will allow us to efficiently test candidate molecules in vivo.

Elettroporazione di Mesenchima craniofacciale

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Due-e tridimensionale Live Cell Imaging di proteine ​​danno al DNA di risposta

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Isolamento e la coltura di miofibre individuali e delle loro cellule satelliti da adulto muscolo scheletrico

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

Analisi delle embrionali e larvali Zebrafish scheletriche miofibre da Preparazioni dissociata

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

Combinando singola molecola di manipolazione e di imaging per lo studio delle interazioni proteina-DNA

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

In tempo reale Imaging del singolo Engineered RNA trascritti in cellule viventi Utilizzando Ratiometric bimolecolari Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

Reduced-gravity Environment Hardware Demonstrations of a Prototype Miniaturized Flow Cytometer and Companion Microfluidic Mixing Technology

Until recently, astronaut blood samples were collected in-flight, transported to earth on the Space Shuttle, and analyzed in terrestrial laboratories. If humans are to travel beyond low Earth orbit, a transition towards space-ready, point-of-care (POC) testing is required. Such testing needs to be comprehensive, easy to perform in a reduced-gravity environment, and unaffected by the stresses of launch and spaceflight. Countless POC devices have been developed to mimic laboratory scale counterparts, but most have narrow applications and few have demonstrable use in an in-flight, reduced-gravity environment. In fact, demonstrations of biomedical diagnostics in reduced gravity are limited altogether, making component choice and certain logistical challenges difficult to approach when seeking to test new technology. To help fill the void, we are presenting a modular method for the construction and operation of a prototype blood diagnostic device and its associated parabolic flight test rig that meet the standards for flight-testing onboard a parabolic flight, reduced-gravity aircraft. The method first focuses on rig assembly for in-flight, reduced-gravity testing of a flow cytometer and a companion microfluidic mixing chip. Components are adaptable to other designs and some custom components, such as a microvolume sample loader and the micromixer may be of particular interest. The method then shifts focus to flight preparation, by offering guidelines and suggestions to prepare for a successful flight test with regard to user training, development of a standard operating procedure (SOP), and other issues. Finally, in-flight experimental procedures specific to our demonstrations are described.

Spaceflight blood diagnostics need innovation. Few demonstrations have been published illustrating in-flight, reduced-gravity health diagnostic technology. Here we present a method for construction and operation of a parabolic flight test rig for a prototype point-of-care flow-cytometry design, with components and preparation strategies adaptable to other setups.

Ridotto gravità ambiente hardware Dimostrazioni di un prototipo miniaturizzato Citofluorimetro e Companion Microfluidic tecnologia di miscelazione

Until recently, astronaut blood samples were collected in-flight, transported to earth on the Space Shuttle, and analyzed in terrestrial laboratories. If ...