Ringraziamo Susan Gerbi, Heidi Smith e David Lambert per la fornitura di materiale per testare in vivo sui centrifugazione Sciara coprophila e Ilyanassa obsoleta, rispettivamente. Ringraziamo i membri del laboratorio Welte per commenti sulla tecnica e manoscritto. Sviluppo e la raffinatezza del test in vivo centrifugazione è stato sostenuto da NIGMS concedere GM64687 a MAW. Le immagini mostrate in fig. 1 e fig. 5E, F sono stati precedentemente pubblicato 2 e sono ristampato qui con il permesso.

<ol>
	<li>Wieschaus, E., N&#252;sslein-Volhard, C., Roberts, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Looking+at+embryos.">Looking at embryos.</a> Drosophila: A Practical Approach. , 179-214 (1998).</li><li>Cermelli, S., Guo, Y., Gross, S. P., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+lipid+droplet+proteome+reveals+that+droplets+are+a+protein+storage+depot.">The lipid droplet proteome reveals that droplets are a protein storage depot.</a> Curr Biol. 16, 1783-1795 (2006).</li><li>Kiehart, D. P., Crawford, J. M., Montague, R. A., Sullivan, W., Ashburner, M., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+microinjection+of+Drosophila+embryos.">Quantitative microinjection of Drosophila embryos.</a> Drosophila Protocols. , 345-359 (2000).</li><li>LaJeunesse, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three+new+Drosophila+markers+of+intracellular+membranes.">Three new Drosophila markers of intracellular membranes.</a> Biotechniques. 36 (784-788), 790-790 (2004).</li><li>Schuh, M., Lehner, C. F., Heidmann, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Incorporation+of+Drosophila+CID/CENP-A+and+CENP-C+into+centromeres+during+early+embryonic+anaphase.">Incorporation of Drosophila CID/CENP-A and CENP-C into centromeres during early embryonic anaphase.</a> Curr Biol. 17, 237-243 (2007).</li><li>Clarkson, M., Saint, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+His2AvDGFP+fusion+gene+complements+a+lethal+His2AvD+mutant+allele+and+provides+an+in+vivo+marker+for+Drosophila+chromosome+behavior.">A His2AvDGFP fusion gene complements a lethal His2AvD mutant allele and provides an in vivo marker for Drosophila chromosome behavior.</a> DNA Cell Biol. 18, 457-462 (1999).</li><li>Welte, M. A., Gross, S. P., Postner, M., Block, S. M., Wieschaus, E. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+regulation+of+vesicle+transport+in+Drosophila+embryos:+forces+and+kinetics.">Developmental regulation of vesicle transport in Drosophila embryos: forces and kinetics.</a> Cell. 92, 547-557 (1998).</li><li>Tran, S. L., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=unpublished+observations.">unpublished observations.</a> , .</li><li>Rothwell, W. F., Sullivan, W., Sullivan, W., Ashburner, M., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+analysis+of+Drosophila+embryos.">Fluorescent analysis of Drosophila embryos.</a> Drosophila Protocols. , 141-157 (2000).</li><li>Guo, Y., Jangi, S., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organelle-specific+Control+of+Intracellular+Transport:+Distinctly+Targeted+Isoforms+of+the+Regulator+Klar.">Organelle-specific Control of Intracellular Transport: Distinctly Targeted Isoforms of the Regulator Klar.</a> Mol Biol Cell. 16, 1406-1416 (2005).</li><li>Meyer, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Overlapping+Functions+of+Argonaute+Proteins+in+Patterning+and+Morphogenesis+of+Drosophila+Embryos.">Overlapping Functions of Argonaute Proteins in Patterning and Morphogenesis of Drosophila Embryos.</a> PLoS Genet. 2, 1224-1239 (2006).</li><li>Shubeita, G. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Consequences+of+motor+copy+number+on+the+intracellular+transport+of+kinesin-1-driven+lipid+droplets.">Consequences of motor copy number on the intracellular transport of kinesin-1-driven lipid droplets.</a> Cell. 135, 1098-1107 (2008).</li><li>Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+lipid-droplet+transport+by+the+Perilipin+homologue+LSD2.">Regulation of lipid-droplet transport by the Perilipin homologue LSD2.</a> Curr. Biol. 15, 1266-1275 (2005).</li><li>Phadnis, N., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unpublished+Observations.">Unpublished Observations.</a> , .</li><li>Phadnis, N., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unpublished+Observations.">Unpublished Observations.</a> , .</li><li>Morgan, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+XXII:+The+redistribution+of+the+visible+materials+of+the+egg+by+centrifuging.">Chapter XXII: The redistribution of the visible materials of the egg by centrifuging.</a> Experimental Embryology. , (1927).</li><li>Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unpublished+Observations.">Unpublished Observations.</a> , .</li><li>Bownes, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Abnormal+oogenesis+and+embryogenesis+resulting+from+centrifuging+Drosophila+melanogaster+females.">Abnormal oogenesis and embryogenesis resulting from centrifuging Drosophila melanogaster females.</a> J Embryol Exp Morphol. 40, 65-81 (1977).</li></ol>

Nessun conflitto di interessi dichiarati.

Passaggi critici Assicurarsi che la concentrazione di agar è abbastanza alta. Se è troppo basso, l&#39;agar-agar si disintegrerà durante la centrifugazione, e gli embrioni saranno scoppiare o essere orientati in modo casuale. Se la concentrazione di agar è solo un po &#39;troppo basso o se l&#39;agar è bagnato (rimozione sufficientemente secchi o insufficiente di TSS eccesso al punto 4.3), gli embrioni galleggerà dai loro buchi durante la centrifugazione e diventare impropriamente oriented. Se l&#39;agar è versato troppo sottilmente, sarà anche il collasso o crack durante la centrifugazione. Assicurarsi che gli embrioni non hanno superato le fasi cellularization. In caso contrario, la separazione di organelli non funziona (Fig. 5C) dal organelli sarà sequestrato all&#39;interno di migliaia di singole cellule. Nei giovani embrioni allo stadio di una cellula, organuli sono liberi di migrare grandi distanze e si accumulano in base alla loro densità caratteristica. Per evitare di analizzare gli embrioni che sono troppo vecchio, assicurarsi di (i) comportare un pre-raccolta per permettere alle femmine di deporre le uova vecchie (passo 2,4), (ii) raccogliere embrioni per 3 ore o meno (punto 2.5), (iii) visivamente selezionare gli embrioni prima di fasi cellularization per includerle in agar (punto 4.2), (iv) tenere incorporando poco tempo per prevenire lo sviluppo fino alla fine di cellularization (punto 4.6), e (v) scartare gli embrioni che non hanno strato correttamente (passo 5.6) . Non lesinare sul tempo di centrifugazione. Anche se alcuni degli embrioni mostrerà stratificazione piacevole anche dopo soli 10 minuti di centrifugazione, la separazione è incoerente dall&#39;embrione embrione. 30 giri min dare risultati altamente coerenti. Inoltre, più lungo è il tempo di centrifugazione, più a lungo i livelli sono stabili dopo la centrifugazione è terminata. Questo è importante per quelle applicazioni in cui ci vuole tempo per elaborare gli embrioni prima di essere utilizzati (per esempio, se hanno bisogno di essere trattati per esperimenti di trapianto o montato per l&#39;analisi confocale). Eventuali modifiche Raccolta degli ovuli e corion rimozione Molti protocolli differenti esistono per i punti 2 e 3 1, 3, 9, e funzionerà così come quelli descritti qui fino a quando gli embrioni utilizzati sono giovani, resa uovo è alto, e la frazione di mis- embrioni in scena è ridotta al minimo. Antero-posteriore allineamento Il protocollo orienta tutti gli embrioni tale che l&#39;estremità posteriore è sepolto nel agar e la fine anteriore è alto (Fig. 3). Questo orientamento è un bene per coerenza, in modo che sia possibile utilizzare il micropilo come punto di riferimento che finiscono rivolta verso l&#39;alto durante la centrifugazione (Fig. 1, 5). Tuttavia, con la pratica, è facile individuare l&#39;orientamento degli embrioni centrifugati dalla comparsa distinte del lipidi goccia e strati tuorlo da campo chiaro microscopia. Allora diventa possibile montare sia con gli embrioni finiscono, questa flessibilità accelera il processo di montaggio. Centrifugazione Unoriented L&#39;aspetto più-in termini di tempo e tecnicamente impegnativo del protocollo è di montaggio degli embrioni in agar (punto 4). Si richiede di toccare ogni embrione due volte (una volta per inserirlo nel foro e una volta per recuperarlo dopo la centrifugazione). In alternativa, si può trasferire gli embrioni a passo 4,1 in provette da microcentrifuga pieno di TSS. Quando questi tubi vengono centrifugati in una microcentrifuga (13.000 giri al minuto, 10-30 min), gli embrioni anche tipicamente strato di ben 10-13. Questa variante permette di processare centinaia di embrioni, allo stesso tempo molto rapidamente. Tuttavia, la stratificazione non è così consistente, una parte degli embrioni non si sviluppano strati distinti, nonostante le forze di centrifugazione molto più alta (~ 16.000 g) che applicato il protocollo di cui sopra. Più impegnativo è il fatto che l&#39;orientamento degli embrioni è variabile, e si può osservare la separazione a vari angoli rispetto all&#39;asse principale embrione. Questa variabilità richiede lo sperimentatore di risolvere dopo la centrifugazione come un embrione particolare è stato organizzato nella provetta da centrifuga. Con l&#39;uso del tuorlo autofluorescenza come marker per la maggior parte alta densità dell&#39;embrione, questo è spesso possibile, anche se richiede molta pratica e giudizio di farlo in modo affidabile. Se ci sono embrioni sufficiente nel campione, si può essere in grado di risolvere quei casi in cui gli embrioni capitato di essere disposti come in fig. 1. Centrifugazione degli embrioni nel loro guscio dell&#39;uovo E &#39;possibile incorporare gli embrioni in agar per il passaggio 4 senza rimuovere il corion primi 2. In questo approccio, gli embrioni sulla piastra di raccolta sono coperti con olio alocarburi invece del 50% di candeggina al punto 3.2 (alocarburi trasforma il traslucido corion). Embrioni di stadi appropriati vengono selezionati e trasferito in una nuova piastra di agar con pinzette; afferrando solo i filamenti d&#39;uovo con le pinze, danniil corretto embrione è evitato. Gli embrioni sono incorporati come nel passo 4,4 per 4,8, con i filamenti d&#39;uovo anteriore attaccare fuori dal buco nel agar. Il guscio d&#39;uovo viene rimosso da un trattamento di candeggina al 50% dopo che gli embrioni sono stati centrifugati e scavato nella agar come al punto 5.4. Centrifugazione, alla presenza del guscio dell&#39;uovo può migliorare il tasso di devitellinization dopo fissazione (passo 6.2), ma la selezione degli stadi corretti per centrifugazione (passo 4.2) è più impegnativo. Applicazioni Centrifugazione embrione è particolarmente utile per gli esperimenti di colocalizzazione, cioè, per verificare se una particolare proteina localizza a un certo organello importante. Per esempio, la proteina Klar è presente sui primi gocce lipidiche embrionale, ma questa localizzazione è difficile da dimostrare in embrioni intatto. In parte, ciò è dovuto sia puncta Klar e gocce lipidiche sono abbondanti nella periferia embrione 10, rendendo colocalizzazione spurie difficili da escludere. Tuttavia, centrifugazione concentra gocce lipidiche in uno strato distinto, e, quindi, colocalizzazione con segnale Klar diventa evidente 10. Un&#39;analisi simile è dimostrato in modo inequivocabile la localizzazione di altre proteine ​​di gocce lipidiche 8, 13-15, con esempi sorprendente come alcuni istoni 2 e nei casi in cui una proteina è presente ubiquitariamente, ma arricchito di gocce lipidiche 8. Come gli embrioni possono essere selezionate visivamente da stadio di sviluppo (punto 4.2), è anche possibile rilevare l&#39;assunzione di proteine ​​evolutivamente regolata a gocce lipidiche 8, 15. Test per colocalizzazione con gocce lipidiche è particolarmente semplice, visto che si accumulano gocce lipidiche in una visivamente distinte &quot;cap&quot;, in modo che nessun ulteriore marker o macchia è necessario per identificare questo strato. Infatti, a causa della sua facilità e risultati visivi potenzialmente sorprendenti (Fig. 1, Fig. 5. D, E), il nostro laboratorio utilizza questa tecnica ormai di routine come un primo test se una proteina è localizzata per il candidato gocce lipidiche. In linea di principio, gli studi di colocalizzazione simile dovrebbe lavorare per il organelli altri mostrato in fig. 1, e probabilmente per gli altri (compresi i parassiti intracellulari), la cui distribuzione negli embrioni centrifugati deve ancora essere documentati. Dal centrifugazione concentra organelli in bande strette, facilita l&#39;isolamento di una frazione arricchita in questi organelli. Questo approccio è già stato utilizzato per il recupero gocce lipidiche per il trapianto in embrioni di donatori 2. Può anche essere possibile analizzare biochimicamente la frazione arricchito (ad esempio, tagliando strati off embrioni dopo la fissazione 15). Il metodo descritto qui ha applicazioni che vanno oltre gli embrioni di Drosophila, la centrifugazione può essere utilizzata per stratificare le uova di diverse specie (tra cui le rane, nematodi, tunicati, anellidi, ricci di mare, molluschi e Cnidari) 16. Nel nostro laboratorio, abbiamo usato il protocollo di cui sopra per gli embrioni di Musca domestica mosca 2 e hanno impiegato centrifugazione unoriented per l&#39;analisi di embrioni dal moscerino fungo Sciara coprophila 17 e la lumaca Ilyanassa obsoleta 17. Infine, questo metodo non è limitato agli embrioni: centrifugazione può anche essere utile per stratificare altre cellule di grandi dimensioni, come gli ovociti (ad esempio, in Drosophila 2, 18 e l&#39;euforbia bug Oncopeltus fasciatus 17). Per diversi esemplari, le condizioni di centrifugazione esatti devono essere ottimizzati: per esempio, 10 minuti a 3000 g è sufficiente per stratificare efficacemente embrioni di Musca domestica 2, e se centrifugati molto più lungo, gli embrioni tendono a rompersi durante la fissazione e immunocolorazione.

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		<th>Catalogue Number</th>
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<td>60x15 mm Petri dishes</td>
<td>&nbsp;</td>
<td>Becton Dickinson</td>
<td>#35-1007</td>
<td>From Falcon</td>
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<tr>
<td>Wire mesh</td>
<td>&nbsp;</td>
<td>Small Parts</td>
<td>CX-0150</td>
<td>stainless steel type 304; mesh size #150 x150, wire diameter: 0.0026 inches</td>
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<td>Tungsten needles</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>10130-10</td>
<td>0.25 mm</td>
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<td>Moria Nickel Plated Pin/Needle Holder</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
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<td>Fly Cages</td>
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<td>Genesee Scientific</td>
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in-vivo centrifugation of drosophila embryos

Una strategia importante per purificare e isolare i diversi tipi di organelli intracellulari è quello di separare gli uni dagli altri sulla base di differenze di densità di galleggiamento. Tuttavia, quando le cellule sono interrotte prima della centrifugazione, proteine ​​e organelli in questo non-native ambiente spesso impropriamente unirsi tra di loro. Qui si descrive un metodo per separare gli organelli dalla densità in intatto, embrioni vivi Drosophila. Embrioni precoci prima cellularization vengono raccolte dalle gabbie della popolazione, e la loro gusci d&#39;uovo esterni vengono rimossi dal trattamento con il 50% di candeggina. Gli embrioni vengono poi trasferiti ad una piastra di agar piccole e inserito, estremità posteriore prima in piccoli fori verticali in agar. Le piastre contenenti embrioni incorporati vengono centrifugati per 30 minuti a 3000g. L&#39;agar supporta gli embrioni e li mantiene in un orientamento definito. In seguito, gli embrioni sono scavate nella agar con un ago smussato. Centrifugazione separa organuli principali in strati distinti, una stratificazione facilmente visibile da campo chiaro microscopia. Un certo numero di marcatori fluorescenti sono disponibili per confermare stratificazione successo in embrioni vivi. Proteine ​​associate con alcuni organelli sarà arricchita da uno strato particolare, dimostrando colocalizzazione. Singoli strati possono essere recuperati per l&#39;analisi biochimiche o il trapianto in uova donatore. Questa tecnica è applicabile per la separazione degli organelli in altre cellule di grandi dimensioni, comprese le uova e ovociti di specie diverse.

Watch this Scientific Journal Video about In-vivo Centrifugation of Drosophila Embryos at JoVE.com

In-vivo Centrifugation of Drosophila Embryos

Descriviamo un metodo per separare gli organelli dalla densità del vivere<em&gt; Drosophila</em&gt; Embrioni. Gli embrioni sono inseriti in agar e centrifugato. Questa tecnica produce separazione riproducibile organuli principali lungo l&#39;asse antero-posteriore dell&#39;embrione. Questo metodo facilita esperimenti colocalizzazione e rendimenti frazioni organello per analisi biochimiche e di esperimenti di trapianto.

In vivo centrifugazione di embrioni di Drosophila

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

in-vivo-centrifugation-of-drosophila-embryos

Research

JoVE Journal

Biology

In-vivo Centrifugation of Drosophila Embryos

18.3K Views.  University of Rochester. Descriviamo un metodo per separare gli organelli dalla densità del vivere Drosophila Embrioni. Gli embrioni sono inseriti in agar e centrifugato. Questa tecnica produce separazione riproducibile organuli principali lungo l'asse antero-posteriore dell'embrione. Questo metodo facilita esperimenti colocalizzazione e rendimenti frazioni organello per analisi biochimiche e di esperimenti di trapianto.

Video: In-vivo Centrifugation of Drosophila Embryos

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

Preparazione delle Culture neuronale da Midgastrula fase embrioni di Drosophila

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

Ricerca

Biologia

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

La preparazione di Drosophila Gli embrioni per Live-Imaging Utilizzando il protocollo goccia

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

This technique exposes the Drosophila embryonic neuromusculature for immunohistochemistry or electrophysiological recording. It is useful for studying early events in neuromuscular development or performing electrophysiology in mutants that cannot hatch.

Raccolta e Preparazione Drosophila Gli embrioni per la registrazione elettrofisiologici e altre procedure

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Mesoscopici fluorescenza per Tomografia In-vivo Imaging di sviluppo Drosophila</em

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

Preparazione di embrioni per la microscopia elettronica del Drosophila Tubo cuore embrionale

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Imaging verticale di Drosophila Embrioni

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Primary Cell Cultures from Drosophila Gastrula Embryos

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos. In brief, a large amount of staged embryos from young and healthy flies are collected, sterilized, and then physically dissociated into a single cell suspension using a glass homogenizer. After being plated on culture plates or chamber slides at an appropriate density in culture medium, these cells can further differentiate into several morphologically-distinct cell types, which can be identified by their specific cell markers. Furthermore, we present conditions for treating these cells with double stranded (ds) RNAs to elicit gene knockdown. Efficient RNAi in Drosophila primary cells is accomplished by simply bathing the cells in dsRNA-containing culture medium. The ability to carry out effective RNAi perturbation, together with other molecular, biochemical, cell imaging analyses, will allow a variety of questions to be answered in Drosophila primary cells, especially those related to differentiated muscle and neuronal cells.

Primary Cell Cultures from Drosophila Gastrula Embryos

We provide a detailed protocol for preparing primary cells dissociated from Drosophila embryos. The ability to carry out the effective RNAi perturbation, together with other molecular, biochemical and cell imaging methods will allow a variety of questions to be addressed in Drosophila primary cells.

Colture cellulari primarie da Drosophila Gastrula embrioni

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos.   In brief, a large amount of staged ...

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 ...

RNAi Interference by dsRNA Injection into Drosophila Embryos

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements and tools for genetic manipulation have become available in Drosophila 2. Soon after the initial discovery by Frie and Mello 3 that double stranded RNA can be used to knockdown the activity of individual genes in Caenorhabditis elegans, RNA interference (RNAi) was shown to provide a powerful reverse genetic approach to analyze gene functions in Drosophila organ development 4, 5.
Many organs, including lung, kidney, liver, and vascular system, are composed of branched tubular networks that transport vital fluids or gases 6, 7. The analysis of Drosophila tracheal formation provides an excellent model system to study the morphogenesis of other tubular organs 8. The Berkeley Drosophila genome project has revealed hundreds of genes that are expressed in the tracheal system. To study the molecular and cellular mechanism of tube formation, the challenge is to understand the roles of these genes in tracheal development. Here, we described a detailed method of dsRNA injection into Drosophila embryo to knockdown individual gene expression. We successfully knocked down endogenous dysfusion(dys) gene expression by dsRNA injection. Dys is a bHLH-PAS protein expressed in tracheal fusion cells, and it is required for tracheal branch fusion 9, 10. dys-RNAi completely eliminated dys expression and resulted in tracheal fusion defect. This relatively simple method provides a tool to identify genes requried for tissure and organ development in Drosophila.

RNAi Interference by dsRNA Injection into Drosophila Embryos

RNA interference has been proven very effective to analyze gene function in Drosophila tracheal development. A detailed protocol used by Jiang lab to inject dsRNA into fly embryos to knockdown gene expression is illustrated. This technique has the potential for screening genes required for tissue and organ development in Drosophila.

L&#39;interferenza di RNAi per iniezione in dsRNA Drosophila Embrioni

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

Imaging Forma Modifica cella in Living Drosophila Embrioni

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...