KB è supportato dal NIH concedere GM54544.

Parte 1: Attrezzature e Forniture <ol><li> Un microscopio a dissezione buona è necessaria per dissezioni embrione; ingrandimento 40X è suggerito, con 25X oculari per aumentare al massimo ingrandimento. </li><li> Una pinza sottile (numero 5) sono necessari per la selezione manuale di embrioni e devitellinization degli embrioni. </li><li> Attrezzature per creare e modificare gli aghi buon bicchiere è richiesto. Tirato in vetro aghi sono necessari per la dissezione. Noi preferiamo vetro solido per la dissezione (durano più a lungo), ma di serie pareti spesse tubo di vetro (diametro esterno 1-1,5 mm) funziona pure. Alcune persone usano aghi tungsteno fine, elettrolizzata per la nitidezza desiderata in un bagno di NaOH 1M con un + 10 Amp batteria autotrasformatore. Aghi affilati dissezione tungsteno hanno il vantaggio di essere relativamente resistenti e di lunga durata. Aghi di vetro cavo (formato simile a elettrodi di patch clamp) sono collegati a tubi di plastica e utilizzato durante la dissezione per l&#39;aspirazione ed espulsione di soluzione salina, così come la consegna controllata di colla. Una varietà di estrattori di vetro sono disponibili per la produzione di aghi di vetro. Computer programmato estrattori può essere regolato per tirare una varietà di forme e dimensioni. Il Brown-Flaming modelli (Sutter Instrument Co.) sono ampiamente favorite. Un microscopio composto (20-40X obiettivo) dovrebbero essere disponibili per controllare e modificare gli elettrodi prima dell&#39;uso. </li><li> Due tipi di soluzioni sono comunemente usati per il bagno durante la dissezione dei tessuti embrionali: 1) &quot;standard&quot; (Jan e Jan, 1976a) o &quot;standard modificato&quot; (Broadie, 2000) saline, basato su soluzioni comunemente usati per la registrazione in altri sistemi di invertebrati , e 2) &quot;emolinfa-like&quot; (HL) saline (Stewart et al 1994), un compromesso tra la soluzione salina standard e le concentrazioni ioniche misurate nella emolinfa Drosophila. Va notato che nessuna di queste saline imitano con precisione la composizione chimica di emolinfa in vivo (Broadie, 2000). Standard di soluzione salina contiene (in mm): 135 NaCl, 5 KCl, 4 MgCl 2, 1.8 CaCl 2, 72 saccarosio, 5 TES, pH 7,2. Sperimentatori possono anche prendere in considerazione salina insetti preparati commercialmente (ad esempio Schneider Media insetti), che non è adatto per la registrazione elettrofisiologica ma molto probabilmente per proteggere la salute dei tessuti esposti. </li></ol> Parte 2: Staging Embryo e dissezione <ol><li> Per la raccolta delle uova ottimale, mantenere giovani (&lt;7 giorni), maschio sano e mosche di sesso femminile (20-40) su cibi freschi vasi che per 2-3 giorni. Pentole posa comunemente costituiti da 100 bicchieri di plastica ml (Tri-Versare) perforata per fornire un&#39;adeguata circolazione dell&#39;aria, che copre una piastra di agar 60 mm. Piastre di agar contenenti mele o succo d&#39;uva indurito con agar. Esistono diverse ricette, ma un esempio è la seguente: 700 ml di acqua, 25-30 grammi di agar, 300 ml di succo concentrato (uva o di mela), 0,5 g metilparaben (p-hydroxymethylbenzoate) e 30g di zucchero. Autoclave l&#39;acqua e agar e far bollire a parte il p-hydroxymethylbenzoate con lo zucchero e il succo. Mescolare l&#39;agar e soluzioni succo di frutta e versare rapidamente in 60 piastre mm, rimuove le bolle. </li><li> Prima della raccolta delle uova, le mosche sono alimentati con pasta di lievito (7 grammi di lievito di birra in 9 ml di acqua conservato a 4 ° C) su piatti freschi, cambiata almeno due volte al giorno per almeno due giorni. Di giorno 3, un piatto buono dovrebbe produrre 100-200 uova all&#39;ora. Deposizione delle uova migliore si osserva in una pentola mantenuto su un lato con l&#39;agar nella piastra graffiato. Molti embrioni saranno stabilite nel o accanto ai graffi. </li><li> Per raccogliere un gran numero di embrioni, delicatamente trasferire gli embrioni dalla piastra di agar in un cesto con un pennello. Un cesto può essere fatto con un tubo da 15 o 50 ml per centrifuga. Tagliare la parte inferiore lasciando spazio sufficiente superficie per intrappolare uno schermo (70 mesh micron) tra il coperchio e la parte superiore del tubo. Sciacquare gli embrioni con dH 2 0 e messo in una piastra di Petri di fresco 50% al 100% di candeggina per rimuovere il corion esterno. In alternativa, le uova possono essere raccolti individualmente utilizzando una pinza sottile, e dechorionated mettendoli manualmente in una goccia di candeggina. De-chorionation può richiedere da 30 secondi a 2 minuti (candeggina fresca è molto più veloce), e devono essere monitorati sotto il microscopio di dissezione. La rimozione del corion espone la lucida, trasparente membrana vitellina. Dechorionated uova devono essere brevemente sciacquati, o posto in soluzione fisiologica, come candeggina distrugge rapidamente i tessuti devitellinized. </li><li> E &#39;fondamentale mettere in scena con attenzione embrioni per l&#39;età evolutiva desiderato. La Drosophila definito stadio embrionale (1-17; Campos-Ortega e Hartenstein, 1985) non sono utili, come tutto lo sviluppo funzionale neuromuscolare avviene in fase avanzata o 16 stadio 17. Così, gli embrioni sono messo in scena da ora dello sviluppo a 25 ° C in un incubatore umidificato, in ore dopo la fecondazione o, più comunemente, dopo la deposizione delle uova (AEL). In queste condizioni, l&#39;embriogenesi dura 21 + / - 1 ora a 25 ° C. </li><li> Uova da un tempo di uovo laici (1-2 hrs) che sono opportunamente invecchiato sono visti per la prossima criteri morfologici. Dopo il lavaggio in ampia dH 2 O, le uova dechorionated sono posti in un piatto di plastica cultura (sotto dH 2 O) per la visualizzazione. Corretta stadiazione può essere confermata con criteri morfologici usando la luce riflessa con un microscopio a dissezione (Campos-Ortega e Hartenstein, 1985). Maturo fase avanzata embrioni (22-24h AEL) sul punto di cova sono generalmente riconosciuti dalla segmentazione della cuticola e trachea gonfiati. Gli embrioni possono anche essere sottoposti a tipizzazione mediante GFP e bilanciatori di dissezione di un microscopio a fluorescenza con filtri appropriati. </li><li> Per la dissezione, embrioni dechorionated ed evolutivamente messo in scena sono fissati (lato dorsale verso l&#39;alto) su un vetrino con una goccia di soluzione salina (vedi punto 1.5). Gli embrioni vengono rimosse dalla membrana vitellina da pungere la membrana verso la fine anteriore o posteriore, con una micropipetta di vetro o un ago di tungsteno, e poi delicatamente liberando l&#39;embrione dalla membrana. In preparazione per la dissezione, l&#39;embrione deve essere posizionato sul vetrino con la superficie dorsale up (lato dorsale fino dissezione esporrà il sistema nervoso ventrale e neuromusculature per gli esperimenti. Tuttavia, l&#39;embrione può ovviamente essere posizionati in altri modi per facilitare l&#39;accesso ad altri tessuti.) </li><li> Embrioni stadio precoce (&lt;16 ore AEL) attaccano direttamente alla pulizia vetro o vetro rivestiti con polilisina (poli-L-lisina bromidrato; Broadie e Bate, 1993a, b). Vecchi embrioni (&gt; 16 ore AEL), a seguito di formazione cuticola, deve essere incollato, idealmente Sylgard rivestite (Dow Corning) coprioggetto. Un&#39;acqua-polimerizzazione colla cianoacrilato chirurgico o veterinario viene utilizzata (Broadie e Bate, 1993c, d). La colla è espresso attraverso una pipetta piccolo elettrodo di vetro (10-20 micrometri punta diametro interno) collegato ad un tubo di gomma con la colla flusso controllato dalla pressione bocca. Si deve prestare attenzione durante il parto colla, come la colla polimerizza rapidamente dopo il contatto con soluzione salina. Si noti inoltre che l&#39;incollaggio non può essere eseguita in soluzione salina con nessuna o scarsa concentrazione di ioni bivalenti, come il collante necessario per polimerizzare gli ioni bivalenti. Piccole quantità di colla prima utilizzazione per fissare saldamente la testa e la coda. </li><li> Una volta che le estremità dell&#39;embrione sono incollati al vetrino, un&#39;incisione è fatta lungo la linea mediana dorsale con un elettrodo di vetro o un ago di tungsteno affilata. Ciò è reso più facile da dolci perforando la linea di incisione prima di effettuare il taglio finale. I lati dell&#39;incisione vengono poi attaccato al vetrino con più colla, per diffondere delicatamente il piatto embrione. Gli organi interni tra cui l&#39;intestino, i corpi grassi e, facoltativamente, trachea vengono eliminati tramite aspirazione con una pipetta di vetro collegato al tubo di gomma (Broadie e Bate 1993a-c). L&#39;embrione dovrebbe essere allegato al piano coprioggetti, epidermide giù con la ventrale sistema nervoso centrale, nervi periferici, e somatiche muscolatura esposta per la sperimentazione. </li></ol>

Precisa stadiazione di embrioni è critica a causa della rapida maturazione delle proprietà funzionali nel corso del tempo solo diverse ore. Diversi problemi complicare questa messa in scena. In primo luogo, molti ricercatori utilizzano uovo tempo stabilisce per gli embrioni palco, ma il tempo di deposizione delle uova può variare enormemente da animale ad animale in condizioni diverse (Broadie et al. 1992). In particolare, le femmine con una dieta limitata tendono a conservare le uova fecondate per periodi prolungati...

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Embryo Collection Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific (www.flystuff.com)</td>
<td>59-100 (for 60 mm dish; other sizes available)</td>
<td>Cages can also be home made using punctured tri-pour beakers, as shown in video</td>
</tr>
<tr>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>Available from various companies</td>
<td>Surgical glue adheres better to sylgard-coated coverslips</td>
</tr>
<tr>
<td>Fine glass tubing, outer diameter 1.0-1.5 mm</td>
<td>&nbsp;</td>
<td>various</td>
<td>&nbsp;</td>
<td>For pulling into fine glass needles for dissection and tubes for glue delivery</td>
</tr>
<tr>
<td>Plastic tubing, to attach to glass pipette for mouth suction and glue application</td>
<td>&nbsp;</td>
<td>Tygon</td>
<td>&nbsp;</td>
<td>Tubing inner diameter needs to match glass outer diameter.</td>
</tr>		</tbody>
		</table>
		</div>

harvesting and preparing drosophila embryos for electrophysiological recording and other procedures

Drosophila è un modello premier genetico per lo studio sia dello sviluppo embrionale e funzionale delle neuroscienze. Tradizionalmente, questi campi sono abbastanza isolati gli uni dagli altri, con storie in gran parte indipendenti e le comunità scientifiche. Tuttavia, l&#39;interfaccia tra questi campi di solito disparati sono i programmi di sviluppo funzionali alla base dell&#39;acquisizione di proprietà elettriche di segnalazione e la differenziazione delle sinapsi chimici funzionali durante le fasi finali della formazione circuito neurale. Questa interfaccia è un&#39;area estremamente importante per le indagini. In Drosophila, queste fasi di sviluppo funzionale si verificano durante un periodo di &lt;8 ore (a 25 ° C) durante l&#39;ultimo terzo di embriogenesi. Questo periodo tardo di sviluppo è stato a lungo considerato intrattabile a causa indagine per la deposizione di un duro, cuticola impermeabile epidermica. Un passo avanti è stato anticipato l&#39;applicazione di acqua-polimerizzazione colla chirurgica che può essere applicata a livello locale per la cuticola per permettere la dissezione controllato di fase avanzata di embrioni. Con una incisione longitudinale dorsale, l&#39;embrione può essere posato piatto, esponendo il cordone ventrale dei nervi e muscolatura della parete del corpo di indagine sperimentale. Questo sistema è stato molto utilizzato per isolare e caratterizzare mutanti genetici che impedire la formazione embrionale sinapsi, e quindi svelare i meccanismi molecolari che regolano la specificazione e differenziazione delle connessioni di sinapsi e funzionali, le proprietà di segnalazione sinaptica.

Watch this Scientific Journal Video about Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures at JoVE.com

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Questa tecnica espone il neuromusculature Drosophila embrionali per immunoistochimica o registrazione elettrofisiologica. E &#39;utile per studiare eventi precoci nello sviluppo neuromuscolare o l&#39;esecuzione di elettrofisiologia in mutanti che non possono schiudersi.

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 ...

harvesting-preparing-drosophila-embryos-for-electrophysiological

Research

JoVE Journal

Biology

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

13.3K Views.  University of Illinois. Questa tecnica espone il neuromusculature Drosophila embrionali per immunoistochimica o registrazione elettrofisiologica. E 'utile per studiare eventi precoci nello sviluppo neuromuscolare o l'esecuzione di elettrofisiologia in mutanti che non possono schiudersi.

Video: Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Harvesting Sperm and Artificial Insemination of Mice

Rodents of the genus Peromyscus (deer mice) are the most prevalent native North American mammals. Peromyscus species are used in a wide range of research including toxicology, epidemiology, ecology, behavioral, and genetic studies. Here they provide a useful model for demonstrations of artificial insemination. 

Methods similar to those displayed here have previously been used in several deer mouse studies, yet no detailed protocol has been published. Here we demonstrate the basic method of artificial insemination. This method entails extracting the testes from the rodent, then isolating the sperm from the epididymis and vas deferens. The mature sperm, now in a milk mixture, are placed in the female&#x2019;s reproductive tract at the time of ovulation. Fertilization is counted as day 0 for timing of embryo development. Embryos can then be retrieved at the desired time-point and manipulated.

Artificial insemination can be used in a variety of rodent species where exact embryo timing is crucial or hard to obtain. This technique is vital for species or strains (including most Peromyscus) which may not mate immediately and/or where mating is hard to assess. In addition, artificial insemination provides exact timing for embryo development either in mapping developmental progress and/or transgenic work. Reduced numbers of animals can be used since fertilization is guaranteed. This method has been vital to furthering the Peromyscus system, and will hopefully benefit others as well.

This protocol demonstrates methods for extracting sperm from the testes of males and then inseminating female mice. This procedure is useful when precise time is needed in developmental studies as well as transgenic work.

Lo sperma raccolto e l&#39;inseminazione artificiale di topi

Rodents of the genus Peromyscus (deer mice) are the most prevalent native North American mammals.  Peromyscus species are used in a wide range of research ...

Ricerca

Biologia

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 ...

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

Metodi elettrofisiologici per la registrazione Potenziali Synaptic dal NMJ Larve di Drosophila

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Electrophysiological Recording in the Drosophila Embryo

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. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. 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.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Registrazione elettrofisiologiche in Drosophila Embryo

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 ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

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 ...

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 ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

Transfection strategia ottimizzata per espressione e registrazione elettrofisiologica di ricombinante tensione canali ionici in cellule HEK-293T

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Isolamento e purificazione di chinesina dal Drosophila Embrioni

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Preparazione individuale Drosophila Egg Chambers per l&#39;imaging dal vivo

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

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

Riepilogo

Esplora Altri Video

Capitoli in questo video

Lo sperma raccolto e l'inseminazione artificiale di topi