Diese Arbeit wurde vom NIH NS27501 zu DKOD unterstützt. Zusätzliche Unterstützung für diese Arbeiten von einem HHMI Professor Grant zu DKOD.

I. Drosophila Embryo-Entnahme <ol><li> Bereiten Eiersammlung Platten <ol><li> Kochen 200 ml dH 2 O mit 8 g Agar </li><li> Abkühlen auf 50 ° C </li><li> 2 ml EtOH und 2 ml Essigsäure </li><li> Gießen Sie in 35 mm Kunststoff-Petrischalen, Tops </li><li> Cool und speichern in luftdichten Behälter bei 4 ° C </li><li> Ergibt ca. 80 Teller </li></ol></li><li> Bereiten Hefepaste <ol><li> Lösen Sie Fleishmans aktive Backhefe in Wasser zu fügen </li><li> Mikrowelle für 20-30 Sekunden, um Hefe zu töten (achten Sie auf Überkochen) </li><li> Shop in 10 ml-Spritze (ohne Kanüle) bei 4 ° C für 2 Wochen </li></ol></li><li> Fliegen: Die Erwachsenen für Eiersammlung werden in der Regel am produktivsten zwischen 3 und 14 Tage nach dem Austritt, sofern sie hatten Zugang zu frischen Lebensmitteln. Viele Mutanten Aktien haben die Fruchtbarkeit, die manifestiert sich in einer abgesenkten Satz von der Eiablage und eine Veränderung in dem Alter, in dem sie am produktivsten sind gesunken. Generell machen mehrere hundert Erwachsene für einen guten Eiablage Bevölkerung. </li><li> Vorgehensweise: Verteilen Sie eine dünne Schicht von Hefe auf einem Ei Sammlung Platte einfügen. Dies bietet ein günstiges Substrat für die Eiablage. Transfer erwachsenen Fliegen auf eine saubere Entnahme der Eizellen Flasche und Band der Sammlung Platte an der Mündung der Flasche. Lassen Sie das Fliegen in dieser Flaschen länger als 3-4 Stunden, da die Luftfeuchtigkeit steigt und die Fliegen zu bekommen in Paste und auf Seiten der Flasche stecken. </li></ol> II. Embryo Dechorionation mit Bleach <ol><li> Richten Sie einen Büchner-Trichter an eine Saugflasche an einen Aspirator. Legen Sie ein Stück Whatman Papier Nr. 1 in den Trichter. Mit Saug-laufen, nass das Papier mit sterilem destilliertem Wasser. </li><li> Spülen Embryonen aus der Sammlung Platten auf das Filterpapier mit einem Strom von Wasser aus einer Spritzflasche. </li><li> Spülen Sie sie in mehreren Bänden von sterilem Wasser, schalten Sie Aspirator, und fügen Sie dann einen Trichter von 50% Bleichmittel. Lassen Eiern sitzen 5-10 Minuten. Das Chorion wird in 1-2 Minuten aufgelöst werden, aber je länger Sie verlassen die Embryonen in der Bleiche die mehr der kontaminierenden Hefe zerstört werden. Suchen Sie sich keine Erwachsenen oder Larven, die aus Schale haben gespült. </li><li> Spülen Sie die Embryonen in eine sterile Petrischale (NICHT Gewebekultur) mit sterilem destilliertem Wasser. Viele der Embryonen wird der Boden der Schale haften, wenn es nicht bereits vorher benetzt. Spülen Sie mehrmals mit sterilem Wasser in dieses Gericht. </li><li> Untersuchen Sie die Embryonen mit Durchlicht herausgreifen Mitte Gastrulastadium Embryonen. </li></ol> III. Vorbereitung der einzelnen Embryokulturen <ol><li> Machen DDM1 (siehe Rezepte in Abschnitt IV) </li><li> Gießen Sie Wasser aus Schüssel von Embryonen kurz vor dem Züchten. Abdeckung Embryonen mit sterilen Medien. </li><li> Set aus 3, 35 mm Petrischalen. Legen Sie in 4 autoklaviert runden Deckgläschen in jedem Gericht. Auf jedem Deckglas, legte 5 ul Medium. Verwenden Sie Bellco Deckgläser-low Bleiglas coverlips. Bellco Biological Glaswaren 800-257-7043 Cat # 1943-00012 </li><li> Ziehen Sie einige ziemlich feinen Pipetten. Die Pipetten verwenden wir sind auf einem Narishige PP-83 Abziehers. Die genauen Abmessungen der Pipette sind nicht entscheidend und wir in der Regel ziehen sie feiner als wir wollen, und sie brechen Sie die Spitze in der gleichen Sichtfeld wie der Embryo, um die Größe zu messen. Sie wollen nicht zu saugen alle Embryonen in einem Rutsch, und Ihr nicht wollen, dass die Spitze so klein, dass es 5 Minuten dauert, saugen die Zellen. Ziel für irgendwo dazwischen. Untersuchen Sie die Embryonen mit Durchlicht und mit einem Mund Saugrohr an der Pipette, um den Inhalt des Embryos zu entfernen. Disperse die Zellen auf die vorbereitete Deckglas durch vorsichtiges Vertreibung der Zellen, um die Deckglas nah wie möglich. Vielleicht möchten Sie die Deckgläser mit höherer Leistung zu prüfen und weiter zu verstreuen die Zellen in der gleichen Weise. </li><li> Lassen Sie die Zellen sitzen nicht länger als 10 Minuten. Flood die Schüssel mit den Medien und setzen in Inkubator, 4-5% CO 2-Umgebung, wenn mit Hilfe von definierten Medium. Temperatur zwischen 22-24 ° C. Wir verwenden routinemäßig 23 ° C und 95% Luftfeuchtigkeit. Feuchtigkeit ist sehr wichtig, da Sie, um Verdunstung zu vermeiden möchten. Sie können eine Standard-Säugetier-Inkubator in einer Kühlzelle mit Temperaturkontrolle platziert wurde auf 23 ° C </li></ol> IV. DDM1 definiertem Medium für den Anbau der Kulturen <ol><li> Machen DMEM: Zu 100 ml Ham F12/DME Medien (DMEM) (Irvine Scientific # 9052). <ol><li> Add 0,476 g Hepes (20 mm) (Sigma # H-3375). Mix. </li><li> Add 1,25 ml L-Glutamin 200 mM (Irvine Scientific # 9317). Mix. </li><li> Filter in Kapuze mit 0,2 mu m Acetat-Spritzenfilter (need 2 Filter). </li><li> Der pH-Wert ist 6.64 und Osm 288. </li><li> Machen Aliquots von 10 ml in 15ml-Zentrifugenröhrchen. </li><li> Lagerung bei 4 ° C für nicht länger als 2 Wochen. Hinweis: Verwenden Sie immer autoklaviert gefiltertem Wasser und in Containern, die für Medien und Ergänzungen vorbehalten sind zu machen. Niemals einer pH-Elektrode oder Rührstab für andere Zwecke in Media.To verwendet machen DDM1: In Ergänzung unten aufgeführtenzu DMEM kurz vor Kultivierung. </li></ol></li><li> Zu 10 ml DMEM hinzufügen: <ul><li> 100 ul Transferrin </li><li> 100 ul Putrescin </li><li> 100 ul Selenium </li><li> 100 ul Progesteron </li><li> 50 pl Insulin </li></ul></li></ol>

<ol>
	<li>Rohrbough, J., O'Dowd, D. K., Baines, R. A., Broadie, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+bases+of+behavioral+plasticity:+Establishing+and+modifying+synaptic+circuits+in+the+Drosophila+Genetic+System.">Cellular bases of behavioral plasticity: Establishing and modifying synaptic circuits in the Drosophila Genetic System.</a> J. Neurobiol. 54, 254-271 (2003).</li><li>O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Voltage-gated+currents+and+firing+properties+of+embryonic+Drosophila+neurons+grown+in+a+chemically+defined+medium.">Voltage-gated currents and firing properties of embryonic Drosophila neurons grown in a chemically defined medium.</a> J. Neurobiol. 27, 113-126 (1995).</li><li>Lee, D., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+excitatory+synaptic+transmission+mediated+by+nAChR+in+cultured+Drosophila+neurons.">Fast excitatory synaptic transmission mediated by nAChR in cultured Drosophila neurons.</a> J. Neurosci. 19, 5311-5321 (1999).</li><li>Lee, D., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyclic-AMP-dependent+plasticity+at+excitatory+cholinergic+synapses+in+Drosophila+neurons:+Alterations+in+the+memory+mutant+dunce.">Cyclic-AMP-dependent plasticity at excitatory cholinergic synapses in Drosophila neurons: Alterations in the memory mutant dunce.</a> J. Neurosci. 20, 2104-2111 (2000).</li><li>Hodges, D. D., Lee, D., Boswell, K., Preston, C. F., Hall, L. M., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=tipE+regulates+sodium-dependent+repetitive+firing+in+Drosophila+neurons.">tipE regulates sodium-dependent repetitive firing in Drosophila neurons.</a> Mol. Cell Neurosci. 19, 402-416 (2002).</li><li>Lee, D., Su, H., Dowd, O., K, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GABA+receptors+containing+Rdl+subunits+mediate+fast+inhibitory+synaptic+transmission+in+Drosophila+neurons.">GABA receptors containing Rdl subunits mediate fast inhibitory synaptic transmission in Drosophila neurons.</a> J. Neurosci. 23, 4625-4634 (2003).</li></ol>

In Drosophila Kulturen aus midgastrula Embryonen bereitete den Neuronen entstehen aus Neuroblasten Vorläufer, von denen viele vor der Differenzierung in vitro zu teilen. Dieses System bietet somit eine einmalige Gelegenheit für die Exploration von genetischen und umweltbedingten Faktoren wichtig in sehr frühen Phasen der neuronalen Entwicklung (Rohrbaugh et al., 2003). Wir haben gezeigt, dass Neurone in einer einfachen Definition-Medium in elektrisch erregbaren Neuronen, bilden auch funktionelle synaptische Verbindungen (O Dowd, 1995; Le...

Die Kulturen in diesem Medien getroffen werden, sind eine 5% ige CO2-Inkubator bei etwa 22-24 ° C gehalten werden Wir verwenden eine Standard-Säugetier-Zellkultur-Inkubator in einem kalten Raum gebracht und erhitzt, um 23 ° C. Immer verwendet autoklaviert gefiltertem Wasser und machen in Containern, die für Medien und Ergänzungen vorbehalten sind. Niemals ein pH-Meter oder Rührstab für andere Zwecke in den Medien verwendet.

preparation of neuronal cultures from midgastrula stage drosophila embryos

Dieses Video zeigt das Verfahren zur Herstellung von primären neuronalen Kulturen aus midgastrula Bühne Drosophila-Embryos. Die Methoden für die Erhebung Embryonen und ihre dechorionation mit Bleichmittel demonstriert. Mit einer Glaspipette an einem Mund-Saugrohr, illustrieren wir die Entfernung aller Zellen, die aus einzelnen Embryonen. Das Verfahren zum Dispergieren Zellen aus jeder embyro in einen kleinen (5 l) Tropfen Medium auf einer unbeschichteten Deckglas wird demonstriert. Ein Blick durch das Mikroskop bei 1 Stunde nach dem Ausplattieren zeigt die bevorzugte Zelldichte. Die meisten der Zellen, wenn in definierten Medium überleben Neuroblasten, dass ein oder mehrere Male in der Kultur teilen, bevor Erweiterung neuritischen Prozesse durch 12-24 Stunden. Ein Blick durch das Mikroskop zeigt den Grad der Neuritenwachstum und Verzweigung in eine gesunde Kultur in 2 Tage in vitro zu erwarten. Die Kulturen werden in einer einfachen Bicarbonat Basis definiert Medium, in einer 5% CO 2-Inkubator bei 22-24 ° C. Neuritischen Prozesse weiter zu erarbeiten über die erste Woche in der Kultur, und wenn sie in Kontakt mit Neuriten aus dem benachbarten Zellen bilden sie oft funktionelle synaptische Verbindungen. Neuronen in diesen Kulturen auszudrücken spannungsabhängigen Natrium, Kalzium und Kalium-Kanälen und sind elektrisch erregbar. Diese Kultur ist nützlich für die Untersuchung molekularer genetischen und umweltbedingten Faktoren, die neuronale Differenzierung, Erregbarkeit und Synapsenbildung / Funktion zu regulieren.

Watch this Scientific Journal Video about Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos at JoVE.com

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

Dieses Video zeigt die Herstellung von primären neuronalen Kulturen aus midgastrula Bühne Drosophila-Embryos. Ansichten von lebenden Kulturen zeigen Zellen 1 Stunde nach dem Ausplattieren und differenzierten Neuronen nach 2 Tagen Wachstum in einem Bicarbonat-basierten definiertem Medium. Die Neuronen sind elektrisch erregbar und bilden synaptische Verbindungen.

Vorbereitung der neuronalen Kulturen aus Midgastrula Bühne Drosophila Embryonen

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

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JoVE Journal

Biology

8.7K Aufrufe.  University of California, Irvine (UCI). Dieses Video zeigt die Herstellung von primären neuronalen Kulturen aus midgastrula Bühne Drosophila-Embryos. Ansichten von lebenden Kulturen zeigen Zellen 1 Stunde nach dem Ausplattieren und differenzierten Neuronen nach 2 Tagen Wachstum in einem Bicarbonat-basierten definiertem Medium. Die Neuronen sind elektrisch erregbar und bilden synaptische Verbindungen.

Serie: Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

Primäre neuronale Kulturen aus den Gehirnen von Late Stage Drosophila Puppen

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

Forschung

Biologie

Placing Growth Factor-Coated Beads on Early Stage Chicken Embryos

The neural tube expresses many proteins in specific spatiotemporal patterns during development. These proteins have been shown to be critical for cell fate determination, cell migration, and formation of neural circuits. Neuronal induction and patterning involve bone morphogenetic protein (BMP), sonic hedgehog (SHH), fibroblast growth factor (FGF), among others. In particular, the expression pattern of Fgf8 is in close proximity to regions expressing BMP4 and SHH. This expression pattern is consistent with developmental interactions that facilitate patterning in the telencephalon.

Here we provide a visual demonstration of a method in which an in ovo preparation can be used to test the effects of Fgfs in the formation of the forebrain. Beads are coated with protein and placed in the developing neural tube to provide sustained exposure. Because the procedure uses small, carefully placed beads, it is minimally invasive and allows several beads to be placed within a single neural tube. Moreover, the method allows for continued development so that embryos can be analyzed at a more mature stage to detect changes in anatomy and in neural patterning. This simple but useful protocol allows for real time imaging. It provides a means to make spatially and temporally limited changes to endogenous protein levels.

A variety of growth factors and proteins interact to induce cells to take on different cell fates during development. Here we demonstrate the use of an in ovo preparation to address possible interactions between different proteins in development by placing beads on E2.5 chick embryos.

Platzieren Growth Factor-Coated Perlen auf Early Stage Hühnerembryonen

The neural tube expresses many proteins in specific spatiotemporal patterns during development.   These proteins have been shown to be critical for cell ...

In Ovo Electroporations of HH Stage 10 Chicken Embryos

Large size and external development of the chicken embryo have long made it a valuable tool in the study of developmental biology.  With the advent of molecular biological techniques, the chick has become a useful system in which to study gene regulation and function.  By electroporating DNA or RNA constructs into the developing chicken embryo, genes can be expressed or knocked down in order to analyze in vivo gene function.  Similarly, reporter constructs can be used for fate mapping or to examine putative gene regulatory elements.  Compared to similar experiments in mouse, chick electroporation has the advantages of being quick, easy and inexpensive.  This video demonstrates first how to make a window in the eggshell to manipulate the embryo.  Next, the embryo is visualized with a dilute solution of India ink injected below the embryo.  A glass needle and pipette are used to inject DNA and Fast Green dye into the developing neural tube, then platinum electrodes are placed parallel to the embryo and short electrical pulses are administered with a pulse generator.  Finally, the egg is sealed with tape and placed back into an incubator for further development.  Additionally, the video shows proper egg storage and handling and discusses possible causes of embryo loss following electroporation.

Chick in ovo electroporation is a technique which allows genetic manipulation of the avian embryo. Common applications of this technique include functional analysis of genes and putative enhancer elements. This video demonstrates neural tube electroporation in HH 10 chick embryos. Injection technique and proper egg handling are discussed.

In Ovo Electroporations von HH Stage 10 Hühnerembryonen

Large size and external development of the chicken embryo have long made it a valuable tool in the study of developmental biology.  With the advent of ...

Neuronal Cell Cultures from Aplysia for High-Resolution Imaging of Growth Cones

Neuronal growth cones are the highly motile structures at the tip of axons that can detect guidance cues in the environment and transduce this information into directional movement towards the appropriate target cell. To fully understand how guidance information is transmitted from the cell surface to the underlying dynamic cytoskeletal networks, one needs a model system suitable for live cell imaging of protein dynamics at high temporal and spatial resolution. Typical vertebrate growth cones are too small to quantitatively analyze F-actin and microtubule dynamics. Neurons from the sea hare Aplysia californica are 5-10 times larger than vertebrate neurons, can easily be kept at room temperature and are very robust cells for micromanipulation and biophysical measurements. Their growth cones have very defined cytoplasmic regions and a well-described cytoskeletal system. The neuronal cell bodies can be microinjected with a variety of probes for studying growth cone motility and guidance. In the present protocol we demonstrate a procedure for dissection of the abdominal ganglion, culture of bag cell neurons and setting up an imaging chamber for live cell imaging of growth cones.

Aplysia californica neurons develop large growth cones in culture that are excellent for high-resolution imaging of growth cone motility and guidance. Here, we present a protocol for dissection and plating of Aplysia bag cell neurons as well as for setting up a chamber for live cell imaging.

Neuronalen Zellkulturen von Aplysia für High-Resolution Imaging of Growth Cones

Neuronal growth cones are the highly motile structures at the tip of axons that can detect guidance cues in the environment and transduce this information ...

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.

Die Herstellung Drosophila Embryonen für Live-Imaging mit dem Hanging-Drop-Protokoll

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

Preparation of Aplysia Sensory-motor Neuronal Cell Cultures

The nervous system of the marine mollusk Aplysia californica is relatively simple, consisting of approximately 20,000 neurons. The neurons are large (up to 1 mm in diameter) and identifiable, with distinct sizes, shapes, positions and pigmentations, and the cell bodies are externally exposed in five paired ganglia distributed throughout the body of the animal. These properties have allowed investigators to delineate the circuitry underlying specific behaviors in the animal1. The monosynaptic connection between sensory and motor neurons is a central component of the gill-withdrawal reflex in the animal, a simple defensive reflex in which the animal withdraws its gill in response to tactile stimulation of the siphon. This reflex undergoes forms of non-associative and associative learning, including sensitization, habituation and classical conditioning. Of particular benefit to the study of synaptic plasticity, the sensory-motor synapse can be reconstituted in culture, where well-characterized stimuli elicit forms of plasticity that have direct correlates in the behavior of the animal2,3. Specifically, application of serotonin produces a synaptic strengthening that, depending on the application protocol, lasts for minutes (short-term facilitation), hours (intermediate-term facilitation) or days (long-term facilitation). In contrast, application of the peptide transmitter FMRFamide produces a synaptic weakening or depression that, depending on the application protocol, can last from minutes to days (long-term depression). The large size of the neurons allows for repeated sharp electrode recording of synaptic strength over periods of days together with microinjection of expression vectors, siRNAs and other compounds to target specific signaling cascades and molecules and thereby identify the molecular and cell biological steps that underlie the changes in synaptic efficacy.
An additional advantage of the Aplysia culture system comes from the fact that the neurons demonstrate synapse-specificity in culture4,5. Thus, sensory neurons do not form synapses with themselves (autapses) or with other sensory neurons, nor do they form synapses with non-target identified motor neurons in culture. The varicosities, sites of synaptic contact between sensory and motor neurons, are large enough (2-7 microns in diameter) to allow synapse formation (as well as changes in synaptic morphology) with target motor neurons to be studied at the light microscopic level.
In this video, we demonstrate each step of preparing sensory-motor neuron cultures, including anesthetizing adult and juvenile Aplysia, dissecting their ganglia, protease digestion of the ganglia, removal of the connective tissue by microdissection, identification of both sensory and motor neurons and removal of each cell type by microdissection, plating of the motor neuron, addition of the sensory neuron and manipulation of the sensory neurite to form contact with the cultured motor neuron.

Preparation of Aplysia Sensory-motor Neuronal Cell Cultures

Primary cultures of Aplysia sensory-motor neurons provide a model preparation for studying synapse formation and synaptic plasticity in vitro. This video demonstrates the identification and microdissection of sensory and motor neurons from Aplysia ganglia as well as the methods for establishing and maintaining sensory-motor neurons in culture.

Vorbereitung der Aplysia Sensory-Motor neuronalen Zellkulturen

The nervous system of the marine mollusk Aplysia californica is relatively simple, consisting of approximately 20,000 neurons.  The neurons are large (up ...

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.

Vorbereitung der Embryonen für Elektronenmikroskopie der Drosophila Embryonale Herz Rohr

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-Zentrifugation von Drosophila Embryonen

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.

Upright Imaging von Drosophila Embryos

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.

Primären Zellkulturen aus Drosophila Gastrula Embryonen

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