This work was supported by NIH grant NS27501 to DKOD. Additional support for this work from an HHMI Professor Grant to DKOD.

I. Drosophila Embryo Collection
<ol>
<li>Prepare egg collection plates <ol>
<li>Boil 200 ml dH2O with 8 g agar</li>
<li>Cool to 50&deg;C</li>
<li>Add 2 ml EtOH and 2 ml Acetic acid</li>
<li>Pour into 35 mm plastic petri dishes, tops </li>
<li>Cool and store in air tight container at 4&deg;C</li>
<li>Makes approximately 80 plates </li>
</ol></li>
<li>Prepare Yeast paste <ol>
<li>Dissolve Fleishmans active baking yeast in water to form paste</li>
<li>Microwave for 20-30 seconds to kill yeast (watch out for boiling over)</li>
<li>Store in 10 cc syringe (without needle) at 4&deg;C for 2 weeks</li>
</ol></li>
<li>Flies:&nbsp;The adults used for egg collection are generally most productive between 3 and 14 days after emerging, providing that they have had access to fresh food. Many mutant stocks have decreased fertility that can manifest itself in a lowered rate of egg laying and a change in the age at which they are most productive. Generally, several hundred adults make for a good egg-laying population.</li>
<li>Procedure: Spread a thin layer of yeast paste on an egg collection plate. This provides a favorable substrate for the laying of eggs. Transfer adult flies to a clean egg collection bottle and tape the collection plate to the mouth of the bottle. Don't leave the flies in these bottles longer than 3-4 hours as the humidity rises and the flies get stuck in paste and on sides of bottle.</li>
</ol>
II. Embryo Dechorionation with Bleach
<ol>
<li>Set up a Buchner funnel connected to a filter flask attached to an aspirator. Put a piece of Whatman #1 paper in the funnel. With suction running, wet the paper with sterile distilled water.</li>
<li>Rinse embryos from collection plates onto the filter paper with a stream of water from a wash bottle.</li>
<li>Rinse them in several volumes of sterile water, turn off aspirator, and then add a funnel volume of 50% bleach. Let eggs sit 5-10 minutes. The chorions will be dissolved in 1-2 minutes, but the longer you leave the embryos in the bleach the more of the contaminating yeast will be destroyed. Pick out any adults or larvae that have rinsed off dish.</li>
<li>Rinse the embryos into a sterile petri dish (NOT tissue culture) with sterile distilled water. Many of the embryos will stick to the bottom of the dish if it has not been pre-wetted. Rinse several times with sterile water in this dish.</li>
<li>Examine the embryos with transmitted light to pick out mid-gastrula stage embryos.</li>
</ol>
III. Preparation of single embryo cultures
<ol>
<li>Make DDM1 (see recipes in section IV)</li>
<li>Pour water off dish of embryos just prior to culturing. Cover embryos with sterile media.</li>
<li>Set out 3, 35 mm petri dishes. Put in 4 autoclaved round coverslips in each dish. On each coverslip, put 5 &micro;l of medium. Use Bellco coverslips-low lead glass coverlips. Bellco Biological Glassware 800-257-7043 Cat #1943-00012 </li>
<li>Pull some fairly fine pipets. The pipets we use are pulled on a Narishige PP-83 puller. The exact dimensions of the pipet are not crucial and we usually pull them finer than we want and them break off the tip in the same field of view as the embryo to gauge the size. You don't want to suck up all of the embryos in one go, and your don't want the tip so small that it takes 5 minutes to suck up the cells. Aim for somewhere in between. Examine the embryos with transmitted light and use a mouth suction tube attached to the pipet to remove the contents of the embryo. Disperse the cells onto the prepared coverslip by gently expelling the cells as close to the coverslip as possible. You may want to examine the coverslips at higher power and further disperse the cells in the same manner.</li>
<li>Let the cells sit for no longer than 10 minutes. Flood the dish with media and put in incubator, 4-5% CO2 environment, if using defined medium. Temperature between 22-24&deg;C. We routinely use 23&deg;C and 95% humidity. Humidity is important, as you want to avoid evaporation. You can use a standard mammalian incubator placed in a coldroom with temperature control set to 23&deg;C.</li>
</ol>
IV. DDM1 Defined medium for growing the cultures
<ol>
<li>Make DMEM: To 100 mls of Ham's F12/DME Media (DMEM) (Irvine Scientific #9052). <ol>
<li>Add 0.476 g Hepes (20mM) (Sigma #H-3375). Mix.</li>
<li>Add 1.25 ml L-Glutamine 200 mM (Irvine Scientific #9317). Mix.</li>
<li>Filter in hood with 0.2 &micro;m acetate syringe filter (need 2 filters). </li>
<li>The pH is 6.64 and Osm 288.</li>
<li>Make aliquots of 10ml in 15ml centrifuge tubes.</li>
<li>Store at 4&deg;C for not more than 2 weeks. Note: Always use autoclaved filtered water and make in containers that are reserved for Media and Supplements only. Never put a pH electrode or stir bar used for other purposes in Media.To make DDM1: Add supplements listed below to DMEM just prior to culturing. </li>
</ol></li>
<li>To 10 mls of DMEM add:
<ul>
<li>100 &micro;l Transferrin</li>
<li>100 &micro;l Putrescine</li>
<li>100 &micro;l Selenium</li>
<li>100 &micro;l Progesterone</li>
<li>50 &micro;l 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 cultures prepared from midgastrula stage embryos the neurons arise from neuroblast precursors, many of which divide prior to differentiation in vitro. This system thus provides a unique opportunity for exploration of genetic and environmental factors important in very early phases of neuronal development (Rohrbaugh et al., 2003). We have shown that neurons grown in a simple defined medium differentiate into electrically excitable neurons that also form functional synaptic connections (O Dowd, 1995; Lee and ...

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<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>Drosophila melanogaster</td>
<td>Animal</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>Fruit flies</td>
</tr>
<tr>
<td>Transferrin</td>
<td>Reagent</td>
<td>Sigma</td>
<td>T-1147</td>
<td>100x Stock: 10 mg/ml in water. Filter through 0.2 um syringe filter (cellulose acetate). Store in 220 ul aliquots (for 20 ml DMEM) at -20C.
</td>
</tr>
<tr>
<td>Insulin</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>I-6634</td>
<td>200x Stock: 10 mg/ml in 0.05N HCl. Filter through 0.2 um syringe filter (cellulose acetate). Store in 120 ul aliquots (for 20 ml DMEM) at -20C.
</td>
</tr>
<tr>
<td>Putrescine</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P-5780</td>
<td>100x Stock: 10 mM in ddH2O. Filter through 0.2 um syringe filter (cellulose acetate). Store in 220 ul aliquots (for 20 ml DMEM) at -20 C
</td>
</tr>
<tr>
<td>Selenium</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>S-5261</td>
<td>100x Stock: 3 uM in ddH2O. Put 0.0051 g Selenium in a 15 ml tube labeled A (3 mM stock). Add 10 ml of sterile water. Take 10 ul from Tube A to Tube B with 10 ml ddH2O (3 uM stock). Filter Tube B through 0.2 um syringe filter (cellulose acetate). Store in 220 ul aliquots (for 20 ml DMEM) at -20C
</td>
</tr>
<tr>
<td>Progesterone</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P-6149</td>
<td>100x Stock: 2 ug/ml
1.	Add 1ml of 100% EtOH to 0.001 g Progesterone bottle
2.	Add 49 ml ddH2O
3.	Transfer 1 ml of this to second tube with 9 ml ddH2O
4.	Filter through 0.2 um syringe filter (cellulose acetate)
5.	Store in 220 ul aliquots (for 20 ml DMEM) at -20C
</td>
</tr>
<tr>
<td>DDM1</td>
<td>Medium</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>To 10 mls of DMEM add from stocks:
	100 ul Transferrin, 
	100 ul Putrescine,
	100 ul Selenium,
	100 ul Progesterone,
	 50 ul Insulin
</td>
</tr>
<tr>
<td>Petri dishes</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Cover-slips</td>
<td>&nbsp;</td>
<td>Bellco Biological Glassware</td>
<td>1943-00012</td>
<td>Low lead glass, autoclaved.</td>
</tr>
<tr>
<td>Paper filter</td>
<td>&nbsp;</td>
<td>Whatman</td>
<td>#1</td>
<td>&nbsp;</td>
<tr>
<td>10cc syringe</td>
<td>Tool</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>
			<div>
			The cultures made in this media must be maintained in a 5% CO2 incubator at about 22-24&deg;C. We use a standard mammalian tissue culture incubator placed in a cold room and heated to 23&deg;C. Always used autoclaved filtered water and make in containers that are reserved for media and supplements only. Never put a pH meter or stir bar used for other purposes in media.		</div>

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.

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

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.

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

preparation-neuronal-cultures-from-midgastrula-stage-drosophila

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Biology

8.8K vistas.  University of California, Irvine (UCI). Este video muestra la preparación de cultivos primarios de neuronas de midgastrula etapa embriones de Drosophila. Puntos de vista de las culturas show en vivo de las células de una hora después de la siembra y las neuronas diferenciadas después de 2 días de crecimiento en un medio de bicarbonato definido. Las neuronas son eléctricamente excitables y forman conexiones sinápticas.

Video: 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.

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

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.

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.

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.

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.

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.

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.

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.

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.

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.

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