J.B. was supported by the Damon Runyon Cancer Research Foundation
Fellowship DRG-1716-02. J.Z. was supported by NIH/NIGMS Fellowship F32 GM082174-02. N.P. is an investigator of the Howard Hughes Medical Institute.

1. Preparation of fly cages
<ol>
<li>Grow Drosophila (wild-type strains such as Oregon-R or any genotype) in convenient containers such as 32-oz insect cups.</li>
<li>Transfer newly eclosed flies to large embryo-collection cages or fly population cages.</li>
<li>Move the cages into a room with a temperature of ~25 &deg;C and humidity ~65% in a light 12-h and a dark 12-h cycle to facilitate the collection of synchronous embryos.</li>
<li>Maintain the flies in the cages by feeding them killed yeast paste streaked on molasses plates. Change the molasses plates (with killed yeast paste) once a day for 2-3 d.</li>
</ol>
2. Collection of synchronous embryos
<ol>
<li>On the day of primary-cell preparation, feed the flies wth fresh plates coated with killed yeast paste for a 1-2 h pre-lay period.</li>
<li>Just before the 12-h dark cycle, replace the pre lay plates with fresh molasses plates streaked with a minimal amount of yeast paste. Discard the pre lay collection, which contains asynchronous older embryos.</li>
<li>Collect embryos for a period of 1-2 h.</li>
<li>Remove the plates containing the relatively synchronous population of embryos and store
at 25&deg;C for 5-6 h. Embryos will be at the advanced gastrula stage at this time.</li>
</ol>
3. Collection and washing of embryos
<ol>
<li>Loosen the embryos from the plates with a wet paintbrush, and rinse thoroughly with tepid tap water through a stacked set of sieves, so that the embryos collect in the finest mesh sieve at the bottom. Rinse for a few minutes to remove as much yeast and fly debris as possible.</li>
<li>Tip the sieves so that the embryos accumulate at one side, and gently wash them off from the sieve into a dechorionation basket.</li>
<li>Move to the tissue culture hood area without letting the embryos dry out.</li>
</ol>
4. Homogenization of embryos and plating of cells
<ol>
<li>Sterilize and dechorionate the embryos by immersing them in 50% (vol/vol) bleach for 5-10 min. Rinse the embryos off the wall of the basket with 70% (vol/vol) ethanol. Alternatively, wash the bleach off embryos with distilled water before ethanol treatment. From this point onward embryos should be handled in sterile conditions and in the tissue culture hood.</li>
<li>Rinse the embryos thoroughly to remove the bleach and/or ethanol with autoclaved distilled water.</li>
<li>During the dechorionation step, fill a Dounce homogenizer with the appropriate volume of room temperature M3 medium based on the amount of embryos. The ratio between the volume of embryos and that of media should not be over 0.5-ml embryos: 40-ml media (or ~100-200 embryos/ml).</li>
<li>Place the rinsed embryos in a sterilized beaker with the amount of M3 media enough to submerge the embryos. Let the embryos soak for 2-5 min.</li>
<li>Disassemble the dechorionation basket and blot the Nitex mesh dry with sterilized paper towels.</li>
<li>Transfer the embryos into the homogenizer with a sterilized paintbrush.</li>
<li>Homogenize gently with short strokes using a loose pestle without reaching the bottom of the tube until all the embryos are suspended. Then gently, but firmly, homogenize with eight full strokes. Do not twist the pestle as it moves up and down.</li>
<li>Transfer the homogenate into a sterile 50-ml conical centrifuge tube either by pouring or pipetting, and cap the tube.</li>
<li>Centrifuge for 10 min at 40g, room temperature to pellet the tissue debris, large cell clumps and vitelline membranes. Transfer the supernatant containing the cells and yolk to a clean tube and repeat the centrifugation step for 5 min.</li>
<li>Carefully transfer the supernatant by pouring or pipetting into a clean tube and spin for 10 min at 360g, room temperature to pellet the cells. Discard the supernatant. Resuspend the cell pellet in 10 ml of culture medium (serum-free medium for RNAi experiments).</li>
<li>Count viable cells using a hemocytometer to estimate the cell density. Meanwhile, repeat step 4.10 to pellet the cells.</li>
<li>Resuspend the cells to a desired concentration. To start, try a range of 1~5 x106 cells /ml and plate out at 1.7 to 2.5 x105 cells /cm2.</li>
</ol>
5. Primary-cell RNAi for cells grown on a 384-well plate
<ol>
<li>Dispense 5 &mu;l of dsRNAs into each well at a final concentration of ~250 ng per well in a 384-well plate.</li>
<li>Use a multi-channel pipette, dispense 10 ml (1~4x106 cells/ml) of primary cells per well in a serum-free M3 medium in a 384-well plate containing a different dsRNA in each well. Alternatively, cells can be cultured on 8-well glass chamber slides for confocal imaging.</li>
<li>Centrifuge the plates for 1 min at 300g, room temperature. Culture the cells in serum-free medium at 25&deg;C for 22 h or at 18&deg;C for 1-2 d.</li>
<li>Add 30 ml of serum-containing culture medium to each well.</li>
<li>Centrifuge the plates for 1 min at 300g, room temperature.</li>
<li>Place the plates in a humid chamber and culture the primary cells for an additional 5~7 d at 25&deg;C.</li>
<li>Image the cells either live or after immunofluorescence staining.</li>
</ol>
6. Representative Results:
In optimal conditions, cells in the culture will look healthy with a round morphology. The culture will have little cell debris, and be 50~80% confluent after initial plating. These cells will undergo morphological changes after being cultured for an extended period of time. It usually takes about 5-8 days for cells in the culture to be fully differentiated at 25&deg;C. Good quality primary cultures are usually judged by how well the cells-of-interest are differentiated in culture. For example, primary muscle cells often have a branch-like morphology and become multinucleated myotubes with stripe-like myofibrils consisting of a series of sarcomeres. Fully differentiated myotubes will usually spontaneously move in the culture. In contrast, differentiated primary neurons form clusters with their cell bodies, and connect with other clusters with their axon cables.
 <img src="/files/ftp_upload/2215/2215fig1.jpg" alt="Figure 1" /> 
Figure 1. Primary cells undergo morphologic changes after being plated in the culture 
 (A) A bright field image of primary cells right after being plated in a well of a 384-well plate. The majority of cells are round and intact and well separated. (B) A bright field image of primary cell culture 20 hours after plating. Primary cells already form clusters (big arrowhead) and muscles with a branch-like morphology (long arrow) can be recognized at this stage. (C) A phase-contrast image of primary culture 5 days after plating. The culture looks much more advanced, with neuronal extensions (small arrows), neuronal clusters (big arrowheads) and muscles (medium-sized arrow).
 <img src="/files/ftp_upload/2215/2215fig2.jpg" alt="Figure 2" /> 
Figure 2. Examples of primary cells treated with dsRNAs cultured in a 384-well plate 
 Primary cells were isolated from the embryos carrying Dmef2-Gal4, D42-Gal4, UAS-mito-GFP transgenes, in which Dmef2-Gal4 and D42-Gal4 drive expression of mito-GFP in muscles and motor neurons, respectively. The cells were treated with dsRNAs targeting either lacZ (A-C) or inflated(if) (D-F). Both primary muscles and neurons can be seen by GFP (A,C,D,F). The white arrows point to the neuronal extensions. Phalloidin staining of actin (arrowheads) reveals a branch-like muscle structure in the culture treated with lacZ dsRNAs (B and C), but round-up muscle morphology in the culture treated with if dsRNAs (E and F). In the merged images (C, F), muscles are stained yellow, but red negative shows neurons and their extensions (white arrows).
 <img src="/files/ftp_upload/2215/2215fig3.jpg" alt="Figure 3" /> 
Figure 3. Confocal micrographs of primary muscles cultured on human vitronectin-coated cover glass slides 
Primary muscles were immunofluorescently stained with anti-Drosophila Titin (KZ) (red in the merge) and with phalloidin for actin (green in the merge). The top panels are the wild-type control muscle and bottom ones are the sals(CG31374) RNAi muscle with shortened
sarcomeres.

<ol>
	<li>Ashburner, M., Roote, J., Sullivan, W., Ashburner, M., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Laboratory culture of Drosophila. , 588-599 (1999).</li><li>Bai, J., Hartwig, J. H., Perrimon, N. S. A. L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=a+WH2-domain-containing+protein,+promotes+sarcomeric+actin+filament+elongation+from+pointed+ends+during+Drosophila+muscle+growth.">a WH2-domain-containing protein, promotes sarcomeric actin filament elongation from pointed ends during Drosophila muscle growth.</a> Dev. Cell. 13, 828-842 (2007).</li><li>Bai, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+interference+screening+in+Drosophila+primary+cells+for+genes+involved+in+muscle+assembly+and+maintenance+.">RNA interference screening in Drosophila primary cells for genes involved in muscle assembly and maintenance .</a> Development. 135, 1439-1449 (2008).</li><li>Sepp, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+neural+outgrowth+genes+using+genome-wide+RNAi.">Identification of neural outgrowth genes using genome-wide RNAi.</a> PloS. Genet. 4, e1000111-e1000111 (2008).</li><li>Donady, J. J., Fyrberg, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mass+culturing+of+Drosophila+embryonic+cells+in+vitro.">Mass culturing of Drosophila embryonic cells in vitro.</a> TCA Manual. 3, 685-687 (1977).</li><li>Seecof, R. L., Alleaume, N., Teplitz, R. L., Gerson, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differentiation+of+neurons+and+myocytes+in+cell+cultures+made+from+Drosophila+gastrulae.">Differentiation of neurons and myocytes in cell cultures made from Drosophila gastrulae.</a> Exp. Cell Res. 69, 161-173 (1971).</li><li>Bai, J., Sepp, K. J., Perrimon, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culture+of+Drosophila+primary+cells+dissociated+from+gastrula+embryos+and+their+use+in+RNAi+screening.">Culture of Drosophila primary cells dissociated from gastrula embryos and their use in RNAi screening.</a> Nature Protocols. 4, 1502-1512 (2009).</li></ol>

No conflicts of interest declared.

The developmental pattern of primary cultures of Drosophila cells can vary greatly, possibly due to the several variables during the primary cell preparation process. First of all, flies used for egg laying should be young (within a week old) and healthy (free of viral or bacterial infection). Unfertilized or infected embryos must be avoided, as they are useless and cells derived from those embryos cannot differentiate and will only survive for 1-2 days. Second, during the homogenization process, it is import...

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<td>Shields and Sang M3 medium</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>S3652</td>
<td>&nbsp;</td>
<tr>
<td>Fetal bovine serum</td>
<td>&nbsp;</td>
<td>JRH Biosciences</td>
<td>12103-500M</td>
<td>&nbsp;</td>
<tr>
<td>Insulin from bovine pancreas</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>I-6634</td>
<td>&nbsp;</td>
<tr>
<td>Large embryo collection cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific</td>
<td>59-101</td>
<td>&nbsp;</td>
<tr>
<td>#25 US standard testing sieve</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>57334-454</td>
<td>&nbsp;</td>
<tr>
<td>#45 US standard testing sieve</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>57334-462</td>
<td>&nbsp;</td>
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<td>#120 US standard testing sieve</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>57334-474</td>
<td>&nbsp;</td>
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<td>Dounce tissue grinder, Wheaton 7 mL</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>62400-620</td>
<td>&nbsp;</td>
<tr>
<td>Dounce tissue grinder,Kontes 40 mL</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>KT885300-0040</td>
<td>&nbsp;</td>
<tr>
<td>Dounce tissue grinder,Kontes 40 mL</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>KT885300-0100</td>
<td>&nbsp;</td>
<tr>
<td>Costar, 384-well plate</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>29444-078</td>
<td>&nbsp;</td>
<tr>
<td>Lab-Tek II Chambered coverglass</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>171080</td>
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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.

Watch this Scientific Journal Video about Primary Cell Cultures from Drosophila Gastrula Embryos at JoVE.com

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.

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 ...

primary-cell-cultures-from-drosophila-gastrula-embryos

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13.8K Views.  Harvard Medical School. 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.

Video: Primary Cell Cultures from Drosophila Gastrula 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 ...

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.

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

Primary Dissociated Midbrain Dopamine Cell Cultures from Rodent Neonates

The ability to create primary cell cultures of dopamine neurons allows for the study of the presynaptic characteristics of dopamine neurons in isolation from systemic input from elsewhere in the brain. In our lab, we use these neurons to assess dopamine release kinetics using carbon fiber amperometry, as well as expression levels of dopamine related genes and proteins using quantitative PCR and immunocytochemistry. In this video, we show you how we generate these cultures from rodent neonates.
The process involves several steps, including the plating of cortical glial astrocytes, the conditioning of neuronal cell culture media by the glial substrate, the dissection of the midbrain in neonates, the digestion, extraction and plating of dopamine neurons and the addition of neurotrophic factors to ensure cell survival.
The applications suitable for such a preparation include electrophysiology, immunocytochemistry, quantitative PCR, video microscopy (i.e., of real-time vesicular fusion with the plasma membrane), cell viability assays and other toxicological screens.

Primary dissociated midbrain dopamine cell cultures allow for the study of presynaptic characteristics of dopamine neurons. They can be used to monitor real-time dopamine release kinetics and protein/mRNA levels of regulators of dopamine exocytosis. Here, we show you how to generate these cultures from rodent neonates.

The ability to create primary cell cultures of dopamine neurons allows for the study of the presynaptic characteristics of dopamine neurons in isolation ...

The Preparation of Primary Hematopoietic Cell Cultures From Murine Bone Marrow for Electroporation

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser MXcell  electroporation system and Gene Pulser  electroporation buffer were specifically developed to transfect nucleic acids into mammalian cells and difficult-to-transfect cells, such as primary and stem cells.This video demonstrates how to establish primary hematopoietic cell cultures from murine bone marrow, and then prepare them for electroporation in the MXcell system. We begin by isolating femur and tibia. Bone marrow from both femur and tibia are then harvested and cultures are established. Cultured bone marrow cells are then transfected and analyzed.

This procedure describes how to establish primary hematopoietic cell cultures from murine bone marrow and is followed by transfection using the Gene Pulser MXCell electroporation system.

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser ...

Tissue Targeted Embryonic Chimeras: Zebrafish Gastrula Cell Transplantation

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups of cells in a larger context are altered.  Watching how one cell interacts with and behaves in a unique environment is essential to characterizing cell functions.  Determining how the localized misexpression of a specific protein influences surrounding cells provides insightful information on the roles that protein plays in a variety of developmental processes.  Our lab uses the zebrafish model system to uniquely combine genetic approaches with classical transplantation techniques to generate genotypic or phenotypic chimeras.  We study neuron-glial cell interactions during the formation of forebrain commissures in zebrafish. This video describes a method that allows our lab to investigate the role of astroglial populations in the diencephalon and the roles of specific guidance cues that influence projecting axons as they cross the midline. Due to their transparency zebrafish embryos are ideal models for this type of ectopic cell placement or localized gene misexpression. Tracking transplanted cells can be accomplished using a vital dye or a transgenic fish line expressing a fluorescent protein. We demonstrate here how to prepare donor embryos with a vital dye tracer for transplantation, as well as how to extract and transplant cells from one gastrula staged embryo to another.  We present data showing ectopic GFP+ transgenic cells within the forebrain of zebrafish embryos and characterize the location of these cells with respect to forebrain commissures.  In addition, we show laser scanning confocal timelapse microscopy of Alexa 594 labeled cells transplanted into a GFP+ transgenic host embryo.  These data provide evidence that gastrula staged transplantation enables the targeted positioning of ectopic cells to address a variety of questions in Developmental Biology.

Zebrafish cell transplantation enables the combination of genetics and embryology to generate tissue specific chimeras. This video demonstrates gastrula staged cell transplantations that have allowed our lab to investigate the roles of astroglial populations and specific guidance cues during commissure formation in the forebrain.

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups ...

Establishing Primary Adult Fibroblast Cultures From Rodents

The importance of using primary cells, rather than cancer cell lines, for biological studies is becoming widely recognized. Primary cells are preferred in studies of cell cycle control, apoptosis, and DNA repair, as cancer cells carry mutations in genes involved in these processes. Primary cells cannot be cultured indefinitely due to the onset of replicative senescence or aneuploidization. Hence, new cultures need to be established regularly. The procedure for isolation of rodent embryonic fibroblasts is well established, but isolating adult fibroblast cultures often presents a challenge. Adult rodent fibroblasts isolated from mouse models of human disease may be a preferred control when comparing them to fibroblasts from human patients. Furthermore, adult fibroblasts are the only available material when working with wild rodents where pregnant females cannot be easily obtained. Here we provide a protocol for isolation and culture of adult fibroblasts from rodent skin and lungs. We used this procedure successfully to isolate fibroblasts from over twenty rodent species from laboratory mice and rats to wild rodents such as beaver, porcupine, and squirrel.

This article describes a protocol for isolation and maintenance of primary fibroblast cultures from skin and lung tissue of wild rodents.

The importance of using primary cells, rather than cancer cell lines, for biological studies is becoming widely recognized.  Primary cells are preferred ...

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 ...

Imaging Cell Shape Change in Living Drosophila Embryos

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

Imaging Cell Shape Change in Living Drosophila Embryos

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

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

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.

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

Na&iuml;ve Adult Stem Cells Isolation from Primary Human Fibroblast Cultures

Over the last decade, several adult stem cell populations have been identified in human skin 1-4. The isolation of multipotent adult dermal precursors was first reported by Miller F. D laboratory 5, 6. These early studies described a multipotent precursor cell population from adult mammalian dermis 5. These cells--termed SKPs, for skin-derived precursors-- were isolated and expanded from rodent and human skin and differentiated into both neural and mesodermal progeny, including cell types never found in skin, such as neurons 5. Immunocytochemical studies on cultured SKPs revealed that cells expressed vimentin and nestin, an intermediate filament protein expressed in neural and skeletal muscle precursors, in addition to fibronectin and multipotent stem cell markers 6. Until now, the adult stem cells population SKPs have been isolated from freshly collected mammalian skin biopsies.
Recently, we have established and reported that a population of skin derived precursor cells could remain present in primary fibroblast cultures established from skin biopsies 7. The assumption that a few somatic stem cells might reside in primary fibroblast cultures at early population doublings was based upon the following observations: (1) SKPs and primary fibroblast cultures are derived from the dermis, and therefore a small number of SKP cells could remain present in primary dermal fibroblast cultures and (2) primary fibroblast cultures grown from frozen aliquots that have been subjected to unfavorable temperature during storage or transfer contained a small number of cells that remained viable 7. These rare cells were able to expand and could be passaged several times. This observation suggested that a small number of cells with high proliferation potency and resistance to stress were present in human fibroblast cultures 7.
We took advantage of these findings to establish a protocol for rapid isolation of adult stem cells from primary fibroblast cultures that are readily available from tissue banks around the world (Figure 1). This method has important significance as it allows the isolation of precursor cells when skin samples are not accessible while fibroblast cultures may be available from tissue banks, thus, opening new opportunities to dissect the molecular mechanisms underlying rare genetic diseases as well as modeling diseases in a dish.

Na&#239;ve Adult Stem Cells Isolation from Primary Human Fibroblast Cultures

We report a method to isolate na&#239;ve multipotent skin-derived precursor (SKP) cells from primary human fibroblast cultures. We show that these SKPs derived from fibroblast cultures share similar stem cell properties to the ones derived directly from human skin biopsies. These cells express the neural crest marker, nestin, in addition to the multipotent markers such as OCT4 and Nanog.

Over the last decade, several adult stem cell  populations have been identified in human skin 1-4.  The isolation of multipotent adult dermal precursors ...