Name: Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae (Video) | JoVE Journal
Uploaded: 2007-05-28T00:00:00.000+00:00
Duration: 15 min 12 s
Description: Learn to prepare primary neuronal cultures from late stage Drosophila pupae brains for neuroscience research. This method enables neurite outgrowth observation and calcium imaging studies.

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5 minutes

This work was supported by NIH grant NS27501 to DKOD. Additional support for this work was provided by a grant to UC Irvine in support of DKOD through the HHMI Professor Program.

Preparations before day of culturing:
<ol>
<li>Make sterile dissecting solution.</li>
<li>Make sterile DMEM and keep in 10 ml aliquots at 4&deg;C for 2 weeks.</li>
<li>Make sterile DDM2 supplements and freeze in 50 or 100 &micro;l aliquots for 1 month.</li>
<li>Make ConA/laminin.</li>
<li>Coat coverslips. &nbsp;&nbsp;&nbsp;&nbsp; Optional: Make sterile CNBM and store frozen for up to 4 months. </li>
</ol>
On day of culturing
I.&nbsp; Prepare Enzyme Solution (ES) in laminar flow hood
<ol>
<li>Put 5 ml of dissecting solution (DS) in a 15 ml centrifuge tube.</li>
<li>Weigh 0.8 mg of L-Cysteine in a 0.6 ml eppendorf and add 150 &micro;l of DS. Pipette up and down until crystals are gone.</li>
<li>Add 150 ml of DS/Cysteine to the 5 ml DS and mix well.</li>
<li>Add 50 U (units) of Papain.</li>
<li>Add 7 ml of 0.1 N Na OH and mix well. Solution will clear within 30 min. when activated.&nbsp;</li>
<li>Filter enzyme solution with 0.2 mm syringe filter into a sterile 1.5 ml screw cap microfuge tube &nbsp; &nbsp;&nbsp; Papain: Want 50 U in 5 ml DS. Each batch is slightly different, so must calculate how much: &nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp; 37.3 mg/ml and 28.3 U/mg &nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp; 37.3 x 28.3 = 1055.59 U/ml&nbsp;&nbsp; (1000 ml) &nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp;&nbsp; 50 U&nbsp; = 47.4 ml </li>
</ol>
II. Make DDM2 media
On day of culturing: add the following supplements to make DDM2
To 10 ml of DMEM add the supplements, just before culturing:
<ol>
<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&nbsp; Insulin</li>
<li>10&micro;l 20-Hydroxyecdysone</li>
</ol>
Note: Make enough DDM2 for all cultures planned (each brain plated on a single coverslip in an individual 35 mm Petri dish, 1.5 ms of media/culture)
Step III-VI can be at a done nonsterile lab bench and should be completed in 45 minutes or less.
III. Pupae Collection
<ol>
<li>Select 10-15 pupae from sides of vial under a dissecting microscope.</li>
<li>Place pupae in the lid of a dry 35 mm Petri dish.&nbsp;</li>
</ol>
Note: We generally use brains from pupae between 55-78 hours after puparium formation (APF). It is slightly harder to dissect out the brain from the younger pupae since the head capsules tend to collapse, while the overall level of neurite outgrowth is slightly lower from the oldest pupae. In the Canton-S wildtype strain, these stages are recognized by pigmented eyes that are light brown to slightly reddish, with little pigment elsewhere.
IV. Decapitation under dissecting microscope
<ol>
<li>Place two drops of sterile dissecting solution in lid of Petri dish, next to pupae.</li>
<li>Gently hold a single pupa with fine forceps in left hand and use a 27 gauge syringe needle attached to a 1 cc syringe in the right hand to remove the cuticle at the anterior end of pupa.</li>
<li>Squeeze pupa slightly with forceps and as head emerges use 27 gauge needle to sever neck.</li>
<li>With tip of the needle or the forceps, transfer head to drop of dissecting solution.</li>
<li>Repeat until you have 10-15 heads in a single drop.</li>
</ol>
V. Removal of the brain from head under dissecting microscope
<ol>
<li>In each hand hold a 1 cc syringe with a 27 gauge needle.</li>
<li>Use these to position the fly head in the drop of dissecting solution so the proboscis is facing you and the dorsal surface of the head is north.</li>
<li>Insert the left needle into the proboscis to pin the head down and use the right needle to make a slit in the right eye that goes from the ventral to the dorsal surface.</li>
<li>Insert the right needle near the left in the region of the proboscis and then use the left needle to gently depress the cuticle over the left eye, pushing the brain toward the slit in the right eye. The brain should emerge from the slit on the right, relatively intact, often with both optic lobes still attached and sometimes the pigmented eye tissue as well.</li>
<li>Push the brain onto the back of the needle and transfer to a drop of sterile dissecting saline in a new sterile, 35 mm Petri dish.</li>
<li>Collect 10-15 brains for each genotype. </li>
</ol>
VI. Removal of optic lobes and enzyme treatment
<ol>
<li>In each hand hold a 1 cc syringe with a 27 gauge needle and use these to gather all brains in one small area of the drop of dissecting solution</li>
<li>Use syringes as cutting tools to remove the optic lobes from each brain so the tissue remaining for culture is the central brain region.</li>
<li>Use a 20 &micro;l pipetman, set at 5 &micro;l, to transfer all the brains of a single genotype to an eppendorf tube containing 1 ml of sterile papain solution. </li>
<li>Brains incubated in papain for 10-15 minutes on a rotator set at 60 rpm.</li>
<li>Centrifuge at 3000 rpm for 3 minutes. </li>
</ol>
Inside sterile laminar flow hood &ndash; all steps where tissue is exposed to air should be done in laminar flow hood for the remaining steps.
VII. Washing
<ol>
<li>Remove enzyme solution and replace with sterile dissecting solution.</li>
<li>Centrifuge at 3000 rpm for 3 minutes.</li>
<li>Wash with sterile dissecting solution a total of 3 times.</li>
<li>Remove dissecting saline and replace with DDM2.</li>
<li>Centrifuge and wash a total of 2 times with DDM2.</li>
<li>Transfer brains and media to a sterile 35 mm Petri dish. </li>
</ol>
VIII. Trituration and Plating under dissecting microscope
<ol>
<li>Place 5 ul of DDM2 in the center of a coverslip.</li>
<li>Transfer 1 brain to this drop of DDM2 with a yellow tip.</li>
<li>Under the microscope, use 27 gauge needles to dice up brain into small pieces (should take &lt;1 minute).</li>
<li>Under visual guidance, break the tip of a&nbsp; glass pipet that has been pulled to a fine point so that the larger pieces can be gently sucked into the tip of the pipet and expelled back into the medium. We use mouth suction tubes that come standard with 100 &micro;l hematocrit capillary glass. Note: Take care not to get bubbles in the media and you will need to empirically determine the best size pipet to use. Good ones allow you to dissociate the brain with some individual cells and still some large clumps, in about 30 seconds/brain. When you look at these at higher power, approximately 10-20% of the neurons should still have some processes remaining and there will still be some large clumps. If you dissociate too much, then all the cells will be round and they will have sustained so much damage they will not survive. This step takes practice and must be done quickly since there is a large surface-to-volume ratio and the osmolality and pH of the drop of DDM2 can change quickly. Tissue needs to be placed in CO2 incubator as quickly as possible. </li>
<li>Write on the petri dish lid: "Genotype, date and time".</li>
<li>Put the 35 mm petri dish in a 100 mm petri dish (with wet kimwipe) and let settle in the CO2 incubator for 30 min.</li>
<li>Flood the 35 mm dish with 1.5 ml of DDM2.</li>
<li>Keep cultures in the incubator at 5% CO2 incubator (23&deg;C). Note: If you don&rsquo;t have access to a cooling CO2 incubator, use a standard mammalian tissue culture incubator and either put it in a cool room and heat to 23&deg;C, or put in the lab, turn off temp, and use ice in a tray at bottom of incubator to regulate temp around ambient. </li>
</ol>
IX. Feeding Cells
<ol>
<li>On Day 1 (next day), add 0.5 ml of CNBM/B27 (Conditioned Media) to make 3:1 DDM2:CNBM/B27.</li>
<li>On Day 5 replace 1 ml of media for 3:1 DDM2:CNBM/B27.</li>
<li>Keep feeding cells with 3:1 DDM2:CNBM/B27 every 4-5 days. Note: Cultures can be grown without the non-neuronal conditioned media but the neurons appear a bit more fragile and are more difficult for use in electrophysiology experiments.</li>
</ol>

<ol><li><a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=%22Culturing+Nerve+Cells%22">Culturing Nerve Cells</a>. Banker, G., Goslin, K. , MIT Press. Cambridge. (1991).</li>
<li>Rohrbough, J., O'Dowd, D. K., Baines, R. A., Broadie, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cellular+bases+of+behavioral+plasticity%3A+Establishing+and+Modifying+synaptic+circuits+in+the+Drosophila+Genetic+System%5BTitle%5D)+AND+%22J.+Neurobiol%22%5BJournal%5D)">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/pubmed?term=((Voltage-gated+currents+and+firing+properties+of+embryonic+Drosophila+neurons+grown+in+a+chemically+defined+medium%5BTitle%5D)+AND+%22J.+Neurobiol%22%5BJournal%5D)">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/pubmed?term=((Fast+excitatory+synaptic+transmission+mediated+by+nAChR+in+cultured+Drosophila+neurons%5BTitle%5D)+AND+%22J.+Neurosci%22%5BJournal%5D)">Fast excitatory synaptic transmission mediated by nAChR in cultured Drosophila neurons.</a> J. Neurosci. 19, 5311-5321 (1999).</li>
<li>Su, H., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fast+synaptic+currents+in+Drosophila+mushroom+body+Kenyon+cells+are+mediated+by+alpha-bungarotoxin-sensitive+nAChRs+and+picrotoxin-sensitive+GABA+receptors%5BTitle%5D)+AND+%22J.+Neurosci%22%5BJournal%5D)">Fast synaptic currents in Drosophila mushroom body Kenyon cells are mediated by alpha-bungarotoxin-sensitive nAChRs and picrotoxin-sensitive GABA receptors.</a> J. Neurosci. 23, 9246-9253 (2003).</li>
<li>Jiang, S. A., Campusano, J. M., Su, H., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Drosophila+mushroom+body+Kenyon+cells+generate+spontaneous+calcium+transients+mediated+by+PLTX-sensitive+calcium+channels%5BTitle%5D)+AND+%22J.+Neurophysiol%22%5BJournal%5D)">Drosophila mushroom body Kenyon cells generate spontaneous calcium transients mediated by PLTX-sensitive calcium channels.</a> J. Neurophysiol. 94, 491-500 (2005).</li>
<li>Campusano, J. M., Su, H., Jiang, S. A., Sicaeros, B., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((nAChR-mediated+calcium+responses+and+plasticity+in+Drosophila+Kenyon+cells%5BTitle%5D)+AND+%22Develop+Neurobiol%22%5BJournal%5D)">nAChR-mediated calcium responses and plasticity in Drosophila Kenyon cells.</a> Develop Neurobiol. 67, 1520-1532 (2007).</li>
<li>Oh, H. -W., Campusano, J. M., Hilgenberg, L. G. W., Sun, X., Smith, M. A., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ultrastructural+analysis+of+chemical+synapses+and+gap+junctions+between+Drosophila+brain+neurons+in+culture%5BTitle%5D)+AND+%22Develop+Neurobiol%22%5BJournal%5D)">Ultrastructural analysis of chemical synapses and gap junctions between Drosophila brain neurons in culture.</a> Develop Neurobiol. , (2007).</li>
</ol>

Neurons harvested from the brains of embryonic/postnatal rodents can be grown in primary cell culture where they extend neurites and form functional synaptic connections. Methods for preparation of these cultures are well established and studies in rodent neuronal cultures have played a critical role in identifying genes and environmental factors involved in regulation of synapse formation and function (Banker and Goslin, 1991). While insect neurons from a variety of species can also be grown in culture, the only insect ...

<table><tbody><tr><td>Concanavalin A</td><td></td><td>Sigma-Aldrich</td><td>C-2010</td><td>To 2.5 mL DS, add 25 mg Concanavalin A bottle. This is 10 mg/ml concentration.Make aliquots of 90 ul and store at -20 &amp;deg;C, no longer than 3 months.</td></tr><tr><td>Laminin</td><td></td><td>Sigma-Aldrich</td><td>L-2020</td><td>Add 1 mL DS to 1mg Laminin bottle. This is 0.5 mg/mL concentration.Make aliquots of 10 &amp;mu;L and store at -20 &amp;deg;C, no longer than 3 months.</td></tr><tr><td>Coverslips</td><td></td><td>Bellco Glass</td><td>1943-00012</td><td>12 mm glass coverslips</td></tr><tr><td>ConA + Laminin solution</td><td></td><td></td><td></td><td>Stock solution, from Con A stock and laminin stock, for coverslip coating: Add in 5 mL DS, 83.5 &amp;mu;L of ConA (167 &amp;mu;g/mL) and 8.35 &amp;mu;uL of Laminin (0.835 &amp;mu;g/mL). Mix. Make aliquots of 100 &amp;mu;L and store at -20 &amp;deg;C, not longer than a month.</td></tr><tr><td>Dissecting Solution</td><td>Buffer</td><td></td><td></td><td>For 500 ml: 400 ml Ultra filtered water + 25 ml Stock Solution A + 14 ml Stock Solution B + 3.0 g (33.3 mM) D (+)-Glucose (Sigma G-8270) + 7.5 g (43.8 mM) Sucrose (Sigma S-0389). Adjust pH to 7.4 with 1N NaOH (around 2 ml). Bring final volume to 500 ml with ultra filtered water. Decant into a clean glass bottle and autoclave. Label &quot;Dissecting Solution&quot; and store at 4?C. </td></tr><tr><td>Dissecting Solution</td><td>Buffer</td><td></td><td></td><td>For 500 ml: 400 ml Ultra filtered water + 25 ml Stock Solution A + 14 ml Stock Solution B + 3.0 g (33.3 mM) D (+)-Glucose (Sigma G-8270) + 7.5 g (43.8 mM) Sucrose (Sigma S-0389). Adjust pH to 7.4 with 1N NaOH (around 2 ml). Bring final volume to 500 ml with ultra filtered water. Decant into a clean glass bottle and autoclave. Label &quot;Dissecting Solution&quot; and store at 4?C. </td></tr><tr><td>Solution B: HEPES</td><td>Buffer</td><td>Sigma-Aldrich</td><td>H-3375</td><td>20.97g (9.9mM). Add ultra filtered water up to 200 ml. Mix until dissolve and bring final volume to 250 ml. Place in a clean bottle and autoclave. Label &quot;Solution B&quot; and store at 4 C. </td></tr><tr><td>Solution A</td><td>Buffer</td><td></td><td></td><td>For 500 ml: 80.0g (137 mM) NaCl (Sigma S-9625) + 4.0g (5.4mM) KCl (Sigma P-4504) + 0.24g (0.17mM) Na2HPO4 (Sigma S-0876) + 0.3g (0.22 mM) KH2PO4 (Sigma P-5379).Weigh out all ingredients and mix until dissolved with 400 ml ultra filtered water. Bring final volume to 500 ml. Place in a clean bottle and autoclave. Label &quot;Solution A&quot; and store at 4&amp;deg;C. </td></tr><tr><td colspan="5">Coating Coverslips:Put autoclaved coverslips in a 60 mm petri dish.Pipet 5 ul of Con A/Laminin mix onto center of each coverslip.Place in 37 C incubator for 2 hours.Rinse 3x coverslips with 100 ul of autoclaved water each. Use vacuum attached to a sterile Pasteur pipet.During the last rinse, pick up the coverslip with forceps and dry both sides.Transfer the coverslip to a 35mm Petri dish.Store at room temperature for up to a month.</td></tr></tbody></table>

primary neuronal cultures from the brains of late stage drosophila pupae

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 ...

My name is Diana Dowd and I'm a professor in the departments of Anatomy and Neurobiology and developmental and cell biology here at uc Irvine. Today I'm going to tell you about the preparation of neuronal cultures from Drosophila Pupi. This involves removing the brain from late stage pupi and we generally culture just the central brain region.And at this stage, the major anatomical structures that are present in the adult brain are already formed. This includes the antenna lobes where there are projection neurons that send their axons all the way up to the mushroom body region and the mushroom body neurons, Kenyan cells, which have sent their axons down to their target neurons through the Alpha, beta and gamma lobes. On the day of culturing, The first thing we do is go into the laminar flow hood and we prepare sterile, the enzyme solution that we're gonna use to dissociate the tissue.And we also make up the medium that we're gonna use for growing the cells. And this is made from stalks that we've prepared ahead of time. The next step is to collect the pupi that we're gonna use for this dissociate or for the dissection.And that's done outside of the hood because these are grown in vials that contain yeast and all sorts of other things. And so we actually do the initial stages of our dissection Outside of the hood. These are peepee that I've just taken off the sides of a vial, and on this left hand row we have animals that have red eyes and they are a good stage for culturing.These ones are slightly, these PPP over here are slightly younger. They have orange or yellow eyes, and this is an okay 1 0 7 line. They have slightly brighter eyes than regular wild types where the color at these stages would be a little bit more brown.All of these stages are fine for culturing, for making the people Brain cultures. Okay, the first step here is To put a couple of drops of dissecting solution right next to the pupi that I have pulled out and lined up in my Petri dish. We use the tops of 35 millimeter Petri dishes.I then take a pair of no number five forceps in my left hand and a one CC tuberculin syringe with a 27 gauge needle at the end. These are my two dissecting tools. And to remove the head from the pupil case or from the pupi, basically I have to use the i, I hold the pupi with my forceps in my left hand, and then I use the needle in my right hand to remove the front of the pupil case.I then put the pull back the the cuticle covering and the head pops out the front. I use my needle then to sever the I at the neck region and I pick up the head and place it into the drop of dissecting saline. I will repeat this procedure 10 to 15 times because to make for a good culture, I like to have 10 to 15 animals of each genotype.All right, I'm going to Remove the brain from the head of the animal. To do that, I use two tuberculin syringes each holding a 27 gauge needle. The left needle I place right through the animal's psis to pin the head down to the Petri dish bottom.And then I use my right needle to open a slit on the right side, and that is where I'm going to push the brain out. Okay, and I'm, the next step then is to put my left needle over the the left eye. And now I'm going to actually just ex push and the brain should come out the right side through the slit.Here it comes. You can see it on the right hand side, and now I can just use my right needle to separate it from the rest of the head capsule and the eyes.Okay. Now, before doing the final dissection, which is to remove the optic lobes, I transfer the brain to a clean drop of saline.I can either use a pipe eman to do that, and I pick it up in the very tip of a 20 microliter pipe eman, and I use it set at five microliters, or I can actually transfer it on the needles As well. This is the last stage in The dissection procedure. It's done in a fresh drop of saline, and generally I'd have about 10 to 15 brains in here.I then remove the optic lobes from each one by using the 27 gauge needles as cutting tools, and I just sever the left optic lobe and then I sever the right optic lobe on each brain. And at this point, all of the brains that have been processed this way then will be transferred into an tube that contains the enzyme solution, which will be used for the enzymatic dissociation. All of the brains from a single genotype are put into a single epi screw cap einor tube that contains one mill of the enzyme solution.And then we just stick them into a a test tube holder and put them on a rotator where they sit for 10 to 15 minutes at room temperature. And that's the actual time of Dissociation. To complete the Enzyme dissociation step, we actually have to remove the enzyme.So we bring the tubes over to an EOR centrifuge, place the tubes in the centrifuge, and we are going to spin them down for three minutes at 3000 RPM. After we spin the brains down, we bring them now to our laminar flow hood. This is where the sterile part of the procedure begins.We now don't open the tube unless we are in the laminar flow hood, and we will now remove the enzyme solution. We'll replace it with dissecting solution, and then we will spin and wash a number of times with both dissecting solution and ending up with a Defined medium, which is what we grow the cells in. All right.The first thing I Need to do here is I need to put a five microliter drop of saline on each of the cover slips that I have in these Petri dishes. There's a single cover slip per Petri dish, and it's been coated with con a laminin, so I put a five microliter drop right in the middle of the cover slip. That's where my drop of laminin is.It's very important cause you want to concentrate your cells right in the middle of the field because we get about 10, 000 cells per brain, and that is what we make a single culture with. Okay, so now looking in the microscope, here are nine brains that have already been gone through the enzyme dissociation procedure. You can see they're still pretty much intact.They don't have their optic lobes. Those were removed, but we just have the central brain region and now I'm going to pipette up a single brain and move it over to the five microliter drop on a cover slip. And I do that four times.I take two one CC syringes put on the 27 gauge needles. Again, it's really important to note that these are very cheap and disposable dissecting tools, so undergraduates can do this without worrying about your budget. Okay, so now I have the two needles and I'm going to do the initial mechanical dissociation, which is to just cut this brain up into several pieces, which will allow me to actually suck these pieces up into a glass pipette in just a minute to complete the final dissociation.When I'm doing this, I don't get them down to tiny pieces, I just want to increase the decrease the total size so I can get a little bit so that the pipette, when I do the dissociation brings up slightly smaller pieces than the full brain. I also wanna do it relatively quickly because you see the drops of media are small. That means we have a high surface to volume ratio and the both the pH and the osmolarity of the solution can change if I leave it sitting out for any length of time.When I was first doing this, I would only actually do two brains at a time, two cultures at a time. Now I usually do four, and now I'm gonna break the pipette, but I can't talk about it or anything, right? This one I've just finished.The dissociation, the size of the pipette was pretty good. I was able to suck the tissue up and up and down a couple of times, and you can see individual cells and those, we still have clumps, but that's okay. They'll wash away.Now, the cultures, as soon as they've been plated, are placed into a second Petri dish and they're placed in the CO2 incubator within a couple of minutes because we wanna make sure that we maintain the pH and the osmolarity at a level that supports their survival. So we stick em in here for 30 minutes before we flood the dish. These cultures were Prepared from the brains of late stage pupi.The culture has been grown for two days in a CO2 incubator, and you can see individual cells that have extended processes. The extent of process elaboration is less than what you would see at five days and more than What you would see at one day. My name is Jorge Ano.I work in the Doubts lab. I'm gonna show you right now how we do the experiment, the calcium imaging experiment in the people cultures that we just prepared. This is a video of a calcium imaging experiment where you see blue cells showing low, indicating low calcium levels and red cells indicating high calcium levels.We'll see that there are some cells that turn to red and it indicates intra increasing intracellular calcium levels. And those are spontaneous and those are transient. They go back to to Baal calcium levels over time, and after a while, we're gonna apply a drug, in this case, nicotine, and you will see the cells increase the intracellular calcium levels.We apply nicotine for 100 seconds, then we watch out nicotine, and then the cells go back to basal calcium levels. So the real advantage Of this culture system is that we're able to make our cultures from both transgenic and mutant lines of flies, different ones in which different subpopulations of neurons are labeled with GFP. This is the brain from one of the lines we've used in which GFP is expressed in the mushroom body, Kenyan cells, neurons involved in mediating complex behaviors, including learning and memory.The cell bodies are located up in the dorsal aspect of the brain around the region where the red arrows are. And these L-shaped structures that are GFP positive are actually the axonal projections from these mushroom body Kenyan cells. The after dissociation of this brain tissue.The neurons regenerate their processes in culture, and you can easily identify the GFP positive Kenyan cells in this case. These are accessible to both then standard electrophysiological recording and calcium imaging studies. Therefore, these cultures are really useful tools for exploring the genetic and environmental factors that are important in regulating function of neurons that we know are important in mediating complex behaviors in the Adult animal.

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Research

JoVE Journal

Biology

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.

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This article describes a method by which one can mimic in vivo development of the Drosophila mushroom body in an ex vivo culture system.

Ex vivo Culturing of Whole, Developing Drosophila Brains

This video demonstrates a protocol to identify exogenous factors that initiate the reactivation of Drosophila brain neural stem cells from their quiescent state. Freshly hatched larvae are dissected to isolate their brains, which are then cultured in media with or without a test hormone to evaluate the hormone's potential to induce neural stem cell reactivation and proliferation.

Analyzing Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae.

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Live Imaging of Drosophila Larval Neuroblasts

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

This article provides detailed protocols for inflicting Penetrating Traumatic Brain Injury (PTBI)  to adult Drosophila and examining the resulting neurogenesis.

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

A method to reactivate quiescent neural stem cells in cultured Drosophila brain explants has been established. Using this method, the role of systemic signals can be uncoupled from tissue-intrinsic signals in the regulation of neural stem cell quiescence, entry and exit.

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Learn to prepare primary neuronal cultures from late stage Drosophila pupae brains for neuroscience research. This method enables neurite outgrowth observation and calcium imaging studies.

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

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