We would like to thank Changhui Pak and Alysia Vrailas Mortimer for initially teaching SMK the brain dissection technique. We also thank members of Ken Moberg&#39;s lab group, especially Chris Rounds, for critically reading the manuscript. Antibodies recognizing Fas2 (1D4) and chaoptin (24B10) were obtained from the Developmental Studies Hybridoma Bank. Antibody 24B10 was deposited to the DSHB by Seymour Benzer and Nansi Colley24,60,61, antibody 1D4 was deposited to the DSHB by Corey Goodman 22,23,56. Fly stocks were obtained from the Bloomington Stock Center. We also wish to thank the Ohio Agricultural Research and Development Center (OARDC) MCIC Imaging Center for use of the confocal microscope to image the brains in Figure 4A and B. SMK is supported by a grant from NICHD (1 R15 HD084241-01A1). M. Kahl was supported by a grant from the NSF (NSF REU #1560061).

1. Drosophila melanogaster Genetics and Optional Heat Shock Procedures
<ol>
	<li>Once flies have been crossed and F1 progeny have hatched, obtain females and/or males of the appropriate genotype. Depending on the brain region being investigated, flies should be collected daily and separated by sex so that age-dependent and/or sexually dimorphic patterns of brain connectivity can be more easily distinguished. 
	NOTE: Optional: If flies are being used for Mosaic Analysis with a Repressible Cell Marker (MARCM) analysis12,47, embryos, larvae, or pupae should be heat shocked at 37 &#176;C for 30 - 45 min to induce mitotic recombination. In order to target specific regions of the mushroom bodies for MARCM analysis, heat shock should be timed according to the schedule determined by12 and outlined in Figure 5B. If using MARCM to investigate brain regions other than the mushroom bodies, pilot experiments should be performed to determine the optimal stage to be heat-shocked. While the steps below are written in regard to eclosed flies, pharate adults can also be dissected using these steps once the pupal case has been removed.</li>
</ol>
2. Dissection Station Preparation
<ol>
	<li>Position the stereomicroscope and light source with attached fiber optic goosenecks on a large benchtop. To promote steady hand movements and reduce hand &#34;shake&#34; while dissecting, it is essential that adequate hand and arm rest space is available around the microscope. Ensure that there is approximately 8 - 10 inches on either side of the microscope and 4 - 6 inches between the base of microscope and edge of the bench.</li>
	<li>Fill 2 or 3 wells of a glass 9-well or 3-well dish with 1.0 mL of PTN buffer (0.1 M Sodium Phosphate Buffer pH 7.2, 0.1% nonionic surfactant, see Materials Table for complete buffer components) and place next to the dissecting station on ice. Newly dissected brains will be transferred to this dish and stored until the fixation step.
	<ol>
		<li>Note: If live imaging is required, dissection buffer should be 1x Phosphate Buffered Saline (PBS) or HL3 buffer48. If intracellular protein localization is required, PBS can also be used as an alternative buffer for dissection and fixation. Following fixation, permeabilization of cell membranes should then be performed using PTN washes containing 0.1% or 0.3% nonionic detergent.</li>
		<li>If PBS is used for dissections and fixation, pipette tips should be rinsed at least one time with a detergent-containing buffer (such as PTN) to prevent brains from sticking to the plastic pipette tips during transfer to microcentrifuge tubes.</li>
	</ol>
	</li>
	<li>Using an empty 35 mm glass or plastic petri dish, construct a dissection dish containing a silicone elastomer. Briefly, mix the elastomer components according to the manufacturer&#39;s directions, pour into 35 mm dishes, and let it polymerize overnight on a flat surface. Elastomer containing dissection dishes should be used to protect the fine tips of dissecting forceps, which can easily be damaged if contact is made between the forceps and a harder surface, such as a glass dish. We also regularly purchase commercially available silicone coated dishes from online retailers. To increase contrast during dissections, silicone elastomer dissection dishes containing inactivated charcoal (and thus colored black) are particularly useful.</li>
</ol>
3. Adult Brain Dissection Procedure
<ol>
	<li>Anesthetize 3 - 5 day old adult D. melanogaster with CO2 or by using ice. If using ice, place the vial containing flies upside down (plug end down) into an ice bucket for ~5 min. Placing the vial into ice upside down prevents flies from becoming lodged in the food. Once flies have been anesthetized, place the flies on a cold metal pad or petri dish sitting in ice or on a CO2 emitting fly pad. If dissecting brains to analyze neurodegeneration, older flies may also be used.</li>
	<li>Place a small amount (150 - 200 &#181;L) of PTN in the center of the dissection dish using a transfer pipette or a p200 pipette to create a &#34;bubble&#34; of PTN. Place the dissection dish under the stereomicroscope and adjust the lighting and focus so that the bubble of PTN fills the field of view and is uniformly illuminated.</li>
	<li>Manipulate flies so that they are &#34;belly up&#34; (i.e., ventral side up) while lying on the metal or CO2 pad.</li>
	<li>Using one pair of #5 forceps, grasp the abdomen of a fly to be dissected and, keeping hold of the fly, completely submerge it in the PTN on the dissection dish. 
	NOTE: For the remainder of the protocol, all steps should be performed while the head is submerged in PTN.</li>
	<li>Using a second pair of #5 forceps, grasp the base of the fly proboscis and pull the two pairs of forceps apart to detach the fly head from the body. Discard the abdomen and thorax. During this step, it is critical that the head is not released and allowed to float on the surface of the PTN. Once the head is floating, it can be very difficult to grasp again without crushing the brain. 
	NOTE: Using this method, connections between the brain and ventral nerve cord are severed. If intact connections between these regions of the CNS are required, an alternative dissection protocol, such as49,50, should be followed. If the proboscis detaches from the fly head before the head is removed, there will be a hole where the proboscis was. In this case, grasp the fly head at the edge of the hole near one eye. Then remove the head using a moderate amount of force while pulling the two pairs of forceps apart from one another. Occasionally, when the head is removed from the body, the gut and/or ventral nerve cord remains attached to the head and should be removed before continuing the dissection.</li>
	<li>While one pair of forceps grasps the proboscis, the second pair should grasp the medial edge of the right fly eye. Slowly, pull the forceps apart from one another. This step should be performed with a small amount of steady lateral force. As the forceps slowly move apart from one another, the proboscis should pull away from the head and create a central hole in the head cuticle. Discard the proboscis with the first pair of forceps without releasing the medial portion of the right eye from the second pair. 
	Note: The adult D. melanogaster brain is in the caudal (i.e., posterior/rear) region of the fly head. Thus, grasping that region of the head should be avoided. Ideally, only the rostral (i.e., front) portion of the head near the medial retina should be directly grasped by the forceps. The brain and associated trachea should now be visible through the central hole in the cuticle. At this time, any white stringy threads of trachea protruding from the hole can be removed and discarded.</li>
	<li>With the second pair of forceps, grasp the medial edge of the left retina (at the edge of the central hole in the head cuticle). To remove the retinas and associated cuticle, slowly pull the forceps away from one another at a 180&#176; angle. As the retina dissociates from the underlying optic lobe, you should feel a slight decrease in tension. Proceed slowly to prevent tearing the optic lobe. 
	NOTE: Separating the forceps too quickly during this step may result in the tearing of the optic lobe or disruption of mushroom body structures. Occasionally, the cuticle will be removed but pieces of the retina will remain attached to the optic lobe. If imaging the mushroom bodies, it is not completely necessary to remove the entire retina. However, analysis of other brain regions (such as retinal neuron innervation of the optic brain lobes) may require the retina to be completely removed as described by48,51. 
	NOTE: As the retina is slowly separated from the underlying optic lobe of the brain, the optic lobe should be observable as an opaque white structure covered with white, stringy trachea. Once one retina has been removed, it can be discarded. When analyzing pathfinding of retinal neurons, particular care should be taken during this step to prevent damage to the optic lobe. An additional protocol focused on dissection and live imaging of the photoreceptor neurons is also available48.</li>
	<li>Now, carefully remove as much of the visible trachea as possible. The trachea may already contain or later fill with air, causing brains to float and, potentially, be lost during later immunostaining steps. To remove trachea, pick it off the brain using a very sharp pair of #5 forceps.</li>
	<li>Remove the remaining retina and surrounding cuticle by using both pairs of forceps to grasp the medial region of the left fly retina. Carefully tear the retina in half to remove pieces of the retina and cuticle. In some cases, removing the remaining cuticle without crushing the brain proves especially challenging. In these cases, we have found that the remaining strands of the ventral nerve cord can instead be grasped by one pair of forceps while the other pair of forceps is used to carefully remove the last of the cuticle.</li>
	<li>Using a p200 pipette, move dissected brains to one well of the 9- or 3-well dish containing PTN. Brains of the same genotype should be pooled together into the same well and kept on ice. Brain tissue should be fixed within one hour of dissection. Brains can be fixed in small batches and pooled if a larger number of brains is required. In most circumstances, an experienced researcher can usually dissect a brain and transfer it to the glass collection dish in approximately 3 - 5 min.</li>
</ol>
4. Fixation and Immunofluorescent Staining Procedure
<ol>
	<li>Using a p200 pipette, transfer dissected brains from the 9-well dish to a 0.5 mL microcentrifuge tube filled with 0.5 mL of 4% paraformaldehyde diluted in PTN. At least 10 - 15 brains of the same genotype can be combined into one microcentrifuge tube. All remaining steps (until mounting of brains onto slides) are completed in 0.5 ml microcentrifuge tubes. 
	CAUTION: Paraformaldehyde (PFA) should be handled in a fume hood. PFA waste should be saved and disposed of properly. 20% paraformaldehyde purchased in glass ampules can be aliquoted into microcentrifuge tubes and stored at -20 &#176;C until needed.</li>
	<li>Inside a fume hood, incubate brains in 4% paraformaldehyde for 20 min with slow speed rocking at room temperature.</li>
	<li>Following fixation, allow brains to settle to the bottom of the microcentrifuge tube by gravity. 
	Note: Occasionally, the brains may stick to the side of the microcentrifuge tube. If this occurs, it is usually helpful to laterally rotate the tube between the index finger and thumb or to very gently tap the tube on the bench to promote sinking of the brains.</li>
	<li>Remove fixative using a p1000 pipette and perform two &#34;quick&#34; washes with 500 &#181;L of PTN, allowing the brains to settle to the bottom of the microcentrifuge tube between washes. During these quick washes, once all the brains have settled by gravity to the bottom of the microcentrifuge tube, the PTN can be immediately exchanged for fresh buffer; no additional wash time is required. 
	NOTE: Typically, leaving extra buffer in the tube is preferable to risking brain removal. Careful examination of the pipette tip is often required to ensure that no brains have been mistakenly removed from the tube. If brains have been accidently pipetted into the tip, dispense them back into the microcentrifuge tube, wait for the brains to settle, and then continue removing any extra PTN that remains.</li>
	<li>After the last quick wash, use a p1000 pipette to perform three &#34;long&#34; washes: add 500 &#181;L of PTN and wash for 20 min at room temperature on a rocker/nutator. All future &#34;long&#34; washes should be for 20 mins. 
	NOTE: Following these washes, fixed brains may be stored overnight at 4 &#176;C in PTN.</li>
	<li>Remove the last wash using a p1000 pipette and incubate brains on a rocker or nutator at room temperature in 0.5 mL of blocking solution [PTN + 5% normal goat serum (NGS)] for at least 30 min at room temperature.
	<ol>
		<li>Goat secondary antibodies will be used in subsequent protocol steps. If secondary antibodies from another species will be used, normal serum from that species (rather than NGS) should be used in the blocking and antibody solutions.</li>
	</ol>
	</li>
	<li>Using a p1000 pipette, remove blocking solution and add the primary antibody diluted in PTN (PTN + 5% NGS + diluted primary antibody). When using a primary antibody for the first time, the optimal dilution of that antibody should be determined empirically. 
	Note: For visualizing mushroom body neurons, antibodies recognizing Fas2 are typically used. These antibodies are available from the Developmental Studies Hybridoma Bank (DSHB) as antibody 1D4 and should be diluted 1:20 in PTN + 5% NGS. 
	NOTE: For visualizing photoreceptor neurons, antibodies recognizing chaoptin are typically used. Chaoptin antibodies are available from the DSHB as antibody 24B10 and should be diluted 1:20 in PTN + 5% NGS. 
	NOTE: The fixation process typically eliminates fluorescence from fluorescent protein such as green fluorescent protein (GFP). Therefore, when using MARCM to analyze axonal guidance of individual MB neurons, use an antibody recognizing GFP.</li>
	<li>Incubate brains in primary antibody solution on a rocker/nutator for 2 - 3 nights at 4 &#176;C.</li>
	<li>Following incubation with primary antibodies, allow brains to settle to the bottom of the microcentrifuge tube and then remove the primary antibody solution.</li>
	<li>Using a p1000 pipette, perform 2 &#34;quick&#34; washes and 3 &#34;long&#34; 20 min washes with 0.5 mL of PTN as described above in Steps 4.4 and 4.5, carefully allowing brains to settle by gravity to the bottom of the microcentrifuge tube between each wash.</li>
	<li>Incubate brains for 3 h at room temperature with appropriate fluorescently-labeled secondary antibodies. Secondary antibodies are usually diluted in 0.5 mL of PTN + 5% NGS at a concentration of 1:200. 
	NOTE: Once fluorescent secondary antibodies have been added, brains should be kept in the dark for the remainder of the experiment. 
	NOTE: In case residual GFP fluorescence remains, when performing MARCM analysis it is advisable to use a secondary antibody labeled with a fluorophore having similar excitation/emission wavelengths as GFP (e.g., Fluoroscein isothyocyanate (FITC) or Alexa488).</li>
	<li>Following secondary antibody incubation, allow brains to settle to the bottom of the microcentrifuge tube and remove the secondary antibody solution.</li>
	<li>Perform 2 &#34;quick&#34; washes and 3 &#34;long&#34; 20 min washes with 0.5 mL of PTN as described above in Steps 4.4 and 4.5, carefully allowing brains to settle to the bottom of the microcentrifuge tube between each wash.</li>
	<li>Following the third &#34;long&#34; 20 min wash, use a p200 pipette to remove as much buffer as possible.</li>
	<li>Add 75 &#181;L of fluorescent anti-fade mounting medium to the brains. Pipette brains and mounting medium into the pipette tip once to mix. Do not invert the tube since brains may become stuck on the cap or sides of the microcentrifuge tube. 
	NOTE: Following suspension of brains in mounting medium, tubes may be wrapped in aluminum foil to slow fluorophore quenching and stored at 4 &#176;C overnight. If necessary, brains can be stored for several days at 4 &#176;C, but should ideally be mounted onto slides as soon as possible.</li>
</ol>
5. Mounting Adult D. melanogaster Brains onto Microscope Slides and Imaging
<ol>
	<li>Build a &#34;bridge&#34; slide. Position two &#34;base&#34; coverslips roughly 1 cm apart on a positively charged slide. Ensure that the positively charged side of the slide is facing up. Adhere the coverslips to the slide with fingernail polish as shown in Figure 3A. It is usually helpful to seal the three outer edges of each base cover slip with fingernail polish to ensure mounting media does not wick under these base cover slips. Let fingernail polish dry completely (10 - 15 min) before proceeding.</li>
	<li>Place the slide under the stereomicroscope and pipette the mounting media solution containing the dissected brains into the space between the two coverslips. To provide more contrast, it is useful to maneuver the gooseneck lights so that they are parallel with the bench top.</li>
	<li>Remove extra mounting media from the slide using a pipette, being careful not to pipette the brains off the slide.</li>
	<li>Wick away extra mounting media. This will allow the brains to be positioned more precisely during the next step.</li>
	<li>Using a pair of forceps and the stereomicroscope, position the brains on the slide in a grid pattern with antennal lobes facing up.</li>
	<li>Place a cover slip (the &#34;bridge&#34;) over the brains (Figure 3A). Use fingernail polish to seal the sides of the bridge cover slip where they contact the &#34;base&#34; cover slips.</li>
	<li>Using a p200 pipette, slowly fill the center cavity under the bridge with fresh mounting media (Figure 3B). Place one drop at a time on the open edge of the center coverslip and allow the mounting media to wick under the center bridge coverslip. Continue until the entire cavity is filled with mounting media, then seal the top and bottom with clear fingernail polish.</li>
	<li>Once the fingernail polish is dry, image the slides immediately or store in a lightproof tight slide box at -20 &#176;C.</li>
	<li>Within one week, image brains using a laser-scanning confocal microscope with excitation lasers and filter cubes appropriate to the chosen fluorescent secondary antibodies. Z-stack images of mushroom body neurons are typically obtained using 20X&#160;or 40X&#160;objectives. Imaging of retinal photoreceptor neurons may require higher magnification.</li>
</ol>

The authors declare that they have no competing financial interests.

The dissection and visualization method described above can be used in a wide variety of immunostaining and live imaging applications. We have outlined a general immunostaining protocol and have highlighted one way in which MARCM can be used to visualize axonal morphology of individual mushroom body neurons. In addition, these general procedures can also be used for imaging other brain regions in fixed or freshly dissected adult brains48,65. Live imaging may provide a more efficient approach when analyzing expression patterns of enhancer traps, reporter genes, or MiMIC lines66, for example. To prevent capturing artifacts from cell death during live imaging of GFP in unfixed tissue, brains should be dissected in 1x Phosphate Buffered Saline (PBS) or HL3 media48, mounted on bridge slides in 1x&#160;PBS (or HL3), and imaged in less than 15 - 20 min. Care should also be taken to remove as much trachea as possible from the dissected brain prior to imaging, since it can interfere with visualization of GFP fluorescence.
While staining conditions for assessing the morphology of mushroom body and photoreceptor neurons are relatively well-established24,61,67, the localization of a specific protein of interest in these cell types may require extensive troubleshooting and optimization. In particular, several critical steps, including antibody dilution, blocking buffer component concentration, and fixation, should be optimized to generate the most reproducible and reliable results. First, the optimal concentration of primary antibody should be determined. Although commercially available antibodies often have a suggested dilution (and we often initially start by using that concentration), these values are rarely determined empirically on Drosophila tissues. Therefore, testing a series of primary antibody dilutions that range above and below the manufacturer&#39;s starting suggestion often results in more specific staining. While primary antibody dilution can have a significant effect on staining accuracy and should be precisely determined, the time brains are incubated in primary antibody containing solutions can vary considerably with little effect on the overall result. For example, we have observed similar results when incubating dissected brains with the Fas2 antibody primary antibody for two, three, or even four days at 4 &#176;C. Although we recommend at least two nights, we have also incubated brains in primary antibody solution for a single night and obtained reproducible staining. Importantly, limiting the amount of time brains are incubated in the secondary antibody to 3 h at room temperature (or one night at 4 &#176;C) helps limit non-specific binding of secondary antibodies to brain tissue.
Second, it may be necessary to use additional &#34;blocking&#34; reagents to eliminate unwanted background signal. Options include increasing the concentration of NGS to ~10%, adding 1 - 10% Bovine Serum Albumin (BSA) to the blocking and primary/secondary antibody solutions, or pre-adsorption of primary antibodies overnight at 4 &#176;C with Drosophila embryos (see reference68, section 2.9, step 3).
While we have written the above protocol using PTN as the suggested buffer for all fixation, wash, and antibody incubation steps, tissue fixation in other buffers (such as PLP and PEM, listed in the Materials Table) may result in profound differences in signal strength. For example, previous studies have demonstrated that Cyclin E is undetectable in larval imaginal discs when tissues are fixed in traditional 4% paraformaldehyde diluted in PBS, but is clearly visible when tissues are fixed in PLP buffer (Ken Moberg, personal communication, and reference69). In addition to changes in buffer components, altering the time and temperature for tissue fixation can also significantly affect immunofluorescent staining and can be empirically determined if needed. Generally, fixation time should be long enough to allow for sufficient crosslinking of cellular components and long-term maintenance of overall cellular morphology while being limited enough to prevent over-crosslinking and &#34;burying&#34; of protein epitopes. Therefore, when initially optimizing staining conditions for a newly acquired primary antibody, we usually limit fixation time to approximately 20 min and will often fix tissues at colder temperatures.
Several controls should be included in any immunofluorescence experiment to investigate the specificity of primary and secondary antibodies as well as the effect of transgenes/genetic background on the observed phenotype. To ascertain whether the primary antibody being used specifically recognizes the protein of interest, tissue from flies lacking this protein and/or tissue overexpressing the protein should be included as controls. The addition of excess purified antigen protein can also be included to determine whether the primary antibody recognizes other epitopes in the fly brain. Finally, any fluorescent signal present when primary antibodies are omitted represents the level of non-specific binding by the selected secondary antibodies.
Several important controls should also be included to assess the contribution of genetic background or the presence of transgenes to staining intensity or neuronal morphology. For example, the flies used in the MARCM experiment in Figure 5 contain multiple transgenes (hsFLP, UAS-CD8-GFP, FRT82B, Tub&#62;GAL80, and OK107-GAL4), each of which should be analyzed separately for effects on mushroom body morphology. At a very minimum, mushroom body morphology of flies containing both OK107-GAL4 and UAS-CD8-GFP should be analyzed. It may also be necessary to assess the effect of heat shock and Flp recombinase production on mushroom body development by analyzing flies containing both hsFLP and FRT82B. Although MARCM can provide significant insight into whether a given protein autonomously controls axonal guidance, the number of controls required to accurately make this conclusion can be a minor limitation of this technique. In less complicated experiments, similar controls are also applicable. For example, take an experiment where the GAL4/UAS system is being used in combination with an RNAi transgene to knockdown expression of a protein of interest in all neurons. In this experiment, at least two transgenes will be present: a pan-neuronal GAL4 driver, such as elav-GAL4, and the UAS-RNAi transgene. The morphology of mushroom body neurons in flies containing each of these transgenes alone should be investigated in addition to the experimental condition where flies harbor both transgenes in the same fly.
Finally, to determine whether the protein of interest is expressed in mushroom body neurons it is usually necessary to co-stain brains with antibodies to Fas2. Alternatively, fluorescent proteins such as GFP, RFP, or the membrane bound CD8-GFP can also be expressed using the GAL4/UAS system and used in combination with antibodies recognizing the protein of interest. Since Fas2 only recognizes the fasciculated axons that form the mushroom body &#945; and &#946; lobes, the use of GAL4-driven, membrane-bound CD8-GFP has been particularly useful for marking mushroom body neurons.
Defects in axonal guidance are often not 100% penetrant and even brains of the same genotype may show some variability. Therefore, when analyzing newly generated mutant alleles for defects in mushroom body or photoreceptor pathfinding, a combination of different alleles and approaches should be utilized. Most studies in the literature use several different approaches to investigate whether a protein plays a cell-autonomous role in controlling axonal guidance: i) analysis of pathfinding defects in homozygous null flies (preferably using different null alleles), ii) analysis of pathfinding defects in flies lacking the protein of interest only in neurons (usually via RNAi), iii) MARCM analysis of pathfinding defects, and iv) rescue experiments where the gene is re-expressed in the neurons of homozygous null flies. Ideally, several dozen brains per genotype should be analyzed for defects in neuronal morphology.
Although the protocol described here primarily focuses on immunofluorescent localization of proteins within fixed tissue, several future applications of this technique are being developed to image live brain tissue. Once brains are dissected (usually in cell culture media), they can then be cultured for several days at 25 &#176;C. These ex vivo culturing methods are being developed in order to investigate a wide variety of biological processes, such as protein activity that promotes axon regeneration following injury70,71, intracellular signaling dynamics72, and neuronal development73.

Although the Drosophila nervous system is smaller than that of humans and rodents, its complexity provides a powerful, approachable model to better understand its vertebrate counterparts. In many cases, the genomes of vertebrates and flies encode very similar proteins that dictate the mechanisms of nervous system development. In fact, many of the genes required for neuronal development in vertebrates have orthologs in flies, including those involved in signaling pathways that control patterning, neurogenesis, and axonal guidance1,2,3,4,5. For example, netrin is a ligand required for axonal guidance in mammals and D. melanogaster6,7,8. While netrin was originally isolated from embryonic chick brain tissue6, subsequent studies have revealed that netrin plays a conserved role during development of the embryonic central nervous system (CNS) in Drosophila8. Other studies have used genetic screens in the embryonic Drosophila CNS to identify conserved ligands and receptors required for pathfinding in both Drosophila and vertebrates9.
While the embryonic fly CNS has been used extensively in the past to identify ligands, receptors, and intracellular signaling proteins required for axonal guidance8,9, recent work has investigated the ways in which many of these proteins also control pathfinding decisions during later stages of development. Specifically, investigation of mushroom body (MB) and retinal photoreceptor neuron development (Figure 1) has provided insight into the mechanisms that control pathfinding, synapse formation, axon pruning, and several other aspects of neuronal development10,11,12,13,14,15,16,17. Photoreceptor neurons connect the fly retina to regions of the adult brain called the lamina and medulla and are critical for relaying visual information to the brain (reviewed by18,19), while mushroom body neurons are centrally located in the fly brain and are required for learning and memory20,21. Both photoreceptor neurons and the intrinsic neurons of the mushroom bodies, called Kenyon cells, utilize evolutionarily conserved diffusible and contact-dependent axonal guidance mechanisms to find their post-synaptic targets. In addition to being visible in adult flies, photoreceptor and MB neurons can also be directly visualized in larvae and pupae with antibodies or reporter genes22,23,24,25. The ability to easily visualize these two sets of neurons at different developmental time points has promoted their use as superb models for many aspects of neuronal development.
In addition to being used as a model to understand the mechanisms of normal nervous system development, recent studies have demonstrated that flies can also serve as accurate models of a wide variety of human diseases, including Fragile X Syndrome (FXS)26, Intellectual Disability (ID)27,28,29,30,31, and others32. For example, to study the molecular function of ZC3H14, a gene recently linked to human intellectual disability, we created a fly model of ID using a null allele of the fly ZC3H14 ortholog, called Nab230. Flies lacking Nab2 have severe memory impairments and extended poly(A) tails, recapitulating what is observed in human patients or patient derived cell lines33,34. Importantly, flies lacking Nab2 also display severe brain morphology defects in their adult mushroom bodies34, similar to what is observed in flies lacking the FXS syndrome gene, FMR135. Thus, flies can serve as an important model organism to study both normal brain development and diseases that disrupt it.
Finally, the accessibility of high throughput methods to monitor behavior, combined with the vast array of available genetic tools, make Drosophila a model organism of choice for identifying the brain regions that control complex behaviors, such as learning and memory, sleep, courtship, thirst, and others36,37,38,39. One particularly useful tool that is at the center of a fly geneticist&#39;s &#34;toolbox&#34; is the GAL4/UAS system (Figure 2). This system40,41 uses tissue specific expression of the Gal4 transcriptional activator to increase the expression of genes or transgenes downstream of an Upstream Activating Sequence (UAS). Modifications of this system have allowed researchers to, for example, precisely control the excitability of specific neurons42,43, overexpress or knock-down specific genes of interest44,45, analyze real-time calcium dynamics in vivo46, and express reporter genes to mark neuronal lineages41. The combination of the GAL4/UAS system with mitotic recombination also allowed for the creation of the Mosaic Analysis with a Repressible Cell Marker (MARCM) system12,47. MARCM has been used extensively in single neuron tracing to identify the cellular signaling components required for axonal guidance12,47. Although these and other techniques have provided a number of valuable insights on the cellular mechanisms required for nervous system function, most require that the Drosophila brain first be dissected; careful removal of the brain is required to retain correct brain morphology and connection patterns between brain regions. The following protocol uses mushroom body and photoreceptor neurons as example neuronal populations as it guides you through the dissection and immunofluorescent staining of adult Drosophila brains.

<table>
	<tbody>
		<tr>
			<td>Microdissection forceps/tweezers</td>
			<td>Ted Pella</td>
			<td>505-NM</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard dishes</td>
			<td>Living Systems Instrumentation</td>
			<td>DD-50-S-BLK</td>
			<td>Available from amazon.com</td>
		</tr>
		<tr>
			<td>Fas2 Antibody</td>
			<td>Developmental Studies Hybridoma Bank</td>
			<td>1D4</td>
			<td></td>
		</tr>
		<tr>
			<td>Chaoptin Antibody</td>
			<td>Developmental Studies Hybridoma Bank</td>
			<td>24B10</td>
			<td></td>
		</tr>
		<tr>
			<td>GFP Antibody</td>
			<td>Aves Lab</td>
			<td>GFP-1010</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa488 goat anti-mouse secondary antibody</td>
			<td>ThermoFisher</td>
			<td>A-11001</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa488 goat anti-chicken secondary antibody</td>
			<td>ThermoFisher</td>
			<td>A-11039</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa647 goat anti-mouse secondary antibody</td>
			<td>ThermoFisher</td>
			<td>A-21236</td>
			<td></td>
		</tr>
		<tr>
			<td>20% paraformaldehyde</td>
			<td>Electron Microscope Services</td>
			<td>RT15713</td>
			<td></td>
		</tr>
		<tr>
			<td>VectaShield</td>
			<td>Vector Labs</td>
			<td>H-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>SuperFrost Plus Slides</td>
			<td>ThermoFisher</td>
			<td>99-910-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>ThermoFisher</td>
			<td>12-553-454</td>
			<td></td>
		</tr>
		<tr>
			<td>Na Phosphate Buffer monobasic</td>
			<td>Sigma</td>
			<td>S3139</td>
			<td></td>
		</tr>
		<tr>
			<td>Na phosphate Buffer dibasic</td>
			<td>Sigma</td>
			<td>S3264</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X 100</td>
			<td>Sigma</td>
			<td>X100-100ml</td>
			<td></td>
		</tr>
		<tr>
			<td>fingernail polish</td>
			<td>Electron Microscope Services (EMS)</td>
			<td>72180</td>
			<td></td>
		</tr>
		<tr>
			<td>stereomicroscope</td>
			<td>Leica S6D with KL300 LED light source</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>9-well dish (spot plate)</td>
			<td>VWR</td>
			<td>89090-482</td>
			<td></td>
		</tr>
		<tr>
			<td>nutator/rocker</td>
			<td>Fisher</td>
			<td>22-363-152 or 88-861-041</td>
			<td></td>
		</tr>
		<tr>
			<td>35mm dish</td>
			<td>Genesee Scientific</td>
			<td>32-103</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard</td>
			<td>Fisher</td>
			<td>50-366-794</td>
			<td></td>
		</tr>
		<tr>
			<td>Kimwipe</td>
			<td>Fisher</td>
			<td>06-666</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Potential Fixation Buffers</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PTN Buffer</td>
			<td></td>
			<td></td>
			<td>0.1M NaPhosphate, pH 7.2, 0.1% Triton-X-100, Typically make up 0.5 L of 0.1M NaPhosphate buffer and aliquote 50ml at a time as needed</td>
		</tr>
		<tr>
			<td>PLP buffer</td>
			<td></td>
			<td></td>
			<td>2% paraformaldehyde, 0.01M NaI04, 0.075M Lysine, 0.037M NaPO4, pH 7.2, Dissolve 0.36 g lysine in 10 ml H2O + 7.5 ml 0.1 M NaH2PO4 pH 7.2 + 2.5 ml 0.1 M Na2HPO4 on ice. Immediately before use, mix 15 ml of this buffered lysine solution with 50 mg NaIO4 (sodium periodate) + 2ml of the 20% high grade paraformaldehyde (EMS) + 3ml H2O</td>
		</tr>
		<tr>
			<td>PEM buffer</td>
			<td></td>
			<td></td>
			<td>0.1M PIPES pH 7.0, 2mM MgS04, 1mM EGTA, This buffer can be conveniently made as a 2x stock and diluted with 8% paraformaldehyde (PFA) to give a final concentration of 4% PFA</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Fly Stocks available from Bloomington</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>elav (c155)-GAL4</td>
			<td></td>
			<td>BL458</td>
			<td>Pan-neuronal GAL4 driver</td>
		</tr>
		<tr>
			<td>w*;;;OK107-GAL4</td>
			<td></td>
			<td>BL 854</td>
			<td>GAL4 driver for all mushroom body neurons (OK107-GAL4 insertion is on the 4th chromosome)</td>
		</tr>
		<tr>
			<td>y(1), w(67c23); c739-GAL4</td>
			<td></td>
			<td>BL 7362</td>
			<td>GAL4 driver for alpha and beta lobes (on 2nd chromosome)</td>
		</tr>
		<tr>
			<td>y(1), w(67c23); c739-GAL4, UAS-CD8-GFP</td>
			<td></td>
			<td>BL 64305</td>
			<td>GAL4 driver for alpha and beta lobes, also contains UAS-CD8-GFP</td>
		</tr>
		<tr>
			<td>w*; 201Y-GAL4</td>
			<td></td>
			<td>BL 4440</td>
			<td>GAL4 driver for primarily the gamma lobes of mushroom body (on 2nd chromosome)</td>
		</tr>
		<tr>
			<td>y(1), w(67c23); 201Y-GAL4, UAS-CD8-GFP</td>
			<td></td>
			<td>BL 64296</td>
			<td>GAL4 driver for mushroom body gamma lobes, also contains UAS-CD8-GFP</td>
		</tr>
		<tr>
			<td>w*, elav (c155)-GAL4, hsFLP; FRTG13, Tub&#62;Gal80/CyO</td>
			<td></td>
			<td>BL 5145</td>
			<td>MARCM stock, contains FRT site and GAL80 on 2nd chromosome</td>
		</tr>
		<tr>
			<td>w*, elav (c155)-GAL4, hsFLP, UAS-CD8-GFP</td>
			<td></td>
			<td>BL5146</td>
			<td>MARCM stock, contains hsFLP, pan-neuronal GAL4, and CD8-GFP on X chromosome</td>
		</tr>
		<tr>
			<td>y(1), w*, hsFLP, UAS-CD8-GFP;;FRT82B, Tub&#62;GAL80/TM3, Sb(1);OK107-GAL4</td>
			<td></td>
			<td>BL 44408</td>
			<td>MARCM stock for flipping 3rd chromosome</td>
		</tr>
		<tr>
			<td>y(1), w*, hsFLP, UAS-CD8-GFP;FRT40A, Tub&#62;GAL80;OK107-GAL4</td>
			<td></td>
			<td>BL44406</td>
			<td>MARCM stock for flipping 2nd chromosome</td>
		</tr>
		<tr>
			<td>w*, hsFLP, tub&#62;GAL80, FRT19A; UAS-CD8-GFP/CyO;;OK107-GAL4</td>
			<td></td>
			<td>BL 44407</td>
			<td>MARCM stock for flipping X chromosome</td>
		</tr>
		<tr>
			<td>y(1), w*; UAS-CD8-GFP/CyO</td>
			<td></td>
			<td>BL 5137</td>
			<td>GFP labels cell surface (CD8 is a transmembrane protein)</td>
		</tr>
		<tr>
			<td>y(1), w*; FRTG13, UAS-CD8-GFP</td>
			<td></td>
			<td>BL 5139</td>
			<td>MARCM stock, contains FRT site and CD8-GFP on 2nd chromosome</td>
		</tr>
		<tr>
			<td>y(1), w*, hsFLP, UAS-CD8-GFP; Pin(1)/CyO</td>
			<td></td>
			<td>BL 28832</td>
			<td>MARCM stock, contains hsFLP and CD8-GFP on X chromosome</td>
		</tr>
		<tr>
			<td>w*; FRTG13, Tub&#62;GAL80</td>
			<td></td>
			<td>BL 5140</td>
			<td>MARCM stock, contains FRT site and GAL80 on 2nd chromosome</td>
		</tr>
		<tr>
			<td>y(1), w*;; FRT82B, Tub&#62;GAL80</td>
			<td></td>
			<td>BL 5135</td>
			<td>MARCM stock, contains FRT site and GAL80 on 3rd chromosome</td>
		</tr>
	</tbody>
</table>

dissection and immunofluorescent staining of mushroom body and photoreceptor neurons in adult drosophila melanogaster brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Watch this Scientific Journal Video about Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains at JoVE.com

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

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

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

dissection-immunofluorescent-staining-mushroom-body-photoreceptor

Research

JoVE Journal

Developmental Biology

19.1K Views.  The College of Wooster. 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.

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

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila melanogaster, inductive microenvironments known as larval Hematopoietic Pockets (HPs) have been identified as anatomical sites for the development and regulation of blood cells (hemocytes), in particular of the self-renewing macrophage lineage.&#160;HPs are segmentally repeated pockets between the epidermis and muscle layers of the larva, which also comprise sensory neurons of the peripheral nervous system. In the larva, resident (sessile) hemocytes are exposed to anti-apoptotic, adhesive and proliferative cues from these sensory neurons and potentially other components of the HPs, such as the lining muscle and epithelial layers. During normal development, gradual release of resident hemocytes from the HPs fuels the population of circulating hemocytes, which culminates in the release of most of the resident hemocytes at the beginning of metamorphosis. Immune assaults, physical injury or mechanical disturbance trigger the premature release of resident hemocytes into circulation. The switch of larval hemocytes between resident locations and circulation raises the need for a common standard/procedure to selectively isolate and quantify these two populations of blood cells from single Drosophila larvae. Accordingly, this protocol describes an automated method to release and quantify the resident and circulating hemocytes from single larvae. The method facilitates ex vivo approaches, and may be adapted to serve a variety of developmental stages of Drosophila and other invertebrate organisms.

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

Drosophila blood cells, or hemocytes, cycle between resident sites and circulation. In the larva, resident (sessile) hemocytes localize to inductive microenvironments, the Hematopoietic Pockets, while circulating hemocytes move freely in the hemolymph. The goal of this protocol is the standardized isolation and quantification of these two, behaviorally distinct but interchanging, hemocyte populations.

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila ...

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is a model for understanding microbial interactions with animal hosts, facilitated by approaches to rear large sample sizes of Drosophila under microorganism-free (axenic) conditions, or with defined microbial communities (gnotobiotic). This work outlines a method for collection of Drosophila embryos, hypochlorite dechorionation and sterilization, and transfer to sterile diet. Sterilized embryos are transferred to sterile diet in 50 ml centrifuge tubes, and developing larvae and adults remain free of any exogenous microbes until the vials are opened. Alternatively, flies with a defined microbiota can be reared by inoculating sterile diet and embryos with microbial species of interest. We describe the introduction of 4 bacterial species to establish a representative gnotobiotic microbiota in Drosophila. Finally, we describe approaches for confirming bacterial community composition, including testing if axenic Drosophila remain bacteria-free into adulthood.

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

A method for rearing Drosophila melanogaster under axenic and gnotobiotic conditions is presented. Fly embryos are dechorionated in sodium hypochlorite, transferred aseptically to sterile diet, and reared in closed containers. Inoculating diet and embryos with bacteria leads to gnotobiotic associations, and bacterial presence is confirmed by plating whole-body Drosophila homogenates.

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is ...

Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila melanogaster

In multicellular organisms, a small group of cells is endowed with a specialized function in their biogenic activity, inducing a systemic response to&#160;growth and reproduction. In insects, the larval prothoracic gland (PG) and the adult female ovary play essential roles in biosynthesizing the principal steroid hormones called ecdysteroids. These ecdysteroidogenic organs are innervated from the nervous system, through&#160;which the timing of biosynthesis is affected by environmental cues. Here we describe a protocol for visualizing ecdysteroidogenic organs and their interactive organs in larvae and adults of the fruit fly Drosophila melanogaster, which provides a suitable model system for studying steroid hormone biosynthesis and its regulatory mechanism. Skillful dissection allows us to maintain the positions of ecdysteroidogenic organs and their interactive organs including the brain, the ventral nerve cord, and other tissues. Immunostaining with antibodies against ecdysteroidogenic enzymes, along with transgenic fluorescence proteins driven by tissue-specific promoters, are available to label ecdysteroidogenic cells. Moreover, the innervations of the ecdysteroidogenic organs can also be labeled by specific antibodies or a collection of GAL4 drivers in various types of neurons. Therefore, the ecdysteroidogenic organs and their neuronal connections can be&#160;visualized simultaneously&#160;by immunostaining and transgenic techniques. Finally, we describe how to visualize germline stem cells, whose proliferation and maintenance are controlled by ecdysteroids. This method contributes to comprehensive understanding of steroid hormone biosynthesis and its neuronal regulatory mechanism.

Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila melanogaster

We describe a protocol for dissection, fixation, and immunostaining of steroidogenic organs in Drosophila larvae and adult females to study steroid hormone biosynthesis and its regulatory mechanism. In addition to steroidogenic organs, we visualize the innervation of steroidogenic organs as well as steroidogenic target cells such as germline stem cells.

In multicellular organisms, a small group of cells is endowed with a specialized function in their biogenic activity, inducing a systemic response ...

Dissection and Culture of Mouse Embryonic Kidney

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney development, the two progenitor tissues, the ureteric bud and the metanephric mesenchyme, communicate and reciprocally induce cellular mechanisms to eventually form the collecting system and the nephrons of the kidney. As mammalian embryos grow intrauterine and therefore are inaccessible to the observer, an organ culture has been developed. With this method, it is possible to study epithelial-mesenchymal interactions and cellular behavior during kidney organogenesis. Furthermore, the origin of congenital kidney and urogenital tract malformations can be investigated. After careful dissection, the metanephric rudiments are transferred onto a filter that floats on culture medium and can be kept in a cell culture incubator for several days. However, one must be aware that the conditions are artificial and could influence the metabolism in the tissue. Also, the penetration of test substances could be limited due to the extracellular matrix and basal membrane present in the explant.
One main advantage of organ culture is that the experimenter can gain direct access to the organ. This technology is cheap, simple, and allows a large number of modifications, such as the addition of biologically active substances, the study of genetic variants, and the application of advanced imaging techniques.

This protocol describes a method for isolating and culturing metanephric rudiments from mouse embryos.

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney ...

Dissection and Staining of Drosophila Pupal Ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

Dissection and Staining of Drosophila Pupal Ovaries

The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a reliable protocol for the dissection of the delicate Drosophila pupal retina. Our surgical approach utilizes readily-available microdissection tools to open pupae and precisely extract eye-brain complexes. These can be fixed, subjected to immunohistochemistry, and retinas then mounted onto microscope slides and imaged if the goal is to detect cellular or subcellular structures. Alternatively, unfixed retinas can be isolated from brain tissue, lysed in appropriate buffers and utilized for protein gel electrophoresis or mRNA extraction (to assess protein or gene expression, respectively). Significant practice and patience may be required to master the microdissection protocol described, but once mastered, the protocol enables relatively quick isolation of mainly undamaged retinas.

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

This paper presents a surgical method for dissecting Drosophila pupal retinas along with protocols for the processing of tissue for immunohistochemistry, western analysis, and RNA-extraction.&#160;

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a ...

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the number of novel cell lines is expected to increase. The DGRC aims to familiarize researchers with using Drosophila cell lines as an experimental tool to complement and drive their research agenda. Procedures for working with a variety of Drosophila cell lines with distinct characteristics are provided, including protocols for thawing, culturing, and cryopreserving cell lines. Importantly, this publication demonstrates the best practices required to work with Drosophila cell lines to minimize the risk of contaminations from adventitious microorganisms or from other cell lines. Researchers who become familiar with these procedures will be able to delve into the many applications that use Drosophila cultured cells including biochemistry, cell biology and functional genomics.

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

Drosophila cell lines are important reagents for both fundamental and biomedical research. This article provides protocols for thawing, subculturing, and the cryopreservation of commonly used Drosophila cell lines to assist researchers in incorporating the use of these reagents in their research.

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the ...

Methods to Test Endocrine Disruption in Drosophila melanogaster

In recent years there has been growing evidence that all organisms and the environment are exposed to hormone-like chemicals, known as endocrine disruptor chemicals (EDCs). These chemicals may alter the normal balance of endocrine systems and lead to adverse effects, as well as an increasing number of hormonal disorders in the human population or disturbed growth and reduced reproduction in the wildlife species. For some EDCs, there are documented health effects and restrictions on their use. However, for most of them, there is still no scientific evidence in this sense. In order to verify potential endocrine effects of a chemical in the full organism, we need to test it in appropriate model systems, as well as in the fruit fly, Drosophila melanogaster. Here we report detailed in vivo protocols to study endocrine disruption in Drosophila, addressing EDC effects on the fecundity/fertility, developmental timing, and lifespan of the fly. In the last few years, we used these Drosophila life traits to investigate the effects of exposure to 17-&#945;-ethinylestradiol (EE2), bisphenol A (BPA), and bisphenol AF (BPA F). Altogether, these assays covered all Drosophila life stages and made it possible to evaluate endocrine disruption in all hormone-mediated processes. Fecundity/fertility and developmental timing assays were useful to measure the EDC impact on the fly reproductive performance and on developmental stages, respectively. Finally, the lifespan assay involved chronic EDC exposures to adults and measured their survivorship. However, these life traits can also be influenced by several experimental factors that had to be carefully controlled. So, in this work, we suggest a series of procedures we have optimized for the right outcome of these assays. These methods allow scientists to establish endocrine disruption for any EDC or for a mixture of different EDCs in Drosophila, although to identify the endocrine mechanism responsible for the effect, further essays could be needed.

Methods to Test Endocrine Disruption in Drosophila melanogaster

Endocrine disruptor chemicals (EDCs) represent a serious problem for organisms and for natural environments. Drosophila melanogaster represents an ideal model to study EDC effects in vivo. Here, we present methods to investigate endocrine disruption in Drosophila, addressing EDC effects on fecundity, fertility, developmental timing, and lifespan of the fly.

In recent years there has been growing evidence that all organisms and the environment are exposed to hormone-like chemicals, known as endocrine disruptor ...

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous system and protects neural tissue from toxins and pathogens. Defects in the formation of this barrier have been correlated with neuropathies, and the breakdown of this barrier has been observed in many neurodegenerative diseases. Therefore, it is critical to identify the genes that regulate the formation and maintenance of the blood-brain barrier to identify potential therapeutic targets. In order to understand the exact roles these genes play in neural development, it is necessary to assay the effects of altered gene expression on the integrity of the blood-brain barrier. Many of the molecules that function in the establishment of the blood-brain barrier have been found to be conserved across eukaryotic species, including the fruit fly, Drosophila melanogaster. Fruit flies have proven to be an excellent model system for examining the molecular mechanisms regulating nervous system development and function. This protocol describes a step-by-step procedure to assay for blood-brain barrier integrity during the embryonic and larval stages of D. melanogaster development.

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Blood-brain barrier integrity is critical for nervous system function. In Drosophila melanogaster, the blood-brain barrier is formed by glial cells during late embryogenesis. This protocol describes methods to assay for blood-brain barrier formation and maintenance in D. melanogaster embryos and third instar larvae.

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous ...

Dissection, Immunohistochemistry and Mounting of Larval and Adult Drosophila Brains for Optic Lobe Visualization

The Drosophila optic lobe, comprised of four neuropils: the lamina, medulla, lobula and lobula plate, is an excellent model system for exploring the developmental mechanisms that generate neural diversity and drive circuit assembly. Given its complex three-dimensional organization, analysis of the optic lobe requires that one understand how its adult neuropils and larval progenitors are positioned relative to each other and the central brain. Here, we describe a protocol for the dissection, immunostaining and mounting of larval and adult brains for optic lobe imaging. Special emphasis is placed on the relationship between mounting orientation and the spatial organization of the optic lobe. We describe three mounting strategies in the larva (anterior, posterior and lateral) and three in the adult (anterior, posterior and horizontal), each of which provide an ideal imaging angle for a distinct optic lobe structure.

Dissection, Immunohistochemistry and Mounting of Larval and Adult Drosophila Brains for Optic Lobe Visualization

This protocol describes three steps to prepare larval and adult Drosophila optic lobes for imaging: 1) brain dissections, 2) immunohistochemistry and 3) mounting. Emphasis is placed on step 3, as distinct mounting orientations are required to visualize specific optic lobe structures.&#160;

The Drosophila optic lobe, comprised of four neuropils: the lamina, medulla, lobula and lobula plate, is an excellent model system for exploring the ...