We would like to thank Claude Desplan for sharing with us an aliquot of the Bsh antibody. The DE-Cadherin, Dachshund, Eyes Absent, Seven-up and Bruchpilot monoclonal antibodies were obtained from the Developmental Studies Hybridoma Bank, created by the NICHD of the NIH and maintained at The University of Iowa, Department of Biology, Iowa City, IA 52242. This work was supported by an NSERC Discovery Grant awarded to T.E.. U.A. is supported by an NSERC Alexander Graham Bell Canada Graduate Scholarship. P.V. is supported by an Ontario Graduate Scholarship.

1. Preparing larval brains for confocal imaging
<ol>
	<li>Dissections 
	NOTE: Before starting the dissection, prepare the fix (4% formaldehyde in phosphate buffered saline (PBS)) and PBT (0.1-0.3% Triton in PBS) solutions. The fix solution should be placed on ice during the dissection. Although paraformaldehyde (PFA) fixative is used in this protocol, alternative fixing strategies (using PLP20 or PEM21) have been described for specific epitopes. For larval dissections, two pairs of forceps (Dumont #5 or #55s) are needed. See the Table of Materials for more details.
	<ol>
		<li>Start by filling each well of the dissecting dish with 400 &#181;L of 1x PBS. Use a pair of forceps to gently pick wandering third-instar larvae crawling on the inside of the vial. Place larvae in the first well of the PBS-filled dish.</li>
		<li>Take one larva and transfer it to the middle well. Using the non-dominant hand, hold the body of the larva down at the base of the well.</li>
		<li>With the dominant hand, gently grasp the larval mouth hook and pull away from the body. The larval brain should be attached to the mouth hook and other accessory tissue and will come off as the tissue is pulled away.</li>
		<li>Remove accessory imaginal discs and transfer the larval brain to the third well of the dissection dish. The imaginal discs are epithelial tissues surrounding the larval brain, which generate adult structures such as the antennae, eyes, legs and wings20. If left on the tissue, the imaginal discs may obstruct proper immunostaining of the larval brain.
		<ol>
			<li>To remove the imaginal discs, use a pair of forceps to firmly grip the brain via the ventral nerve cord. Using another pair of forceps, gently pluck away the discs.</li>
			<li>Remove the eye disc carefully, since it wraps over the surface of the brain lobes. To remove the eye disc, use a pair of forceps to hold the brain against the base of the dish. Instead of holding the brain via the ventral nerve cord, use the forceps to lightly grasp the brain lobe being careful to not squeeze the lobe. Using another pair of forceps, gently pull away the eye disc. 
			NOTE: The eye imaginal disc will eventually give rise to photoreceptors of the retina. For those interested in studying neural connectivity between the retina and optic lobe, the eye disc can be left on the brain. However, for those solely interested in optic lobe structures, it is recommended to remove the eye disc as it may hinder image quality due to its location on the surface of the brain.</li>
		</ol>
		</li>
		<li>Repeat steps 1.1.2&#8210;1.1.4 until a sufficient number of brains have been collected (or 30 min of dissection time have elapsed). Dissections should not go longer than 30 min to ensure the integrity of the tissue is not compromised.</li>
		<li>Once the dissection period is over, use a P200 pipette to carefully remove PBS from the third well. Ensure that a small volume of liquid remains in the well to keep the brains immersed. 
		NOTE: It is crucial to keep the brains immersed in liquid at all points of the protocol. If the tissue dries out, the quality of the stain may be compromised by unwanted autofluorescence in the confocal image.</li>
		<li>Add 500 &#181;L of the cold fix solution into the third well. Use a pair of forceps to stir the liquid gently, allowing the brains to swirl in the dish. Cover the dish with a glass slide and place it on ice for 30 min to allow for tissue fixation.</li>
		<li>Remove the fixation solution and wash the brains with 400 &#181;L PBT, 5 times. 
		NOTE: Triton is a detergent in PBT that keeps the brains from sticking to each other or the dish, and prepares the tissue for optimal antibody penetration.</li>
	</ol>
	</li>
	<li>Immunohistochemistry 
	NOTE: There are several primary antibodies available from the Developmental Studies Hybridoma Bank (DSHB) that can be used to label specific optic lobe structures or cell types. DE-Cadherin labels the IPC and OPC neuroepithelia, Bruchpilot (Brp) marks the developing neuropil and Dachshund labels lamina and lobula neurons (as well as a small subset of medulla neurons). Additionally, Elav can be used to label neurons, Prospero to mark ganglion mother cells and Repo to identify glia.
	<ol>
		<li>Prepare the primary antibody solution by adding antibodies to PBT upto a total volume of 100 &#181;L. A blocking agent (10% normal goat serum) can be used to prevent non-specific binding between the primary antibody and tissue20,22,23. The antibodies used in this protocol specifically label brain tissue without the need for blocking agents.</li>
		<li>Remove the final post-fixation wash and add the primary antibody solution. Ensure that there is only a small amount of PBT in the well prior to adding the antibody solution. After adding the solution, gently stir the brains with forceps 5&#8210;10 times. Cover with a glass slide, seal with laboratory film and incubate at 4 &#176;C overnight.
		<ol>
			<li>Alternatively, if a refrigerated orbital shaker is available, place the dish on the shaker to optimize the overnight incubation. 
			NOTE: The primary antibody incubation and all subsequent steps can alternatively be performed in a 1.5 mL microcentrifuge tube. The volume of primary and secondary incubations can remain 100 &#181;L, but the wash steps should be performed with 800 &#181;L or more of PBT.</li>
		</ol>
		</li>
		<li>After incubation, wash out the primary antibodies with PBT as in step 1.1.8 of the previous section. 
		NOTE: The primary antibody solution should be saved and stored at 4 &#176;C as it can be reused for subsequent experiments. Many primary antibodies can be reused up to four times.</li>
		<li>Add 400 &#181;L of PBT to the brains for a final wash. Cover the dish with a slide, seal with laboratory film and place it on an orbital shaker at low speed (100 rpm) and room temperature (RT) for ~4 h. 
		NOTE: The protocol can be paused here. Brains can be left in wash for 2&#8210;3 days at 4 &#176;C.</li>
		<li>Prepare the secondary antibody solution in a total volume of 100 &#181;L. 
		NOTE: In this protocol, all secondary antibodies are used at a dilution of 1:500, without blocking agents.</li>
		<li>Remove the PBT wash from the well and add the secondary antibody solution. Mix the tissue in the solution using a pair of forceps. Cover the dish with a slide and laboratory film. Place the dish on an orbital shaker at RT for a minimum incubation period of 2 h. Since secondary antibodies contain light-sensitive fluorophores, cover the dish with aluminum foil. 
		NOTE: The protocol can be paused here. Brains can be left in secondary antibody solution overnight at 4 &#176;C.</li>
		<li>Wash out the secondary antibodies as described in step 1.2.3. Add 400 &#181;L of PBT to the brains for a final wash. Cover the dish with a slide, seal with laboratory film and foil. Place the brains on a shaker at RT for 4 h. 
		NOTE: The protocol can be paused here. Brains can be left in wash for 2&#8210;3 days at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Mounting 
	<ol>
		<li>Remove final PBT wash and replace with two drops of fluorescence-safe mounting media. Place a drop of the mounting media onto the center of a slide.</li>
		<li>Transfer brains from the well to the drop on the slide using forceps. To avoid damaging the optic lobes, the brains can be held by the ventral nerve cord while transferring to the slide.</li>
		<li>Mount the brains as per the mounting strategies that follow (Figure 1A).
		<ol>
			<li>Anterior side up 
			NOTE: For those interested in visualizing the anterior medulla, lamina or lobula plug, this mounting orientation is ideal. The best way to distinguish the anterior versus posterior mounts is by examining the position of the ventral nerve cord relative to the brain lobes.
			<ol>
				<li>Use the anterior orientation, to observe the ventral nerve cord projecting out over the lobes (Figure 1B).</li>
			</ol>
			</li>
			<li>Posterior side up 
			NOTE: This orientation is recommended for those interested in visualizing the posterior tips of the OPC (pOPC) or IPC 10,11.
			<ol>
				<li>Use the posterior orientation, to view the ventral nerve cord projecting out from under the brain lobes (Figure 1C).</li>
			</ol>
			</li>
			<li>Lateral view 
			NOTE: A lateral mounting orientation is used to visualize the crescent of lamina, medulla or lobula plug neurons in both the dorsal-ventral and anterior-posterior axes in a single plane (Figure 1G).
			<ol>
				<li>For a lateral mount, split the brain lobes from each other to fall flat on their side, with the lamina and lobula plug facing up.</li>
				<li>Using two pairs of sharp forceps, split the ventral nerve cord in half, starting from where the cord attaches to the lobes. Note that the two lobes are already separate from one another, with the ventral nerve cord holding them intact. Thus, split the ventral nerve cord down from the cleavage point between the lobes, to separate them. Then flip each brain lobe and attached ventral nerve cord on their sides with the lateral surfaces facing up. 
				NOTE: For lateral view mount, it is recommended to use a tungsten needle with a fine tip to orient the brain lobe on its side. To clearly visualize the orientation of the lobes and the direction of the ventral nerve cord when mounting, the microscope lighting can be adjusted. If using a gooseneck LED light source, the goosenecks should be positioned parallel to the surface of the slide for clear visibility. Also, the lighting intensity can be increased or decreased to enhance contrast between the brain structures and provide clarity while mounting.</li>
			</ol>
			</li>
		</ol>
		</li>
		<li>Place a small drop of PBS on either side of the mounting media containing the brains. Place one cover slip onto each drop, and a final coverslip over the brains. The right and left edges of the top coverslip should rest on the two other coverslips (Figure 1A). 
		NOTE: Prior to placing a coverslip over the brains, a bridge must be built. Larval brains are approximately 200 &#181;m thick, and thus, to keep the integrity of the tissue, a bridge is created that increases the distance between the cover slip and the surface of the slide.</li>
		<li>Seal the edges of the bridge with nail polish to secure the mounted brains.</li>
		<li>Image the brains using confocal microscopy.</li>
	</ol>
	</li>
</ol>
2. Preparing adult brains for confocal imaging
<ol>
	<li>Dissections 
	NOTE: Adult brain dissections are more challenging than larval brains and require careful handling. It is strongly recommended to use at least one pair of ultra-fine forceps (Dumont #55) for this part of the protocol.
	<ol>
		<li>Anesthetize flies with CO2, using either a needle or flypad, and place them on a laboratory wipe or paper towel over ice (to keep them anesthetized).</li>
		<li>Add 400 &#181;L of PBS to all three wells of the glass dish. Use a pair of forceps to gently grab one adult fly by the wings and place it into the first well of the dish.</li>
		<li>Use a pair of forceps held in the non-dominant hand to hold the thorax of the fly against the base of the well. With the dominant hand, gently pull off the head from the rest of the body.</li>
		<li>Transfer the head into the second well. Often, the head will float in the PBS and can be difficult to handle. Use a pair of forceps to hold it down against the base of the well by the proboscis.</li>
		<li>Peel away a region of the cuticle between the eyes using both forceps. Since the brain sits right below the cuticle, be as gentle as possible to prevent damaging the underlying tissue. Continue to peel cuticle away one piece at a time, until the brain is exposed. Remove any accessory tissue (i.e., retina, trachea, air sacs) that may remain attached to the brain. 
		NOTE: Removal of the retina has been described in previous protocols22,24, but the lamina can also be separated from the rest of the optic lobe. This is useful for those wanting to study the medulla or lobula complex, since the lamina sits at the surface and may obstruct visualization of these other optic lobe neuropils when imaging. To remove the lamina, use a pair of sharp forceps to gently pull at the indentation between the lamina and medulla neuropils. The lamina will slowly begin to peel away from the medulla. Repeat this motion until the entire lamina is removed. During this process it is important to maintain a firm grip of the brain. Use another pair of forceps to hold the brain against the base of the dish by positioning it so that the top and bottom of the central brain fit perfectly between the tips of the forceps. A firm grip around the central brain will prevent unwanted damage to the optic lobe.</li>
		<li>Transfer the clean brain to the third well. Repeat steps 2.1.2&#8210;2.1.6 until a sufficient number of brains are obtained or a maximum dissection period of 30 min has elapsed.</li>
		<li>Remove the PBS in the third well and add 500 &#181;L of fix solution. Ensure that the brains do not dry out here or during any wash steps as this will lead to background fluorescence in the confocal images. Cover the dish with a glass slide and allow the brains to incubate in fix for 20 min at RT.</li>
		<li>After fixation, wash the brains with 400 &#181;L of PBT, 5 times. 
		NOTE: The protocol can be paused here. Brains can be covered with a glass slide and laboratory film and left in wash for 2&#8210;3 days at 4 &#176;C. For researchers new to the system, it may be easier to leave accessory tissue on the brains before and during fixation. This will allow researchers to maximize the number of brain samples obtained in a given dissection period. After the tissue has been fixed and washed, the brains can be carefully cleaned before adding primary antibody solution. Adult brains with accessory tissue tend to float, and therefore risk drying out. As a result, it is recommended to use a pair of forceps to carefully pool all of the brains into the center of the dish upon adding fix solution, and to clean the brains immediately after fixation.</li>
	</ol>
	</li>
	<li>Immunohistochemistry 
	<ol>
		<li>Perform immunohistochemistry as described for larval brains in section 1.2.</li>
		<li>Useful primary antibodies from the DSHB include those that are mentioned in the protocol of larval immunohistochemistry in section 1.2. See the Table of Materials for a complete list of antibodies and corresponding dilutions. 
		NOTE: Extra care must be taken during the wash steps for adult brains, since they may float to the surface of the wash solution and risk drying out on the sides of the well when the solution is removed (which will lead to background fluorescence). A good practice is to hold the pipette in one hand to add or draw away liquid, and a pair of forceps in the other to keep the brains submerged.</li>
	</ol>
	</li>
	<li>Mounting 
	<ol>
		<li>Prior to the mounting step, perform a final wash with PBS (instead of PBT). PBS causes the brains to become slightly sticky, which allows them to remain in the desired orientation during mounting.</li>
		<li>Remove the final wash and replace with three drops of fluorescence-safe mounting media. Place a drop of the mounting media onto the center of a microscope slide.</li>
		<li>Transfer brains from the well to the drop on the slide using forceps. To avoid damaging the optic lobes and prevent the tissue from drying out, carefully retrieve a brain via capillary action. Slowly close a pair of forceps around a brain until a small volume of liquid containing the brain is drawn up between the tips. 
		NOTE: When transferring the brain to the slide, be careful to not close the forceps as this will damage the brain. Alternatively, a P200 pipette can be used to transfer the brains. Cut off the end of the tip to widen the opening and pre-rinse with PBT to prevent the brains from sticking to the inside of the tip.</li>
		<li>Mount the brains as per the following strategies (Figure 2A).
		<ol>
			<li>Anterior side up 
			NOTE: To visualize lamina and medulla neurons, orient the brain with the anterior side up (Figure 2B). This can be achieved by examining the anatomy and curvature of the central antennal lobes.
			<ol>
				<li>With a pair of forceps, gently turn the brain on its side to examine both the anterior and posterior sides. Observe that on the anterior side, the curvature of the brain is pronounced and the antennal lobes protrude slightly outward from the center.</li>
			</ol>
			</li>
			<li>Posterior side up
			<ol>
				<li>Position the brains with the posterior (flat) side facing up and the antennal lobes down (Figure 2C). 
				NOTE: This will bring the lobula and lobula plate closer to the imaging surface and is ideal for those interested in visualizing lobula complex neurons.</li>
			</ol>
			</li>
			<li>Horizontal view 
			NOTE: To visualize all optic lobe neuropils simultaneously, use a horizontal mounting strategy (Figure 2G). In this orientation, neuronal cell bodies, axonal trajectories and dendritic arborizations can be visualized in the same plane.
			<ol>
				<li>Initially, orient the brain with the anterior side up. With a tungsten needle, gently tilt the brain 90&#176; upwards so that it sits on its dorsal side with its antennal lobes facing outwards. Check that both the anterior and posterior sides of the brain are visible in this orientation.</li>
			</ol>
			</li>
		</ol>
		</li>
		<li>Instead of a cover slip bridge, use clay to elevate the cover slip from the slide. The use of clay allows for re-mounting of the brains after imaging (described in the discussion section).
		<ol>
			<li>Place a small piece of clay on each corner of a cover slip. Ensure that each piece of clay is relatively the same thickness. The pieces of clay should be between 0.5&#8211;1 mm in thickness. Gently place the coverslip over the brains and apply slight pressure to the corners. The coverslip should immediately catch the liquid and form a seal. The slide is now ready for imaging. 
			NOTE: For long-term storage of the slide, it is recommended to seal the coverslip with nail polish and store at 4 &#176;C away from light.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+developing+and+adult+Drosophila+brains+and+retinae+for+live+imaging.">Preparation of developing and adult Drosophila brains and retinae for live imaging.</a> Journal of Visualized Experiments. , e1936 (2010).</li><li>Bertet, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+patterning+of+neuroblasts+controls+notch-mediated+cell+survival+through+regulation+of+hid+or+reaper.">Temporal patterning of neuroblasts controls notch-mediated cell survival through regulation of hid or reaper.</a> Cell. 158, 1173-1186 (2014).</li><li>Pinto-Teixeira, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+Concurrent+Retinotopic+Maps+in+the+Fly+Motion+Detection+Circuit.">Development of Concurrent Retinotopic Maps in the Fly Motion Detection Circuit.</a> Cell. 173, 485-498 (2018).</li><li>Plevock, D. A., Rusan, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+Imaging+of+Drosophila+Larval+Neuroblasts.">Live Imaging of Drosophila Larval Neuroblasts.</a> Journal of Visualized Experiments. , e51756 (2014).</li><li>Ting, C. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analyzing+dendritic+morphology+in+columns+and+layers.">Analyzing dendritic morphology in columns and layers.</a> Journal of Visualized Experiments. , e55410 (2017).</li><li>&#214;zel, M. N., Langen, M., Hassan, B. A., Hiesinger, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Filopodial+dynamics+and+growth+cone+stabilization+in+Drosophila+visual+circuit+development.">Filopodial dynamics and growth cone stabilization in Drosophila visual circuit development.</a> eLife. 4, (2015).</li></ol>

The authors declare that they have no competing financial interests.

In this protocol, we describe a method to immunostain larval and adult Drosophila brains and mount them in several orientations. While methods to stain larval and adult brains have been previously described22,23,24,27,28, mounting strategies for the optimal visualization of specific optic lobe structures have received less attention2...

The Drosophila visual system, comprised of the compound eye and underlying optic lobe, has been an excellent model for the study of neural circuit development and function. In recent years, the optic lobe in particular has emerged as a powerful system in which to study neurodevelopmental processes such as neurogenesis and circuit wiring1,2,3,4,5,6,7,8. It is made up of four neuropils: the lamina, medulla, lobula and lobula plate (the latter two comprise the lobula complex)1,2,3,4,5,6. Photoreceptors from the eye, target neurons of the lamina and medulla, which process visual inputs and relay them to the neuropils of the lobula complex1,2,3,4,5,6. Projection neurons in the lobula complex subsequently send visual information to the higher order processing centers in the central brain1,5,9. The complex organization of the optic lobe, necessitated by a need to maintain retinotopy and to process different types of visual stimuli, makes it an attractive system for studying how sophisticated neural circuits are assembled. Notably, the medulla shares striking similarities in both its organization and development with the neuroretina, which has long been a model for vertebrate neural circuit development3,8.
Optic lobe development begins during embryogenesis, with the specification of ~35 ectodermal cells that form the optic placode2,4,5,6,7,8. After larval hatching, the optic placode is subdivided into two distinct primordia: 1) the outer proliferation center (OPC), which generates the neurons of the lamina and outer medulla and 2) the inner proliferation center (IPC), which generates neurons of the inner medulla and lobula complex4,5,6,10. In the late second-instar larva, the neuroepithelial cells of the OPC and IPC begin to transform into neuroblasts that subsequently generate neurons via intermediate ganglion mother cells4,5,11,12. Optic lobe neuroblasts are patterned by spatially and temporally-restricted transcription factors, which act together to generate neural diversity in their progeny11,12,13,14. In the pupa, the circuits of the optic lobe neuropils are assembled via the coordination of several processes, including programmed cell death11,15, neuronal migration12,16, axonal/dendritic targeting10,17, synapse formation18,19 and neuropil rotations10,17.
Here, we describe the methodology by which larval and adult brains are dissected, immunostained and mounted for imaging the optic lobe. 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. Thus, we put special emphasis on how the orientation of mounting relates to the spatial organization of the optic lobe structures. We describe three mounting strategies for larval brains (anterior, posterior and lateral) and three for adult brains (anterior, posterior and horizontal), each of which provide an optimal angle for imaging a specific optic lobe progenitor population or neuropil.

<table border="0" cellpadding="0" cellspacing="0" style="width:1077px;" width="1077">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">10x PBS</td>
			<td style="width:325px;">Bioshop</td>
			<td style="width:164px;">PBS405</td>
			<td style="width:357px;"></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">37% formaldehyde</td>
			<td style="width:325px;">Bioshop</td>
			<td style="width:164px;">FOR201</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Alexa Fluor 488 (goat) secondary</td>
			<td style="width:325px;">Invitrogen</td>
			<td style="width:164px;">A-11055</td>
			<td>use at 1:501</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Alexa Fluor 555 (mouse) secondary</td>
			<td style="width:325px;">Invitrogen</td>
			<td style="width:164px;">A-31570</td>
			<td>use at 1:500</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Alexa Fluor 647 (guinea pig) secondary</td>
			<td style="width:325px;">Invitrogen</td>
			<td style="width:164px;">A-21450</td>
			<td>use at 1:503</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Alexa Fluor 647 (rat) secondary</td>
			<td style="width:325px;">Invitrogen</td>
			<td style="width:164px;">A-21247</td>
			<td>use at 1:502</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Cover slips</td>
			<td style="width:325px;">VWR</td>
			<td style="width:164px;">48366-067</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Dissecting forceps - #5</td>
			<td style="width:325px;">Dumont</td>
			<td style="width:164px;">11251-10</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Dissecting forceps - #55</td>
			<td style="width:325px;">Dumont</td>
			<td style="width:164px;">11295-51</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Dissection Dish</td>
			<td style="width:325px;">Corning</td>
			<td>722085</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Dry wipes</td>
			<td style="width:325px;">Kimbery Clark</td>
			<td style="width:164px;">34155</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Goat anti-Bgal primary antibody</td>
			<td>Biogenesis</td>
			<td></td>
			<td>use at 1:1000</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Guinea pig anti-Bsh primary antibody</td>
			<td style="width:325px;">Gift from Claude Desplan</td>
			<td style="width:164px;"></td>
			<td style="width:357px;">use at 1:500</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Guinea pig anti-Vsx1 primary antibody</td>
			<td style="width:325px;">Erclik et al. 2008</td>
			<td style="width:164px;"></td>
			<td style="width:357px;">use at 1:1000</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Laboratory film</td>
			<td style="width:325px;">Parfilm</td>
			<td style="width:164px;">PM-996</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Microcentrifuge tubes</td>
			<td style="width:325px;">Sarstedt</td>
			<td style="width:164px;">72.706.600</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Microscope slides</td>
			<td style="width:325px;">VWR</td>
			<td style="width:164px;">CA4823-180</td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Mouse anti-dac primary antibody</td>
			<td style="width:325px;">Developmental Studies Hybridoma Bank (DSHB)</td>
			<td style="width:164px;">mabdac2-3</td>
			<td>use at 1:20</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Mouse anti-eya primary antibody</td>
			<td style="width:325px;">DSHB</td>
			<td style="width:164px;">eya10H6</td>
			<td>use at 1:20</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Mouse anti-nc82 primary antibody</td>
			<td style="width:325px;">DSHB</td>
			<td style="width:164px;">nc82</td>
			<td>use at 1:50</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Mouse anti-svp primary antibody</td>
			<td style="width:325px;">DSHB</td>
			<td style="width:164px;">Seven-up 2D3</td>
			<td>use at 1:100</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Polymer Clay</td>
			<td>Any type of clay can be used</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Rabbit anti-GFP</td>
			<td style="width:325px;">Invitrogen</td>
			<td style="width:164px;">A-11122</td>
			<td>use at 1:1000</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Rat anti-DE-Cadherin primary antibody</td>
			<td style="width:325px;">DSHB</td>
			<td style="width:164px;">DCAD2</td>
			<td>use at 1:20</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Slowfade mounting medium</td>
			<td style="width:325px;">Invitrogen</td>
			<td style="width:164px;">S36967</td>
			<td>Vectashield mounting medium ( cat# H-1000) can also be used</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:231px;">Triton-x-100</td>
			<td style="width:325px;">Bioshop</td>
			<td style="width:164px;">TRX506</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Confocal images of larval and adult optic lobes mounted in the orientations described in the protocol are presented in Figure 1 and Figure 2.
Figure 1 shows schematics and representative confocal slices of larval brains positioned in the anterior, posterior and lateral orientations. In the anterior mounting orientation, the OPC epithelium (DE-Cadherin), medulla neuroblasts (deadpan&#62;&#946;gal)...

Watch this Scientific Journal Video about Dissection, Immunohistochemistry and Mounting of Larval and Adult Drosophila Brains for Optic Lobe Visualization at JoVE.com

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;

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 ...

dissection-immunohistochemistry-mounting-larval-adult-drosophila

Research

JoVE Journal

Developmental Biology

3.7K Views.  University of Toronto - Mississauga. 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. 

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

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.

Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. ...

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

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.

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.

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

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 ...

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry (GC-MS)-based Metabolomics

Recent advances in the field of metabolomics have established the fruit fly Drosophila melanogaster as a powerful genetic model for studying animal metabolism. By combining the vast array of Drosophila genetic tools with the ability to survey large swaths of intermediary metabolism, a metabolomics approach can reveal complex interactions between diet, genotype, life-history events, and environmental cues. In addition, metabolomics studies can discover novel enzymatic mechanisms and uncover previously unknown connections between seemingly disparate metabolic pathways. In order to facilitate more widespread use of this technology among the Drosophila community, here we provide a detailed protocol that describes how to prepare Drosophila larval samples for gas chromatography-mass spectrometry (GC-MS)-based metabolomic analysis. Our protocol includes descriptions of larval sample collection, metabolite extraction, chemical derivatization, and GC-MS analysis. Successful completion of this protocol will allow users to measure the relative abundance of small polar metabolites, including amino acids, sugars, and organic acids involved in glycolysis and the TCA cycles.

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry GC-MS-based Metabolomics

This protocol describes how to prepare Drosophila larvae for GC-MS-based metabolomic analysis.

Recent advances in the field of metabolomics have established the fruit fly Drosophila melanogaster as a powerful genetic model for studying animal ...

Visualization of Tangential Cell Migration in the Developing Chick Optic Tectum

Time-lapse imaging is a powerful method to analyze migrating cell behavior. After fluorescent cell labeling, the movement of the labeled cells in culture can be recorded under video microscopy. For analyzing cell migration in the developing brain, slice culture is commonly used to observe cell migration parallel to the slice section, such as radial cell migration. However, limited information can be obtained from the slice culture method to analyze cell migration perpendicular to the slice section, such as tangential cell migration. Here, we present the protocols for time-lapse imaging to visualize tangential cell migration in the developing chick optic tectum. A combination of cell labeling by electroporation in ovo and a subsequent flat-mount culture on the cell culture insert enables detection of migrating cell movement in the horizontal plane. Moreover, our method facilitates detection of both individual cell behavior and the collective action of a group of cells in the long term. This method can potentially be applied to detect the sequential change of the fluorescent-labeled micro-structure, including the axonal elongation in the neural tissue or cell displacement in the non-neural tissue.

We describe the methods for fluorescent labeling of tangentially migrating cells by electroporation, and for time-lapse imaging of the labeled cell movement in a flat-mount culture in order to visualize migrating cell behavior in the developing chick optic tectum.

Time-lapse imaging is a powerful method to analyze migrating cell behavior. After fluorescent cell labeling, the movement of the labeled cells in culture ...

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 ...

Visualization and Analysis of Pharyngeal Arch Arteries using Whole-mount Immunohistochemistry and 3D Reconstruction

Improper formation or remodeling of the pharyngeal arch arteries (PAAs) 3, 4, and 6 contribute to some of the most severe forms of congenital heart disease. To study the formation of PAAs, we developed a protocol using whole-mount immunofluorescence coupled with benzyl alcohol/benzyl benzoate (BABB) tissue clearing, and confocal microscopy. This allows for the visualization of the pharyngeal arch endothelium at a fine cellular resolution as well as the 3D connectivity of the vasculature. Using software, we have established a protocol to quantify the number of endothelial cells (ECs) in PAAs, as well as the number of ECs within the vascular plexus surrounding the PAAs within pharyngeal arches 3, 4, and 6. When applied to the whole embryo, this methodology provides a comprehensive visualization and quantitative analysis of embryonic vasculature.

Here, we describe a protocol to visualize and analyze the pharyngeal arch arteries 3, 4, and 6 of mouse embryos using whole-mount immunofluorescence, tissue clearing, confocal microscopy, and 3D reconstruction.

Improper formation or remodeling of the pharyngeal arch arteries (PAAs) 3, 4, and 6 contribute to some of the most severe forms of congenital heart ...

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

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

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Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue