This work was supported by grants from the Welch Foundation (I-1657) and the NIH (RO1EY18884). P.R.H. is a Eugene McDermott Scholar in Biomedical Research.

1. Introduction
In this video, we will demonstrate how to prepare Drosophila brains and retinae for live imaging and immunohistochemistry with a focus on photoreceptor cells. First, we will show how to prepare for brain dissections. Next, we will demonstrate two basic dissections commonly used for immunohistochemistry that form the basis for the live preparations. These two preparations are the adult brain and eye dissections. Next, we will demonstrate the dissections used for live imaging of the adult and developing brain with emphasis on their usage for live imaging of photoreceptors. Finally, we will describe how we image live tissue and show some examples of live imaging using resonant confocal microscopy.
2. Dissection preparation
<ol>
	<li>For good dissections, you need sharp forceps. Using a sharpening block or very fine sand paper (&gt;1500), gently pass the forceps back and forth on each side until the ends meet at a fine point. We use a standard sharpening stone. We will not cover the sharpening technique here, but note that sharp forceps are essential for live dissections. We sharpen forceps before every dissection.</li>
	<li>For adult brain dissections, place the flies on a CO2 pad to anesthetize them and sort out the desired genotype. For pupal dissections, simply reach into the vial and carefully remove a pupa with your forceps, being careful to avoid breaking the pupal case.</li>
	<li>Moisten a Kimwipe with water. You will use this during the dissection to remove debris from your forceps. Position a dissecting dish on the stage of a stereoscope and fill it with HL3 solution 1. All dissections will be conducted in HL3 to ensure that the tissue remains alive and healthy during the dissection procedure. Also, we use a dissection dish that has a bottom covering of Sylgard to protect forceps during the dissection. </li>
	<li>Now, prepare fixing solution if you plan to perform immunohistochemistry. Place 180&mu;L of HL3 into a 500&mu;L microcentrifuge tube. Add 20&mu;L of 37% formaldehyde to obtain a 3.7% formaldehyde solution.</li>
	<li>Finally, proper hand position is important for good dissections. With good hand position, your forceps will be steady and capable of controlled, subtle movements. First, place the forceps on the side of your thumb. Then bring the index finger straight down such that the tip of the finger rests on top. Now rest the side of the forceps on the side of your middle finger and move to the dissection dish. Make physical contact with the dish by your thumb and middle finger. While resting your thumb and middle finger on the dish, plant your wrist on the microscope stage. This way, you have three points planted firmly on surfaces during the dissection and it is now possible to make very subtle manipulations of the forceps. This technique may feel a bit awkward at first, but with practice, you will soon be a master of brain dissections.</li>
</ol>
3. Adult brain
<ol>
	<li>On the CO2 pad, orient an adult fly ventral side up with the head away from your hand. Grab the thorax just below the head. If done properly, the legs and proboscis will extend. While viewing under a stereoscope, grab the extended proboscis with your forceps to remove the head. Discard the body and submerge the head. Refocus onto the submerged head. The entire dissection will be performed under solution. It is very important to hold onto the head with forceps at all times. Otherwise, the head will float and is difficult to retrieve. If the head moves out of focus during the dissection, simply move it back into the focal plane without adjusting the microscope. This seemingly simple task may be difficult at first, but keeping your head in focus will become easier over time. </li>
	<li>Begin the dissection by first tearing the connective tissue between the proboscis and the eye. Tear through the eye while holding firmly with at least one forceps at all times. Be certain that the toe of the forceps is only just beneath the retina or cuticle, to avoid damaging the brain underneath. Alternating left and right, use your forceps to tear away the retina, working toward the back of the head. Tearing away the retina will increase the chances that the lamina remains attached to the optic lobe throughout this dissection. Continue around the back of the head, tearing cuticle along the way. Tear through the other eye and begin pulling away cuticle. As long as you are grabbing cuticle, you know you are not crushing the brain! With this in mind, continue to pull cuticle and retina away until the brain is revealed.</li>
	<li>Once the brain is visible, you can begin to remove trachea attached to the brain (Fig. 1A). Continue to pull at trachea and cuticle until the brain is isolated. At this point, you should refocus to the bottom of your dish for the remaining steps. Remove the remaining trachea because your brain may otherwise float during overnight antibody washes. For imaging of photoreceptor terminals, it is desirable to retain the lamina, but you need to remove the retina, which contains the cell bodies and strongly autofluorescent pigment. To do this, carefully pull at the bushy and pigmented cellular remains without damaging the lamina. This is a finer skill and takes practice.</li>
	<li>The adult brain can be kept alive for hours or even days using conditions that we will discuss in section 6. For immunohistochemistry, the adult brain is now ready to be fixed in formaldehyde.2 Use a P20 pipetteman set to 5&mu;L to move the adult brain to the fixing solution you've already prepared. Continue dissecting adult brains for 20 minutes, which should yield 4-10 brains. Leave the brains in fix for an additional 40 minutes, however this duration may vary for optimal primary antibody staining. Remove the formaldehyde solution and wash the brains 3 times for 5 minutes each in 400&mu;L of a solution containing PBS and 0.4% TritonX-100, hereafter referred to as PBS-Triton . After completely washing out the fix solution, you may add primary antibody solution.</li>
</ol>
4. Adult eye
<ol>
	<li>Begin this dissection as described for the adult brain. After submerging the head and refocusing the microscope, start by tearing the connective tissue between the proboscis and the eye. Above the proboscis, separate the cuticle from the eye. Now, separate the cuticle from the underside of the eye, working toward the back of the head. With one forceps, grab the center of the back of the head. With the other, grab the eye and pull. Allow the eye to settle to the bottom of the dish (Fig. 1B). The lamina will probably still be attached to the eye and can be used for live imaging of the terminals of completely intact photoreceptor cells at this time. To continue with live imaging of photoreceptor cell bodies, go to section 4.5. The next three sections explain fixed eye immunohistochemistry.</li>
	<li>For immunohistochemistry, move the eye to fixing solution using a P20 pipetteman set to 5&mu;L. Continue dissecting adult eyes for a total of 10 minutes. Leave the eyes in fix for an additional 30 minutes. Remove the fix solution and conduct washes as described in the adult brain dissection. </li>
	<li>If you ultimately wish to image the ommatidial structure of the eye the lamina must be removed now. Return the fixed eyes to your dissecting dish and remove any trachea or fat cells that may still be attached. Next, while holding the outside of the eye with one forceps, remove the lamina with the other. It should come off in one piece, exposing the cellbodies underneath (Fig. 1B). Be careful to grab the lamina only! Otherwise, you risk detaching the photoreceptors from the retina. </li>
	<li>Using a P20 pipetteman, move the eyes to 400uL PBS-Triton in a 500uL tube. Gently rock them at 4 degrees overnight to remove the autofluorescent red pigment from the photoreceptors. The following day, you may discard the wash solution and add primary antibody solution.</li>
	<li>For live imaging of adult eyes we only use pharate adult flies, i.e. late stage pupae shortly before eclosion. At this stage, the photoreceptor cell bodies separate from terminals in the lamina during the dissection.</li>
	<li>Pharate adult pupae have well-defined wings, eyes, and cuticle visible through the pupal case. Select a pupa and remove the anterior portion of the pupal case to expose the head region. Carefully remove the thin inner membrane that additionally encloses the developing fly. Grab the base of the head with your forceps and pull. Discard the rest of the pupa while keeping the head submerged.</li>
	<li>Proceed as demonstrated in the eye dissection for immunohistochemistry. At the pharate adult stage, however, the photoreceptor cell bodies will detach more easily from the lamina, allowing you to image the cell bodies nestled in the retina. Also, the lamina remains attached to the optic lobe, allowing you to image the distal lamina. If you wish to image the proximal lamina, where the photoreceptor terminal cartridges can be seen, then perform the eye dissection on an adult fly where the lamina is more strongly attached to the eye.</li>
</ol>
5. Developing eye disc-brain complexes
<ol>
	<li>The developing eye disc at 20% pupal development (P+20%) has become a favorite for live imaging in our lab because this single layer of relatively large developing cells is easily observable using confocal microscopy.3</li>
	<li>To select a 20% pupa, look for the malpighian tubules, a green spot the appears beneath the pupal case, and avoid pupae that have a "dark body"4. After selecting a P+20% pupa, submerge it in HL3 in your dissecting dish and carefully remove an anterior portion of the pupal sack, being careful not to penetrate the thin inner membrane. Grasp the inner membrane with your forceps and pull apart.</li>
	<li>Carefully search through the released contents to find the developing brain and eye discs which are quite prominent (Fig. 1C). The eye disc-brain complex is easily separated at this stage of metamorphosis. Isolate the brain at the bottom of the dissecting dish. Carefully separate the central brain from the optic lobe. The optic lobe/eye disk will be used in live imaging.</li>
</ol>
6. Live Imaging: At the microscope
<ol>
	<li>If you wish to image for a short period of time in only one or two solutions, you may image your tissue on a glass slide. In this case, prepare the slide by first coating the surface with Sylgard using the manufacturer's instructions. Place 20&mu;L HL3 in the middle of the slide. Transport your dissected tissue in 20 additional &mu;L of HL3 to the slide. The tissue must NEVER be exposed to air. In addition, any stress or strain from pipette handling or solution surface tension must be avoided.</li>
	<li>For live imaging, the tissue must be firmly secured with minimal handling and contact. We securely position the brain using the water-polymerizing surgical glue GluShield applied through a microelectrode, a technique routinely used for embryonic preparations for electrophysiology5. Obtain a glass electrode like those pulled for electrophysiological recordings. Break off the tip and fill the end with GluShield using negative air pressure generated by the mouth. Break off only enough of the electrode that glue may enter. Now use positive pressure to apply a small amount of glue to the Sylgard under a drop of HL3. Now, quickly set the tissue on the glue, maintaining the desired orientation for live imaging. A water-immersion lens can now be used for imaging. If you want to add a membrane permeable dye, you can add it to the bath while imaging. </li>
	<li>We use a Resonant Scanning Confocal Microscope that allows fast imaging with reduced photobleaching.6 For imaging over long periods of time or to exchange solutions completely or multiple times, we use a perfusion chamber (Harvard IC30 confocal imaging chamber) which connects to a peristaltic pump that slowly perfuses culture solution over the live tissue (Fig. 2A). Movement of constant liquid exchange and minimal GluShield contact reduce the flattening of the tissue that has been observed over longer periods of time7. Note that for the imaging of developmental time periods, the eye disc-brain complex needs to remain intact and should have minimal contact with the glue.</li>
	<li>Use a cover slip coated with a drop of Sylgard that is shaved to render it thin and flat. Dispense 20&mu;L HL3 onto the center of the coverslip and glue the live tissue to the Sylgard. Generate a gasket that will allow bubbles to pass through the chamber without exposing the live tissue to air (Fig. 2B). The gasket should be thick enough to avoid crushing the tissue but thin enough to bring the tissue close to the confocal microscope lens; we use 250&mu;m in our setup. Assemble the perfusion chamber, being careful to keep the tissue completely submerged at all times. Begin perfusion initially at a rate close to 100&mu;L per minute using a peristaltic pump. This speed is rather fast depending on the gasket thickness that defines the chamber height. For imaging on the scale of hours, you may need to add 20-hydroxyecdysone and antibiotic, but not antifungal, and perform perfusion at much lower speeds around 10&mu;l per minute. Finally, we bubble oxygen into the culture medium that supplies the perfusion chamber.</li>
</ol>
7. Representative Results:
At the end of our video protocol we show an example for live imaging in developing photoreceptor cells using the preparation presented in Section 5. This preparation takes advantage of Drosophila transgenesis to cell-specifically express fluorescent reporters. In the example shown, two fluorescent reporters are expressed under control of the photoreceptor-specific GMR-Gal4 driver8: the membrane-tagged myrRFP and the endosomal marker 2xFYVE-GFP9. Two-channel 3D datasets with a bounding box of 70x70x17.5&mu;m and a voxel size of 137x137x700nm (512x512x26) were obtained every 1 min. The movie shows a selected plane over time in this 4D dataset that reveals dynamics of intracellular endosomal trafficking and vesicle fusion events. Representative images for fixed tissue preparations of the eye and lamina are shown in Fig. 3.

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Fluorescent Probes Used for Imaging
Here we describe the strengths and limitations of the Drosophila visual system preparation for a selection of fluorescent probes from three classes: (1) Probes that are added to a live tissue preparation exogenously; (2) fluorescent proteins that are genetically encoded for both live and fixed imaging; (3) fluorescent dyes that are added to fixed tissue for immunohistochemistry.
1. Exogenous probes:
Lysotracker Green DND-26 or Red DND-99 (Invitrogen, Inc.) These are commercially available fluorescent membrane-permeable compounds that accumulate in strongly acidified compartments in the cell, most prominently lysosomes, late endosomes, and autophagosomes. Lysotracker at nanomolar concentrations (we typically use 50 nM) penetrates the L3 and P+20% eye disc fully in less than one minute. Since Lysotracker continuously accumulates and exerts an alkalizing effect, we image in less than 5 minutes. In contrast to the eye discs, Lysotracker does not penetrate the brain without disruption of the tissue. However, careful tearing of the outer membrane allows penetration of a few cell diameters in the optic lobe.
Lysosensor DND-189 (Invitrogen, Inc.) In contrast to Lysotracker, Lysosensor does not accumulate in acidified compartments, but exhibits increased fluorescence at acidic pH. Lysosensor has penetration characteristics that are very similar to Lysotracker. We use Lysosensor at a concentration of 1μM.
2. Genetically encoded probes:
For many live imaging experiments, we express genes tagged with various fluorescent molecules like GFP, TagRFP10 or the pH-sensitive pHluorin11 under control of the binary Gal4/UAS expression system12. Gal4-lines that express in developing photoreceptors include GMR-Gal4 (all photoreceptors)8 and mDelta0.5-Gal4 (developing R4 and R7)13. Several subtype-specific Gal4-driver lines exist that use different Rhodopsin promoters14. Particularly useful fluorescent probes in live imaging are photoconvertible proteins. We have successfully used Dendra2, a monomeric green-to-red photoconvertable protein15. Note that Dendra2 is designed to be photoconvertible using a 488nm Argon laser line. However, we are unable to achieve conversion using visible blue light using our set-up in either conventional or resonant confocal scanning mode. In contrast, conversion with 405nm UV laser is efficient.
3. Immunohistochemistry:
For the fixed adult eye, we often choose either rhodamine phalloidin (Invitrogen, Inc.) or anti-Chaoptin (a photoreceptor-specific antibody16) to visualize rhabdomeric structure (Fig 1A). A good way to analyze lamina cartridge structure is to combine anti-chaoptin16, the glial marker anti-ebony17, and anti-sec618(Fig 1B). As shown in Fig. 3, this combination of antibodies distinguishes the major presynaptic (chaoptin) and postsynaptic (sec6) processes as well as epithelial glia (ebony) in the lamina19.
Assembling the Perfusion Chamber
For live imaging, we employ a perfusion chamber that allows slow exchange of solutions without disturbing the tissue preparation. A demonstration of perfusion chamber assembly is included in the video and schematically shown in Figure 2. Assembly of the perfusion chamber is illustrated in Fig. 2A. Begin by laying the gasket over the specimen first, and then continue with assembly as shown. The purpose of the gasket is to create a space between the base of the chamber and the lid, a space inside which the tissue is sealed and through which the solution will flow. The space inside the gasket must include the inlet (solution enters here), the tissue prep, and the outlet through which the solution leaves the chamber. Two features of the gasket are critical: First, the gasket must be thick enough for the brain yet thin enough to allow imaging with a high-resolution lens. The ideal gasket thickness depends on set-up-specific parameters. Second, a notch in the gasket provides a compartment that allows a bubble to pass through without contacting the specimen (Fig 2B).
Imaging at the Microscope
We use a 63x (aperture 1.3) glycerine immersion lens for the perfusion chamber or a 63x water immersion lens directly into the culture medium on a slide. Glycerine lenses provide more working distance than oil lenses. We use a resonant scanning confocal microscope which scans at much faster rates than a conventional scanner (8000 Hz versus 1400 Hz, respectively). Resonant scanning allows 3-dimensional recordings over time at faster frame rates. In addition, faster scan speeds significantly reduce laser phototoxicity6 which is a major concern for repeated scans of live tissue. A balance of laser intensity and PMT voltage (gain) must be achieved to maximize signal while minimizing noise and photobleaching. An important further consideration is that the amount of laser impact on a given region is determined by resolution and zoom. For example, a region of choice is scanned at the same resolution at 256x256, zoom 2x as at 512x512, zoom 1x, yet the maximally possible live imaging speed is 4 times faster at 256x256, zoom 2x. To record the activity of quickly moving intracellular compartments, we have successfully recorded with the following settings: 5 frames per z-stack at a rate of one z-stack per 10 seconds with a scan speed of 8000 Hz and 2x line averaging. For long imaging sessions on the scale of hours, we often reduce the frame rate to one per several minutes and choose larger areas as the preparation is more likely to shift.
Drosophila Visual System Anatomy
Figure 3 shows the anatomy of the Drosophila adult eye (Fig. 3A), the adult brain (Fig. 3B), and the 20% -25% pupal eye disc/brain complex (Fig. 3C). Figure 3A illustrates the adult eye both with and without the lamina and fat cells attached. During the adult eye dissection, the lamina, which is in the center of the eye nestled in a bowl formed by the photoreceptor cell bodies (left), must be removed without disrupting the cell bodies underneath (right). For the adult brain dissection, one should be able to distinguish the photoreceptor cell bodies, the lamina, the optic lobe, the central brain, and trachea (Fig. 3B). The photoreceptors and trachea need to be removed for this dissection while leaving the lamina attached to the optic lobe. To aid in the identification of the 20-25% pupal eye disc/brain complex, a live image is presented in Figure 3C.
<img src="/files/ftp_upload/1936/1936fig1.jpg" alt="Insect brain anatomy, optic lobe, lamina, trachea, diagram, cell bodies, photoreceptor analysis." data-altupdatedat="1747911167000" data-alt="Figure 1"> Figure 1. (A) adult brain. (B) adult eye. (C) 20%-25% pupal eye-disc/brain complex.
<img src="/files/ftp_upload/1936/1936fig2.jpg" alt="Flow chamber diagram; microscopy setup for specimen observation; fluid dynamics and assembly process." data-altupdatedat="1747911167000" data-alt="Figure 2"> Figure 2. (A) Perfusion chamber assembly and (B) gasket positioning relative to the specimen.
<img src="/files/ftp_upload/1936/1936fig3.jpg" alt="Fluorescent microscopy images: cell structures, protein localization, experiment result comparison." data-altupdatedat="1747911167000" data-alt="Figure 3"> Figure 3. (A) Adult rhabdomere staining using anti-chaoptin. (B) Adult lamina staining using anti-chaoptin (green, photoreceptors), anti-ebony (red, glia), and sec6 (blue, interneurons).

<table><tbody><tr><td>Sylgard 184</td><td>Dow Corning</td><td>103251252</td><td></td></tr><tr><td>GluShield</td><td>GluStitch, Inc.</td><td>GB055QO</td><td></td></tr><tr><td>20-Hydroxyecdysone</td><td>Sigma-Aldrich</td><td>H5142-10mg</td><td>5mg/ml in EtOH</td></tr><tr><td colspan="4">&lt;p class=&quot;jove_title&quot;&gt;Special Equipment:&lt;/p&gt;&lt;ol&gt;&lt;li&gt;Dissection Scope: Leica Mz12.5 with digital camera&lt;/li&gt;&lt;li&gt;Leica TCS SP5 Confocal Tandem-Scanner Microscope for conventional and resonant scanning live&lt;br/&gt; imaging microscopy, including blue (488nm), green (561nm), orange (594nm) and far-red (633nm) lasers as well as special 63x, 1.4 glycerine and a 63x, 1.3 water immersion lenses.&lt;/li&gt;&lt;li&gt;Harvard Instruments Confocal Imaging Chamber RC-30 and peristaltic pump model 720&lt;/li&gt;&lt;li&gt;Sutter Pipette puller P-97&lt;/li&gt;&lt;li&gt;Software: Amira 5.2, Visage Imaging&lt;/li&gt;&lt;/ol&gt;</td></tr></tbody></table>

preparation of developing and adult drosophila brains and retinae for live imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Watch this Scientific Journal Video about Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging at JoVE.com

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

preparation-developing-adult-drosophila-brains-retinae-for-live

Research

JoVE Journal

Biology

35.4K Views.  University of Texas Southwestern Medical Center. Dit protocol beschrijft drie Drosophila Voorbereiding: 1) volwassen hersenen dissectie, 2) volwassen netvlies dissectie en 3) het ontwikkelen van eye-schijf-hersen-complexen dissectie. De nadruk wordt gelegd op speciale voorbereiding technieken en voorwaarden voor de live-imaging, maar alle voorbereidingen kunnen worden gebruikt voor vaste weefsel immunohistochemie.

Video: Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Ex vivo Culturing of Whole, Developing Drosophila Brains

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for investigating the cell biology and development of central brain structures by allowing acute pharmacological interventions and live imaging of cellular processes. As an example of the technique, prior work from our lab1 has shown that a previously unrecognized subcellular compartment lies between the axonal and somatodendritic compartments of axons of the Drosophila central brain. The development of this compartment, referred to as the axon initial segment (AIS)2, was shown genetically to depend on the neuron-specific cyclin-dependent kinase, Cdk5. We show here that ex vivo treatment of wild-type Drosophila larval brains with the Cdk5-specific pharmacological inhibitors roscovitine and olomoucine3 causes acute changes in actin organization, and in localization of the cell-surface protein Fasciclin 2, that mimic the changes seen in mutants that lack Cdk5 activity genetically.
A second example of the ex vivo culture technique is provided for remodeling of the connections of embryonic mushroom body (MB) gamma neurons during metamorphosis from larva to adult. The mushroom body is the center of olfactory learning and memory in the fly4, and these gamma neurons prune their axonal and dendritic branches during pupal development and then re-extend branches at a later timepoint to establish the adult innervation pattern5. Pruning of these neurons of the MB has been shown to occur via local degeneration of neurite branches6, by a mechanism that is triggered by ecdysone, a steroid hormone, acting at the ecdysone receptor B17, and that is dependent on the activity of the ubiquitin-proteasome system6. Our method of ex vivo culturing can be used to interrogate further the mechanism of developmental remodeling. We found that in the ex vivo culture setting, gamma neurons of the MB recapitulated the process of developmental pruning with a time course similar to that in vivo. It was essential, however, to wait until 1.5 hours after puparium formation before explanting the tissue in order for the cells to commit irreversibly to metamorphosis; dissection of animals at the onset of pupariation led to little or no metamorphosis in culture. Thus, with appropriate modification, the ex vivo culture approach can be applied to study dynamic as well as steady state aspects of central brain biology.

Ex vivo Culturing of Whole, Developing Drosophila Brains

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

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for ...

Neuroscience

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

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 ...

Developmental Biology

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

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

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

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 ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

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

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

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

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

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

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