Name: preparation of developing and adult drosophila brains and retinae for live imaging
Uploaded: 2010-03-15T17:55:00
Duration: 16 min 47 s
Description: Watch this Scientific Journal Video about Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging at JoVE.com

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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R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optic+lobe+development.">Optic lobe development.</a> Adv Exp Med Biol. 628, 115-136 (2008).</li><li>Ashburner, M., Golic, K. G., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+-+A+Laboratory+Handbook.">Drosophila - A Laboratory Handbook.</a> , (2005).</li><li>Featherstone, D. E., Chen, K., Broadie, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Harvesting+and+preparing+Drosophila+embryos+for+electrophysiological+recording+and+other+procedures.">Harvesting and preparing Drosophila embryos for electrophysiological recording and other procedures.</a> J Vis Exp. , (2009).</li><li>Borlinghaus, R. 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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 1&mu;M.
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="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="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="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).

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		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>103251252</td>
<td>&nbsp;</td>
<tr>
<td>GluShield</td>
<td>&nbsp;</td>
<td>GluStitch, Inc.</td>
<td>GB055QO</td>
<td>&nbsp;</td>
<tr>
<td>20-Hydroxyecdysone</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>H5142-10mg</td>
<td>5mg/ml in EtOH</td>
</tr>		</tbody>
		</table>
		</div>
			<div>
			Special Equipment:
<ol>
	<li>Dissection Scope: Leica Mz12.5 with digital camera</li>
	<li>Leica TCS SP5 Confocal Tandem-Scanner Microscope for conventional and resonant scanning live 
 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.</li>
	<li>Harvard Instruments Confocal Imaging Chamber RC-30 and peristaltic pump model 720</li>
	<li>Sutter Pipette puller P-97</li>
	<li>Software: Amira 5.2, Visage Imaging</li>
</ol>		</div>

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

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

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 ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function. Our studies have found that dying cells send signals to their live neighbors to form and contract a ring of actin and myosin that ejects it out from the epithelial sheet while closing any gaps that might have resulted from its exit, a process termed cell extrusion1. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe a method to induce and image extrusion in the larval zebrafish epidermis. To visualize extrusion, we inject a red fluorescent protein labeled probe for F-actin into one-cell stage transgenic zebrafish embryos expressing green fluorescent protein in the epidermis and induce apoptosis by addition of G418 to larvae. We then use time-lapse imaging on a spinning disc confocal microscope to observe actin dynamics and epithelial cell behaviors during the process of apoptotic cell extrusion. This approach allows us to investigate the extrusion process in live epithelia and will provide an avenue to study disease states caused by the failure to eliminate apoptotic cells.

Dying cells are extruded from epithelial tissues by concerted contraction of neighboring cells without disrupting barrier function. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe methods to induce and image extrusion in the larval zebrafish epidermis at cellular resolution.

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function.  Our studies have found ...

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 ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...