This protocol was developed under a Wellcome Trust grant. We thank Ms. Maureen Sinclair for maintaining the fly stocks and preparing the fly food over the years. We would also like to thank Mr. Michael Aitchison for his help and technical support in developing of this protocol.

1. Transgenic Flies and Maintenance
<ol>
 <li>The transgenic flies used in this demonstration: Ubq-gfp-Cdc20 (II), Ubq-rfp-His2B (X*); Ubq-gfp-Cdc20 (II*), Ubq-gfp-Cdc20 (II*); mad2EY(III*) and Ubq-gfp-Cdc20 transgenic flies were previously generated in the lab via a standard P-element mediated transgenic approach10,11 and Ubq-rfp-His2B is a kind gift from Yohanns Bela&iuml;che at UMR 144 CNRS/Institute Curie, Paris, France. They were introduced into a Mad2 mutant background via standard Drosophila genetics. The mad2EY original mutant line was purchased from the Bloomington stock center. We will not discuss the procedure used for raising the transformants in this protocol. 
 Note: * represents the chromosome number.</li>
 <li>Maintenance: Transgenic flies were maintained at 25 &deg;C in plastic vials containing fly food and with additional dry yeast powder on the top. The vial was routinely replaced every 3-4 weeks depending on growing conditions (Figure 1).</li>
</ol>
2. Fly Food Preparation (Lab scale)
<ol>
 <li>An appropriate amount of the fly food mix was heated with constant stirring to dissolve the components.</li>
 <li>About 8-10 ml of this medium was distributed as slurry into each plastic vial (2.5 cm diameter x 8 cm length) using a Jencons Scientific Ltd peristaltic pump.</li>
 <li>When the food slurry was set and has cooled to room temperature, the vial is then plugged with a cotton foam plug. These foods vials are placed in a tray that is then sealed in a plastic bag and kept at 4 &deg;C for later use.</li>
</ol>
3. Small-scale Egg Collection
<ol>
 <li>About 50 pairs of 2-3 day old adult flies were transferred to a fresh fly food vial supplied with additional dry yeast powder on its surface at 25 &deg;C for laying embryos.</li>
 <li>The flies are then transferred to a fresh vial every hour and leave the embryos in the vial for 30 minutes to ensure some of the collected embryos are aged around nuclear division cycle 8-10 when the nuclei are gradually migrating to the cortex and organized as a single monolayer. The first hour collection is normally discarded as it often contains aged embryos that were retained in the female bodies when conditions weren't suitable for laying.</li>
</ol>
4. Preparing Coverslips and Slides
<ol>
 <li>Take out a 50 x 22 mm coverslip and slightly wet its four corners on one side with a very small amount of water using a moist fine pen brush and put it on a microscope slide so that the coverslip does not move because of the capillary surface tension caused by the thin liquid film.</li>
 <li>Apply a thin strip of heptane glue across the middle of the coverslip, the heptane should evaporate in seconds to leave the glue on coverslip.</li>
 <li>Cut another coverslip with a diamond pen into small squares (~1.5 mm2), pick up one and place it on one end of the glue strip (this is used to open the needle tip when microinjection is required) and save the others for future use.</li>
 <li>Take another microscope slide and stick a piece of double-sided sticky tape about 2 cm long to it and peel off the cover paper. (See Figure 2A &amp; B).</li>
</ol>
5. Dechorionate Embryos
<ol>
 <li>Transfer the flies to a fresh food vial and use a moistened fine pen brush to transfer about 50 eggs onto the double-sided sticky tape on the slide with an appropriate amount of liquid to allow the eggs to be spread out. When the liquid has evaporated the eggs will stick to the tape.</li>
 <li>Under a dissecting microscope touch the eggs several times along their long axis using the outer side of one half of a pair of tweezers gently till the chorion is broken and open but leave the embryo in the chorion shell to prevent rapid dehydration. Continue this process till 10-20 of the eggs has been treated (depending on the number of the eggs required for a single experiment).</li>
 <li>The embryos can then be picked up and removed from the chorion shell and gently placed on the glue strip one by one. The eggs could be placed and aligned either with the long side of the embryo parallel or anti-parallel to the long side of the glue strip depend on if they are to be microinjected.</li>
 <li>The embryos can either be covered immediately with an appropriate amount of Voltalef 10S oil (a 5:1 mixture of Holocarbon 700 oil and 27 oil, Sigma) to prevent further desiccation for immediate imaging or placed in a desiccation chamber for 3-8 minutes to dehydrate them further depending on the humidity of the room and the purpose of the experiment. The dehydrated embryos should be covered with 10S oil when appropriate to prevent over-dehydration if microinjection is required. (See Figure. 2C, D &amp; E).</li>
</ol>
6. Imaging Embryos
<ol>
 <li>Time-lapse or single images of living embryos are collected using a Leica TCS SP2 inverted confocal microscopy system with a HCX PL APO CS 40X 1.25 oil immersion objective (60 or 100X objectives can be used too depending on the purpose of the experiment, the higher the magnification used the more photo bleaching will occur).</li>
 <li>When imaging multiple fluorophores, which have overlapping spectra and are co-expressed in the same embryo, each fluorescent protein emission wavelength was collected sequentially. The excitation and emission wavelengths available on the Leica TCS SP2 system are: GFP, &lambda;Ex: 488 nm and &lambda;Em: 500-600 nm. RFP, &lambda;Ex: 543 nm and &lambda;Em: 555-700 nm.</li>
 <li>A single image was commonly obtained as an average of 2 scanlines with an average of 2 frame-scans. This takes a minimum of 6.3 seconds to scan a 512x512 pixel image in a simultaneous scan mode, and 15.44 seconds for a sequential scan mode and produces a good signal-to-noise ratio. These times need to be taken into account when setting up the time frames for time-lapse image collecting.</li>
 <li>The pinhole was normally set to 1-2 AU (airy unit), and the laser power was adjusted to the lowest level possible to avoid any significant photobleaching.</li>
</ol>
7. Provoke the SAC by Microinjecting the Embryos with Appropriate Reagents
<ol>
 <li>Prepare the needle before the experiment. We prefer to pull the needle from siliconised glass capillaries (GC-100-10, Harvard) using the flaming/brown micropipette puller (Sutter Instrument, Model: P-97) with parameters: heat = 600, pull = 90, vel = 70, del = 150.</li>
 <li>Backfill the needle with 1-2 ml of an appropriate concentration of the reagent in injection buffer using a fine loading tip (20 &mu;l Eppendorf lot: W215818J).</li>
 <li>Mount the needle in the needle holder on the microscope with the pressure pipe connected to the Eppendorf injection control system.</li>
 <li>Find an embryo under the 40X objective and find the needle shadow without changing the focal plane by moving the needle into the center of the field of view and gradually lowering it to the right focal plane.</li>
 <li>Move the embryo away gently to find the small square of the broken coverslip pre-mounted at one end of the glue strip. Very gently move the needle tip until it hits the edge of the small square coverslip and gently breaks it open.</li>
 <li>Move the embryos back into view and select the embryo of the right age for injection. I normally inject the embryo at nuclear division cycle at 5-7 as this is before the nuclei migrate outwards to the cortex. The nuclear division cycle stages can be determined by a quick scan of the fluorescent signals of the GFP-Cdc20 or the nuclear signal of the RFP-Histone 2B prior to injection9.</li>
 <li>Carefully move the embryo close to the needle and readjust the focal plane for both the embryo and the needle. Move the embryo into the needle and inject a drop of the reagent into the desired position and move the embryo away (or follow the instructions for the microinjection system). The embryo can be injected from its posterior or from the side depending on the purpose of the experiment.</li>
 <li>Once injected the embryo is ready to be imaged at the appropriate time. (See Figure 2F &amp; G)</li>
</ol>
8. Representative Results
<img alt="Figure 1" src="/files/ftp_upload/3763/3763fig1.jpg" /> 
Figure 1. Required materials: a. fine pen brush, b. small square broken-glasses container, c. 22 x 50 mm coverslip, d. microscope slide, e. double-sided sticky tape, f. dry yeast powder in test tube with holes on the lid, g. yeast granule used for fly maintenance, h. fly food vial, i. a half of a tweezers used for dechorionate embryos, j. a pair tweezers, k. heptane glue liquid container (volumetric flask).
<img alt="Figure 2" src="/files/ftp_upload/3763/3763fig2.jpg" /> 
Figure 2. The diagrams show the preparation of the coverslips and slides in step 4, embryo dechorionation in step 5 and needle preparations for microinjection in step 7. A. The top picture shows a coverslip with a strip of heptane glue across its middle and a small square of a broken coverslip stuck to one end. The bottom picture shows a microscope slide with a thin layer of water at its four corners to hold a coverslip. The reason for using a slide to hold the coverslip is to keep roughly the same focal distance as that found when you have double-sided tape on the slides and thus avoiding refocusing the microscope while you are dissecting and transferring the embryos between tape and coverslip. B. A slide with a short length of double-sided Scotch tape applied before peeling off its cover paper used for dechorionating the embryo. C. The picture on the left shows embryos with intact chorion shells with marked anterior and posterior ends; the picture on the right shows the embryos in the broken chorion shells before they were transferred onto the coverslip with the glue strip. D &amp; E. Diagrams showing two ways for transferring, placing and aligning the dechorionated embryos on glue strips. The embryos were covered by 10S oil after an appropriate desiccation period. F &amp; G. Diagrams showing how to open the tip of the needle and position it so as to inject.
Single or time-lapse images obtained from the above described experiments can be directly analyzed using Leica software or saved in .TIF files as an open format available for further quantification or editing analyses using other common image analysis programs such as Image J, Photoshop and MetaMorph etc. Two examples to illustrate this are discussed below:
Example 1: A time-lapse movie (Figure 3) shows the dynamic kinetochore recruitment of GFP-Cdc20 and chromosome motion in living Drosophila syncytial embryos. The region of interest from original time-lapse sequence images was determined/edited and automated-batch converted using Photoshop software. The movie was assembled using QuickTime software.
<img alt="Figure 3" src="/files/ftp_upload/3763/3763fig3.jpg" /> 
Figure 3. A time-lapse movie shows the dynamic kinetochore recruitment of GFP-Cdc20 and chromosome motion in living Drosophila syncytial embryos. (A) Time-lapse images were taken from a transgenic syncytial embryo co-expressing GFP-Cdc20 (in green) and RFP-Histone 2B (in red) fusion proteins and were recorded using a Leica TCS SP2 confocal system at 18 &deg;C during nuclear division cycles 7-8. Frames were taken every 10 seconds. The frame with the cells already in prophase is treated as the zero time point10. <a href="https://www.jove.com/files/ftp_upload/3763/3763movie3A.mov" target="_blank">Click here to view movie for Figure 3A</a>. (B) GFP-Cdc20 can be readily observed on prophase and prometaphase kinetochores (B3 &amp; 4, white arrows) and persists on metaphase and anaphase kinetochores (B5 &amp; 6, white arrows). GFP-Cdc20 is excluded from interphase nucleus (White arrowhead), entering the nucleus by early prophase. Chromatin morphologies were determined using co-expressed His2BmRFP as markers (B8-14). Bars= 5mm.
Example 2: SAC functions can be studied by manipulating the embryos by microinjecting antibodies, fluorescently labeled proteins or chemical compounds of interest that potentially trigger the SAC. For instance injecting colchicine to depolymerize the microtubules so provoking the SAC as indicated in Figure 410.
<img alt="Figure 4" src="/files/ftp_upload/3763/3763fig4.jpg" /> 
Figure 4. Mad2 is essential for colchicine-invoked SAC function10. gfp-cdc20; mad2+/+, gfp-cdc20; mad2EY (mad2-/- null mutant) and gfp-mad2; mad2EY embryos were treated with colchicine by microinjection. Time-lapse confocal images were taken before (01, 07 &amp; 13) or after injection (02-06, top panels; 08-12, middle panels; 14-18, bottom panels). GFP-Cdc20 (top and middle panels) or GFP-Mad2 (bottom panel) kinetochore signals were used as cell cycle progression markers. Top panel, the arrows indicate the arrested kinetochores in 02-06. The area marked by an arrow in 01 indicates that before colchicine treatment, GFP-Cdc20 is excluded from the late interphase nucleus. Middle panel, in the absence of endogenous Mad2, GFP-Cdc20 signals continue to oscillate in and out of the nucleus, and on and off the kinetochores, although cytokinesis appears to be defective, as would be expected in embryos lacking microtubules, in the presence of colchicine (indicated by arrows in 07-12) suggesting a failed SAC function in arresting cells in response to colchicine treatment. Separation of daughter nucleus failed in picture 10. Bottom panel, arrowheads in 14-18 indicate arrested kinetochores with accumulated GFP-Mad2 fusion proteins to suggest the functional GFP-Mad2 in rescue the SAC defect phenotype in Mad2 mutant embryo. Arrowhead in 13 indicates GFP-Mad2 accumulation in a late interphase nucleus. Bar=5mm. The embryos were microinjected with ~1% egg volume of a 100mg/ml colchicine stock solution in 1 x PBS.

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flies+without+a+spindle+checkpoint.">Flies without a spindle checkpoint.</a> Nat. Cell Biol. 9, 565-572 (2007).</li><li>Tang, Z., Shu, H., Oncel, D., Chen, S., Yu, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phosphorylation+of+Cdc20+by+Bub1+provides+a+catalytic+mechanism+for+APC/C+inhibition+by+the+spindle+checkpoint.">Phosphorylation of Cdc20 by Bub1 provides a catalytic mechanism for APC/C inhibition by the spindle checkpoint.</a> Mol. Cell. 16, 387-397 (2004).</li><li>Huang, J., Raff, J. 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We have nothing to disclose.

The protocol described here is a generic method for imaging fly syncytial embryos using a Leica TCS SP2 confocal laser scanning microscope and can be modified to suit other microscope systems and can also be adapted to study other gene functions using Drosophila syncytial embryos. We have used this protocol to study many aspects of the spindle assembly checkpoint, protein dynamics and protein proteolysis using transgenic flies or polyclonal antibodies to visualize the protein localisation in living or fixed samples4,6,12-14 .
It is simplest to keep the flies in fresh fly food vials when collecting small numbers (~10 - 100) of eggs. This reduces the hassle of making and preparing additional apple juice agar plates and egg collection chambers as described in other publications15,16. The addition of a small square of broken coverslip glass at one end of the glue strip on the coverslip makes it a lot easier to open the needle and determine the injection volume by adjusting the microinjection system settings while the needle is immersing in the 10S oil. The degree of desiccation of the embryo is important for the success of the injection in avoiding leakages and maintaining the embryos viability especially for those experiments which require a long period of time or when raising transformants after injection. However, there is no golden rule for exactly how long desiccation is required. This varies from experiment to experiment and depends on the amount of the solution injected and the humidity of the working environment.
The functions of a specific protein of interest can be manipulated by microinjecting specific protein inhibitors, mono- or poly-clonal antibodies against a specific protein, messenger RNA or fluorescently labeled peptides under wild type or genetic mutation backgrounds. Many of these mutant lines or GFP-tagged transgenic lines are publicly available from Drosophila stock centers and most of them are listed on Flybase (<a href="http://flybase.org/" target="_blank">http://flybase.org/</a>) and can be searched for online.

Essential equipment and reagents:
Confocal imaging system: The imaging system described in this protocol is a Leica TCS SP2 laser scanning confocal inverted microscope system. The method described is also suitable perhaps with some minor modifications for other imaging systems.
Dissecting microscope: We use an Olympus SZX7 with DF PLAPO 1X-4 lens.
Needle puller: Flaming/brown micropipette puller (from Sutter Instrument, Model: P-97).
Microinjection system: An Eppendorf microinjection system is described but any other appropriate system can be used.
Fly food distributor: Jencons Scientific Ltd peristaltic pump.
Reagents:
<table border="1">
	<tbody>
		<tr>
			<td>Ingredients</td>
			<td>Weights</td>
			<td>Resources</td>
			<td>Lot No.</td>
		</tr>
		<tr>
			<td>Maize meal</td>
			<td>100.0g</td>
			<td>SUMA, UK</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Brown sugar</td>
			<td>50.0g</td>
			<td>Billington's, UK</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dry yeast</td>
			<td>25.0g</td>
			<td>DCL YEAST Ltd. UK</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Agar</td>
			<td>12.5g</td>
			<td>Fisher Scientific</td>
			<td>106556</td>
		</tr>
		<tr>
			<td> Sorbic acid</td>
			<td>0.4g</td>
			<td>BDH, VWR International Ltd. UK</td>
			<td>8829310</td>
		</tr>
		<tr>
			<td>Benzoic acid</td>
			<td>2.9g</td>
			<td>Fisher Scientific</td>
			<td>1019599</td>
		</tr>
		<tr>
			<td>Nipagin 
			(Methyl -4-Hydro 
			xybenzoate)</td>
			<td>0.9g</td>
			<td>BDH, VWR International Ltd. UK</td>
			<td>K35969015</td>
		</tr>
		<tr>
			<td colspan="3">H2O up to 1L</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>
Table 1. Fly food ingredients.
Holocarbon 700 and 27 oils purchased from Sigma.
Dry yeast: Thomas Allison, UK.
Preparing the heptane glue: Take about 1.5 meters of double-sided Scotch tape, put it into a 15ml Falcon tube with 5ml of heptane and rotate for 3-5 hours or overnight. After that time split into 4 equal aliquots in 1.5 ml eppendorf centrifuge tubes and spin for 5 min at a fast speed in a bench-top centrifuge to remove any debris. Store the heptane glue solution in a 10ml volumetric flask and keep for use. The glue strength will vary according to the concentration and the amounts of the glue used. Normally, the thinner glue produces lesser noisy background when scanned by confocal microscope.
<ul>
	<li>Coverslip: BDH, VWR International Ltd. UK</li>
	<li>Microscope slide: BDH, VWR International Ltd. UK</li>
	<li>Siliconised glass capillaries: GC-100-10, Harvard</li>
	<li>Fine loading tip: 20&mu;l eppendorf lot: W215818J</li>
	<li>Tweezers</li>
	<li>Fine pen brush</li>
</ul>

studying mitotic checkpoint by illustrating dynamic kinetochore protein behavior and chromosome motion in living drosophila syncytial embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Watch this Scientific Journal Video about Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos at JoVE.com

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

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JoVE Journal

Biology

11.3K Views.  University of Newcastle, United Kingdom. The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

Video: Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

Studying Aggression in Drosophila (fruit flies)

Aggression is an innate behavior that evolved in the framework of defending or obtaining resources. This complex social behavior is influenced by genetic, hormonal and environmental factors. In many organisms, aggression is critical to survival but controlling and suppressing aggression in distinct contexts also has become increasingly important. In recent years, invertebrates have become increasingly useful as model systems for investigating the genetic and systems biological basis of complex social behavior. This is in part due to the diverse repertoire of behaviors exhibited by these organisms. In the accompanying video, we outline a method for analyzing aggression in Drosophila whose design encompasses important eco-ethological constraints. Details include steps for: making a fighting chamber; isolating and painting flies; adding flies to the fight chamber; and video taping fights. This approach is currently being used to identify candidate genes important in aggression and in elaborating the neuronal circuitry that underlies the output of aggression and other social behaviors.

Studying Aggression in Drosophila fruit flies

Aggression is an innate behavior that evolved in the framework of defending or obtaining resources.  This complex social behavior is influenced by ...

Application of Light-cured Dental Adhesive Resin for Mounting Electrodes or Microdialysis Probes in Chronic Experiments

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these acrylics do not bond to the bone, screws have been used as anchors. However, in small experimental animals like finches or mouse, their craniums are very fragile and can not successfully hold the anchors. In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium. Therefore, anchor screws are not required and surgical field can be reduced considerably. Past experiences show that the bonding effect maintains more than 2 months. Conventional resin's window of time when the materials are pliable and workable is a few minutes. However, the window of working time for these dental adhesives is significantly wider and adjustable.

In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium.

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these ...

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

RNAi Interference by dsRNA Injection into Drosophila Embryos

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements and tools for genetic manipulation have become available in Drosophila 2. Soon after the initial discovery by Frie and Mello 3 that double stranded RNA can be used to knockdown the activity of individual genes in Caenorhabditis elegans, RNA interference (RNAi) was shown to provide a powerful reverse genetic approach to analyze gene functions in Drosophila organ development 4, 5.
Many organs, including lung, kidney, liver, and vascular system, are composed of branched tubular networks that transport vital fluids or gases 6, 7. The analysis of Drosophila tracheal formation provides an excellent model system to study the morphogenesis of other tubular organs 8. The Berkeley Drosophila genome project has revealed hundreds of genes that are expressed in the tracheal system. To study the molecular and cellular mechanism of tube formation, the challenge is to understand the roles of these genes in tracheal development. Here, we described a detailed method of dsRNA injection into Drosophila embryo to knockdown individual gene expression. We successfully knocked down endogenous dysfusion(dys) gene expression by dsRNA injection. Dys is a bHLH-PAS protein expressed in tracheal fusion cells, and it is required for tracheal branch fusion 9, 10. dys-RNAi completely eliminated dys expression and resulted in tracheal fusion defect. This relatively simple method provides a tool to identify genes requried for tissure and organ development in Drosophila.

RNAi Interference by dsRNA Injection into Drosophila Embryos

RNA interference has been proven very effective to analyze gene function in Drosophila tracheal development. A detailed protocol used by Jiang lab to inject dsRNA into fly embryos to knockdown gene expression is illustrated. This technique has the potential for screening genes required for tissue and organ development in Drosophila.

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

An in vivo Crosslinking Approach to Isolate Protein Complexes From Drosophila Embryos

Many cellular processes are controlled by multisubunit protein complexes. Frequently these complexes form transiently and require native environment to assemble. Therefore, to identify these functional protein complexes, it is important to stabilize them in vivo before cell lysis and subsequent purification. Here we describe a method used to isolate large bona fide protein complexes from Drosophila embryos. This method is based on embryo permeabilization and stabilization of the complexes inside the embryos by in vivo crosslinking using a low concentration of formaldehyde, which can easily cross the cell membrane. Subsequently, the protein complex of interest is immunopurified followed by gel purification and analyzed by mass spectrometry. We illustrate this method using purification of a Tudor protein complex, which is essential for germline development. Tudor is a large protein, which contains multiple Tudor domains - small modules that interact with methylated arginines or lysines of target proteins. This method can be adapted for isolation of native protein complexes from different organisms and tissues.

An in vivo Crosslinking Approach to Isolate Protein Complexes From Drosophila Embryos

Multi-component protein complexes play crucial roles during cellular function and development. Here we describe a method used to isolate native protein complexes from Drosophila embryos after in vivo crosslinking followed by purification of the crosslinked complexes for subsequent structure-function analysis.

Many cellular processes are controlled by multisubunit protein complexes. Frequently these complexes form transiently and require native environment to ...

Genome-wide Protein-protein Interaction Screening by Protein-fragment Complementation Assay (PCA) in Living Cells

Proteins are the building blocks, effectors and signal mediators of cellular processes. A protein&#8217;s function, regulation and localization often depend on its interactions with other proteins. Here, we describe a protocol for the yeast protein-fragment complementation assay (PCA), a powerful method to detect direct and proximal associations between proteins in living cells. The interaction between two proteins, each fused to a dihydrofolate reductase (DHFR) protein fragment, translates into growth of yeast strains in presence of the drug methotrexate (MTX). Differential fitness, resulting from different amounts of reconstituted DHFR enzyme, can be quantified on high-density colony arrays, allowing to differentiate interacting from non-interacting bait-prey pairs. The high-throughput protocol presented here is performed using a robotic platform that parallelizes mating of bait and prey strains carrying complementary DHFR-fragment fusion proteins and the survival assay on MTX. This protocol allows to systematically test for thousands of protein-protein interactions (PPIs) involving bait proteins of interest and offers several advantages over other PPI detection assays, including the study of proteins expressed from their endogenous promoters without the need for modifying protein localization and for the assembly of complex reporter constructs.

Genome-wide Protein-protein Interaction Screening by Protein-fragment Complementation Assay PCA in Living Cells

Proteins interact with each other and these interactions determine in a large part their functions. Protein interaction partners can be identified at high-throughput in vivo using a yeast fitness assay based on the dihydrofolate reductase protein-fragment complementation assay (DHFR-PCA).

Proteins are the building blocks, effectors and signal mediators of cellular processes. A protein&#8217;s function, regulation and localization often ...