The authors thank members of the Veraksa laboratory for helpful comments on the manuscript and suggestions for protocol improvements. A.V. was supported by the NIH grants GM105813 and NS096402. L.Y. was supported by the University of Massachusetts Boston Sanofi Genzyme Doctoral Fellowship.

1. Preparation of 5 L Fly Cages (to Fit 15 cm Plates)
<ol>
	<li>Obtain the materials needed: 5-quart (4.7-L) container with lid (Figure 1A), nylon mesh, and a razor blade.</li>
	<li>Mark a 12 cm diameter circle and cut a hole in the bottom of the container using the razor blade (Figure 1B). Cut a 15 cm diameter hole in the lid (Figure 1C).</li>
	<li>Cut nylon mesh into 25 x 25 cm squares.</li>
	<li>Place the mesh over the top rim of the container and press down with the lid (Figure 1C). Make sure the mesh is tight and the lid is not tilted. The cage is ready to be populated with flies (see step 5.3).</li>
</ol>
2. Preparation of 1 L Fly Cages (to Fit 10 cm Plates)
<ol>
	<li>Obtain the materials needed: an electric drill, a 3 inch (76 mm) cutting tool (Figure 1E and Figure 1F), a straight-sided 1 L plastic jar with lid, nylon mesh, a razor blade, sandpaper, goggles, and work gloves.</li>
	<li>Cut 3 inch (76 mm) holes in the bottom and the lid of the plastic jar (Figure 1G), with the lid tightly screwed onto the container while drilling. Position the cutting tool in the center of the circle when starting to make a hole. 
	Caution: Wear eye protection and work gloves during this step.</li>
	<li>Smoothen the edges of the holes with a razor blade and sandpaper.</li>
	<li>Cut nylon mesh into 18 x 18 cm squares.</li>
	<li>Place the mesh over the top rim of the jar and tighten by screwing the lid on (Figure 1G). Make sure the mesh is tight and the lid is not tilted. The cage is ready to be populated with flies (see step 5.3).</li>
</ol>
3. Preparation of Apple Juice (AJ) Plates
<ol>
	<li>Add 19 g agar and 1 L of 18.2 M&#937;&#183;cm water into a 2 L flask. Add a stir bar, and mix briefly.</li>
	<li>Autoclave for 30 min. Meanwhile, thaw 100% apple juice concentrate from frozen stock, and prepare 10 mL of 10% (weight/volume) methyl 4-hydroxybenzoate solution in ethanol.</li>
	<li>After autoclaving, start mixing the agar on a stir plate, and add 80 mL of 100% apple juice concentrate and 10 mL of 10% (weight/volume) methyl 4-hydroxybenzoate solution to the flask. Let cool at room temperature with constant mixing.</li>
	<li>Pour 15 cm Petri dishes (for 5 L cages) or 10 cm Petri dishes (for 1 L cages) so that the level of agar in each dish is approximately 5 mm thick. Allow to solidify at room temperature, and wrap in plastic bags. AJ plates can be stored at 4 &#176;C for two weeks. 
	Note: Petri dishes can be washed and reused.</li>
	<li>Prepare wet yeast paste: pour some active dry yeast into a 100 mL container with a lid that would allow easy mixing, and add water in small amounts while stirring, to obtain a thick but thoroughly mixed paste. Store at 4 &#176;C for up to 2 weeks.</li>
</ol>
4. Preparation of Lysis Reagents
<ol>
	<li>Prepare 5x lysis buffer: 250 mM Tris pH 7.5, 25% glycerol, 1% octylphenoxy poly(ethyleneoxy)ethanol (IGEPAL CA-630), 7.5 mM MgCl2, 625 mM NaCl, 125 mM NaF, 5 mM Na3VO4.
	<ol>
		<li>To make 200 mL: completely dissolve 1 g NaF powder and 184 mg Na3VO4 powder in 71 mL water with constant stirring. Add 50 mL 1 M Tris pH 7.5, 2 mL 100% IGEPAL, 1.5 mL 1 M MgCl2, and 25 mL 5 M NaCl. Then add 50 mL glycerol (add last) and continue stirring for 1 h.</li>
		<li>Filter-sterilize using 250-mL filter units. This step can be slow, but filtering the solution is important to remove insoluble particulates. Aliquot by 10 mL in 50 mL tubes and store at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare 1 M dithiothreitol (DTT) solution in water. Store at -20 &#176;C in 1 mL aliquots.</li>
	<li>Obtain protease inhibitor cocktail tablets, with ethylenediaminetetraacetic acid (EDTA). Store at 4 &#176;C.</li>
	<li>Obtain 10 mL syringes and syringe filters: 26 mm diameter, surfactant-free cellulose acetate (SFCA) membrane, 0.45 &#181;m pore size.</li>
</ol>
5. Collection of Fly Embryos
<ol>
	<li>Amplify the transgenic fly lines carrying the tagged protein of interest in bottles (approximately 100 flies per bottle), and transfer the bottles every 2 days until 25-30 bottles are accumulated for each fly line. Amplify the control stock (e.g. the y w line) in a similar way.</li>
	<li>Warm AJ plates to room temperature (about 1-2 h, can be done overnight). Using a gloved finger, spread some wet yeast paste in the center of AJ plates, to make an approx. 6-cm circle for a 15 cm plate and a 4 cm circle for a 10 cm plate.</li>
	<li>Anesthetize the flies with a flow of carbon dioxide using a standard Drosophila stage setup, and transfer into the cages (approx. 15 well-grown bottles of flies per 5 L cage, 5 bottles per 1 L cage).
	<ol>
		<li>Cover the cage bottom with prepared AJ plates from step 5.2, and tape the AJ plate to the cage using two strips of tape (Figures 1D and 1H). It is useful to fold over the tips of the tape that go over the plate for easy peeling when changing plates.</li>
		<li>After the flies recover from anesthesia, keep the cages with the plates at the bottom and mesh on top. 
		Note: It is important to wait until the flies fully recover, as they will otherwise stick to the AJ plate and yeast paste.</li>
		<li>At the end of the experiment, to prepare the cage for cleaning, place the cage in the freezer for several hours. Soaking the mesh in water for a few hours will facilitate cleaning.</li>
	</ol>
	</li>
	<li>Incubate the cages with flies at room temperature for 2-3 days and change AJ plates twice a day. To change the plates, turn the cage upside down, peel the tape from the plate but hold the plate to the cage, tap the whole cage gently on the bench, and quickly swap the old plate for a new one, trying not to allow flies to escape. 
	Note: Flies need to adjust to the cage and will lay best starting on day 3. They will continue laying at peak level for about 4-5 days. Two 5 L cages at top laying capacity can produce up to 1 g of embryos in an overnight collection. Use 1 L cages for smaller-scale experiments.</li>
	<li>Allow flies to lay embryos on fresh AJ plates overnight (approx. 16 h), or for any other desired length of time.</li>
	<li>The next morning, mark and weigh mesh containers for embryo collection, one per each fly line used.</li>
	<li>Prepare 1x lysis buffer, using reagents from section 4 (Note: ready-to-use 1x lysis buffer has to be prepared right before extraction): add 40 mL water to 10 mL of 5x concentrated lysis buffer (stored at -80 &#176;C) in a 50 mL tube.
	<ol>
		<li>Add 50 &#181;L of 1 M DTT to a final concentration of 1 mM, mix well and separate into two 50 mL tubes, 25 mL in each. To one of the tubes, add one protease inhibitor tablet and rotate at 4 &#176;C for 30 min. While the tablet is dissolving, collect and dechorionate embryos (steps 5.8 to 6.6). Check to make sure the tablet has fully dissolved, and keep the buffer on ice. 
		Note: The second tube of 1x buffer with DTT can be stored at -80 &#176;C and will only require addition of the tablet before the next experiment. 25 mL of lysis buffer is sufficient for up to 2 samples, taking into account subsequent washes during protein purification steps.</li>
	</ol>
	</li>
	<li>Mark the AJ plates on the cages with the genotypes of the fly lines used, and swap the plates for fresh ones. 
	Note: Excess yeast paste can be scraped off the plate at this step. Embryos can be aged on the plates at 25 &#176;C or room temperature to obtain the desired window of developmental time.</li>
	<li>Add enough water to cover each AJ plate. Gently dislodge the embryos from the plate with a soft paint brush (flat head approx. 10 mm wide works well for large plates). Pour the embryos into the previously weighed and marked mesh container, and drain excess water.
	<ol>
		<li>If needed, perform a second rinsing and brushing of the plate to remove all the embryos. Combine the embryos from additional plate(s) for the same fly line into the same mesh container.</li>
	</ol>
	</li>
	<li>Briefly blot the mesh on paper towels to remove excess water.</li>
</ol>
6. Embryo Dechorionation
<ol>
	<li>Half-fill a 6 cm Petri dish with 50% bleach (vol/vol solution in water). 
	Note: See important comment about bleach brand in the Table of Materials and Reagents. 
	Caution: Wear gloves and avoid getting bleach on clothes.</li>
	<li>Prepare 2 larger Petri dishes with water.</li>
	<li>Immerse the mesh container with embryos in 50% bleach for 90 s. Tap the mesh gently during incubation a couple of times to suspend the embryos in the bleach solution.</li>
	<li>At the end of the 90 s incubation, wash the mesh container in each of the 2 dishes with water for about 10 s.</li>
	<li>Rinse the mesh container thoroughly with 18.2 M&#937;&#183;cm water using the squirt bottle. Also, wash the sides and the outside of the mesh container. 
	Note: No bleach smell should be detected after a thorough wash.</li>
	<li>Blot the mesh container on paper towels. Weigh the mesh container with the embryos and subtract the weight of the empty container obtained in step 5.6. Record the resulting weight of the embryos. A good amount of embryos to aim for is 0.5 - 1 g.</li>
</ol>
7. Preparation of Embryo Extract
<ol>
	<li>Add 1 mL of lysis buffer with protease inhibitors (from step 5.7) to a dounce glass homogenizer and keep it on ice. Transfer the embryos from the mesh container into the homogenizer. 
	Note: Embryos can be picked up with a flat end of a metal spatula or a brush and transferred directly into the buffer. Perform all steps in this section on ice.</li>
	<li>Wash off the remaining embryos on the mesh with an additional 1 mL lysis buffer and add to the material in the homogenizer. Let the embryos settle down to the bottom of the homogenizer, and carefully aspirate the supernatant.</li>
	<li>Add lysis buffer with protease inhibitors (from step 5.7) to the homogenizer, aiming for 5-6 mL of lysis buffer per gram of embryos.</li>
	<li>Homogenize the embryos by 4-6 strokes with a loose fitting pestle. 
	Note: During the first strokes, there will be more resistance. Try to move the pestle in a continuous motion to avoid splashing the solution.</li>
	<li>Homogenize the embryos by 8-10 strokes with a tight fitting pestle.</li>
	<li>Incubate the homogenate on ice for 20 min.</li>
	<li>Transfer the homogenate into microcentrifuge tubes and centrifuge at 4 &#176;C for 15 min at top speed (approx. 13,000 x g).</li>
	<li>Avoiding the pellets and the very top lipid layer, transfer the supernatants into fresh microcentrifuge tubes and centrifuge at 4 &#176;C for 15 min at top speed (approx. 13,000 x g).</li>
	<li>Carefully aspirate supernatants with 1 mL tip and load into chilled 10 mL syringes with 0.45 &#181;m filters attached (combine aliquots of the same sample into one syringe). Push all of the solution into a labeled 15 mL tube on ice. The extract can be immediately used for subsequent protein purification experiments or snap-frozen in liquid nitrogen and kept at -80 &#176;C for future use. 
	Note: It is better to freeze the extracts rather than the embryos, because during subsequent thawing of embryos proteases may become activated and lead to protein degradation.</li>
</ol>

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N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+expression+during+the+life+cycle+of+Drosophila+melanogaster.">Gene expression during the life cycle of Drosophila melanogaster.</a> Science. 297 (5590), 2270-2275 (2002).</li><li>Soeller, W. C., Poole, S. J., Kornberg, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+transcription+of+the+Drosophila+engrailed+gene.">In vitro transcription of the Drosophila engrailed gene.</a> Genes Dev. 2 (1), 68-81 (1988).</li><li>Kamakaka, R. T., Kadonaga, J. 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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Raising+large+quantities+of+Drosophila+for+biochemical+experiments.">Raising large quantities of Drosophila for biochemical experiments.</a> Methods Cell Biol. 44, 99-108 (1994).</li><li>Sisson, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culturing+large+populations+of+Drosophila+for+protein+biochemistry.">Culturing large populations of Drosophila for protein biochemistry.</a> CSH Protoc. 2007, (2007).</li><li>Caravaca, J. M., Lei, E. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maintenance+of+a+Drosophila+melanogaster+Population+Cage.">Maintenance of a Drosophila melanogaster Population Cage.</a> J Vis Exp. 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The authors have nothing to disclose.

The protocol presented here is a simple and general procedure for setting up Drosophila population cages at medium scale and making whole-cell protein extracts from embryos. The resulting extracts can be used in a variety of downstream applications, such as purification of protein complexes on affinity resins. It is critical to perform the extraction steps on ice and use strong protease inhibition, to minimize protein degradation. As described, the protocol is suitable for isolation of most cytoplasmic and some ...

Genetic screens and, more recently, genomic approaches have revolutionized the study of biological functions. However, important cellular information is encoded in proteins and their ensemble of interacting partners. While traditional genetic modifier screens can identify rate-limiting pathway components and recover indirect interactions, the strength of the proteomic approaches lies in their ability to identify complete immediate interaction networks of proteins of interest. Proteomics is thus a valuable orthogonal method to study biological systems, and complements genomics, transcriptomics, and traditional genetic screens. Affinity purification-mass spectrometry (AP-MS) has proven to be a powerful approach to study protein-protein interactions (PPIs) in their native environment in cells and tissues1,2. This method allows for the identification of direct or indirect interactions at specific developmental stages or tissue contexts, and has been successfully used to identify multiple novel PPIs in a variety of developmental pathways (reviewed in reference1). Despite the undisputed success of PPI studies, most of them have been carried out in cultured cells, in which the &#34;bait&#34; proteins of interest were overexpressed. There are two issues with studying PPIs in cell culture: first, a specific cell line may not provide a full complement of interactions due to lack of expression of certain proteins. Second, high overexpression usually employed in such analyses might lead to artefacts such as protein misfolding or identification of false positive interactions.
Both of these limitations can be overcome by analyzing PPIs in vivo. A limiting step in such experiments is the availability of the starting material for purifying protein complexes. Drosophila melanogaster has long been used as a model for functional analysis, and recently it has also been shown to be an excellent system for studying PPIs in vivo. Drosophila embryogenesis represents a particularly attractive tissue type to study PPIs, because embryos can be easily collected in large quantities, and also because most genes (&#62;88%) are expressed over the course of embryogenesis, thus providing a rich in vivo environment for detecting relevant PPIs3.
Traditionally, biochemical studies in flies utilized very large-scale embryo collections (100-150 g), such as those necessary for purifying functional transcriptional lysates4,5. Previous AP-MS studies in Drosophila also needed large amounts of embryos (5-10 g), because they relied on a two-step purification approach such as tandem affinity purification (TAP), with the associated loss of material at each step6. Large amounts of starting material necessitated setting up embryo collections in large population cages, which can be both expensive (when purchased commercially) and time-consuming to maintain and clean7,8,9.
Recent advances in the development of single-step affinity purification approaches, as well as the increasing sensitivity of mass spectrometers, have reduced the necessary amount of starting material by an order of magnitude. Using tags such as the streptavidin-binding peptide (SBP) or green fluorescent protein (GFP) and starting from less than 1 g of embryos, it is possible to isolate the amounts of the bait protein and interacting components that would be sufficient for identification by mass spectrometry10,11.
The goal of the protocol presented here is to help the researchers overcome a perceived barrier to biochemical analysis of PPIs in vivo. To that end, we provide a simple and inexpensive procedure to collect Drosophila embryos at medium scale (0.5-1 g), followed by one-step preparation of whole-cell protein extracts that are suitable for subsequent analysis by AP-MS or other approaches. Our method relies on the use of custom-made 1-L or 5-L population cages that can be easily produced by any laboratory. Furthermore, the extraction conditions presented here have been validated in several studies, both in cultured cells and in vivo10,12,13,14,15,16,17.

<table border="0" cellpadding="0" cellspacing="0" width="1384">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Leaktite 5-Qt. Natural Multi Mix Container (Pack of 3)</td>
			<td style="width:183px;">Home Depot</td>
			<td style="width:119px;">209330</td>
			<td style="width:644px;">For making 5-L cages.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Leaktite 5-Qt. Natural Multi Mix Lid (Pack of 3)</td>
			<td style="width:183px;">Home Depot</td>
			<td style="width:119px;">209320</td>
			<td style="width:644px;">Diameter 8.37 in. Lids for 5-L cages.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Fisherbrand Petri dishes with clear lid, 100 x 15 mm</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">FB0875712</td>
			<td style="width:644px;">Material: Polystyrene. To be used with 1-L cages.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Fisherbrand Petri dishes with clear lid, 150 x 15 mm</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">FB0875714</td>
			<td style="width:644px;">Material: Polystyrene. To be used with 5-L cages.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Fisherbrand Petri dishes with clear lid, 60 x 15 mm</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">FB0875713A</td>
			<td style="width:644px;">Material: Polystyrene. Used during embryo dechorionation.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Sefar NITEX nylon mesh</td>
			<td style="width:183px;">Sefar</td>
			<td style="width:119px;">03-180/44</td>
			<td style="width:644px;">180 micron, 44% open area, used for fly cages.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Milwaukee 3 in. Hole Dozer Hole Saw with Arbor</td>
			<td style="width:183px;">Home Depot</td>
			<td style="width:119px;">49-56-9670</td>
			<td style="width:644px;">3 inch cutting tool for making 1-L cages.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:439px;">Nalgene Wide-Mouth Straight-Sided PMP Jars with White Polypropylene Screw Closure</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">11-823-33</td>
			<td style="width:644px;">For making 1-L cages. Thermo Scientific cat # 21171000. Alternative 1-L containers can be used.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Red Star Active Dry Yeast, 2 pound pouch</td>
			<td style="width:183px;">Red Star&#160;</td>
			<td style="width:119px;"></td>
			<td style="width:644px;">can be purchased from Amazon.com or other suppliers.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Frozen 100% Apple Juice Concentrate&#160;</td>
			<td style="width:183px;"></td>
			<td style="width:119px;"></td>
			<td style="width:644px;">can be purchased from a grocery store. Has to say &#34;100% apple juice&#34; on the can.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Methyl 4-hydroxybenzoate</td>
			<td style="width:183px;">Acros Organics</td>
			<td style="width:119px;">AC126965000</td>
			<td style="width:644px;">also known as methylparaben or Tegosept. Used as a preservative in AJ plates.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">IGEPAL CA-630 for molecular biology, 100 ml</td>
			<td style="width:183px;">Sigma</td>
			<td style="width:119px;">I8896</td>
			<td style="width:644px;">used for preparing lysis buffer.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:439px;">Nalgene Rapid-Flow Sterile Disposable Filter Units with CN Membrane&#160;</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">09-740-2A</td>
			<td style="width:644px;">0.2 &#956;m pore size. Used for filtering 5x lysis buffer. Thermo Scientific cat # 1260020.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">CO-RO ROCHE cOmplete Protease Inhibitor Cocktail</td>
			<td style="width:183px;">Sigma</td>
			<td style="width:119px;">11697498001</td>
			<td style="width:644px;">Vial of 20 tablets. Used for protease inhibition in lysis buffer.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Corning SFCA syringe filters&#160;</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">09-754-21</td>
			<td style="width:644px;">SFCA membrane, diameter 26 mm, pore size 0.45 &#956;m. Used for filtering final extract samples.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">BD&#160;Luer-Lok Disposable Syringes without Needles</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">14-823-2A</td>
			<td style="width:644px;">for filtering final extract samples. BD cat # 309604.</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:439px;">Bleach</td>
			<td style="width:183px;">Clorox</td>
			<td style="width:119px;"></td>
			<td style="width:644px;">used for embryo dechorionation at 50% (vol/vol) in water, can be purchased at any supermarket. It is important to use the Clorox brand, as other brands may result in incomplete dechorionation or may be toxic for embryos.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Corning Costar Netwell Plates</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">07-200-213</td>
			<td style="width:644px;">mesh containers for embryo collections. 74 &#956;m mesh size, 6-well.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:439px;">Wheaton Dounce Tissue Grinders, capacity 15 ml</td>
			<td style="width:183px;">Fisher Scientific</td>
			<td style="width:119px;">06-435B</td>
			<td style="width:644px;">homogenizer for embryos, with loose and tight pestles. Wheaton cat # 357544.</td>
		</tr>
	</tbody>
</table>

medium-scale preparation of drosophila embryo extracts for proteomic experiments

Analysis of protein-protein interactions (PPIs) has become an indispensable approach to study biological processes and mechanisms, such as cell signaling, organism development, and disease. It is often desirable to obtain PPI information using in vivo material, to gain the most natural and unbiased view of the interaction networks. The fruit fly Drosophila melanogaster is an excellent platform to study PPIs in vivo, and lends itself to straightforward approaches to isolating material for biochemical experiments. In particular, fruit fly embryos represent a convenient type of tissue to study PPIs, due to the ease of collecting animals at this developmental stage and the fact that the majority of proteins are expressed in embryogenesis, thus providing a relevant environment to reveal most PPIs. Here we present a protocol for collection of Drosophila embryos at medium scale (0.5-1 g), which is an ideal amount for a wide range of proteomic applications, including analysis of PPIs by affinity purification-mass spectrometry (AP-MS). We describe our designs for 1 L and 5 L cages for embryo collections that can be easily and inexpensively set up in any laboratory. We also provide a general protocol for embryo collection and protein extraction to generate lysates that can be directly used in downstream applications such as AP-MS. Our goal is to provide an accessible means for all researchers to carry out the analyses of PPIs in vivo.

To illustrate the use of this protocol in a protein complex purification experiment, we generated a homozygous viable fly line, arm-EGFP-ERK, expressing EGFP-tagged Drosophila extracellular signal-regulated kinase (ERK, encoded by the rolled gene) under the control of a ubiquitously expressed armadillo (arm) promoter18,19. The arm promoter is active during most stages of Dr...

Watch this Scientific Journal Video about Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments at JoVE.com

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

The goal of this protocol is to provide a straightforward and inexpensive approach to collecting Drosophila embryos at medium scale (0.5-1 g) and preparing protein extracts that can be used in downstream proteomic applications, such as affinity purification-mass spectrometry (AP-MS).

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

Analysis of protein-protein interactions (PPIs) has become an indispensable approach to study biological processes and mechanisms, such as cell signaling, ...

medium-scale-preparation-drosophila-embryo-extracts-for-proteomic

Research

JoVE Journal

Developmental Biology

7.8K Views.  University of Massachusetts Boston. The goal of this protocol is to provide a straightforward and inexpensive approach to collecting Drosophila embryos at medium scale (0.5-1 g) and preparing protein extracts that can be used in downstream proteomic applications, such as affinity purification-mass spectrometry (AP-MS).

Video: Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Since mitochondria play roles in amino acid metabolism, carbohydrate metabolism and fatty acid oxidation, defects in mitochondrial function often compromise the lives of those who suffer from these complex diseases. Detecting mitochondrial metabolic changes is vital to the understanding of mitochondrial disorders and mitochondrial responses to pharmacological agents. Although mitochondrial metabolism is at the core of metabolic regulation, the detection of subtle changes in mitochondrial metabolism may be hindered by the overrepresentation of other cytosolic metabolites obtained using whole organism or whole tissue extractions. 
Here we describe an isolation method that detected pronounced mitochondrial metabolic changes in Drosophila that were distinct between whole-fly and mitochondrial enriched preparations. To illustrate the sensitivity of this method, we used a set of Drosophila harboring genetically diverse mitochondrial DNAs (mtDNA) and exposed them to the drug rapamycin. Using this method we showed that rapamycin modifies mitochondrial metabolism in a mitochondrial-genotype-dependent manner. However, these changes are much more distinct in metabolomics studies when metabolites were extracted from mitochondrial enriched fractions. In contrast, whole tissue extracts only detected metabolic changes mediated by the drug rapamycin independently of mtDNAs.

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Mitochondria play central roles in the regulation of metabolism and homeostasis. Subtle changes in mitochondrial metabolism that affect organismal physiology could be difficult to detect in whole organism metabolomics studies. Here we describe an isolation method that enhances the detection of subtle metabolic shifts in Drosophila melanogaster.

Since mitochondria play roles in amino acid metabolism, carbohydrate metabolism and fatty acid oxidation, defects in mitochondrial function often ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout (Oncorhynchus Mykiss) and Brown Trout (Salmo Trutta Morpha fario)

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on fish abdomens to expel the eggs. An alternative method uses the pressure of gas injected into the body cavity, which causes the subsequent release of the eggs. This method is believed to have less negative effects on both the welfare and egg quality of the broodstocks. Herein, we compare the results of air and hand stripping methods with respect to one-year survival and egg quantity and quality in two salmonid fish, rainbow trout (Oncorhynchus mykiss) and brown trout (Salmo trutta morpha fario). Our results indicate that air stripping yielded a better quality of eggs and higher one-year survival rate in rainbow trout. In addition, air stripping resulted in lower mortality rate than the group subjected to hand stripping (25% vs. 35%). The pH and hatching rate of the hand stripped group was lower than those of the air stripped group. In the case of brown trout, the quality of eggs obtained by both hand and air-stripping methods was similar; however, the one-year losses in fish were higher in air stripped group (15% compared to 0% in hand stripped fish). Although the advantages of air stripping method over hand stripping in terms of egg quality might not be observed in all salmonid species, the air-stripping procedure might be a promising option to be adopted in hatcheries as it ensures a high level of reproducibility and efficiency.

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout Oncorhynchus Mykiss and Brown Trout Salmo Trutta Morpha fario

The principal aim of this study is to standardize and test the pneumatic method (air stripping) of collecting eggs in rainbow trout and brown trout. This method allows effective and simple collection of the eggs without the necessity of fish abdomen massage.

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on ...

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

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

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

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

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

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to understand morphogenetic processes is to directly measure cellular forces and mechanical properties in vivo during embryogenesis. Here, we present a setup of optical tweezers coupled to a light sheet microscope, which allows to directly apply forces on cell-cell contacts of the early Drosophila embryo, while imaging at a speed of several frames per second. This technique has the advantage that it does not require the injection of beads into the embryo, usually used as intermediate probes on which optical forces are exerted. We detail step by step the implementation of the setup, and propose tools to extract mechanical information from the experiments. By monitoring the displacements of cell-cell contacts in real time, one can perform tension measurements and investigate cell contacts&#39; rheology.

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

We present a setup of optical tweezers coupled to a light sheet microscope, and its implementation to probe cell mechanics without beads in the Drosophila embryo.

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to ...

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal and postnatal consequences. To ensure the innocuousness of ART applications, studies in animal models are necessary. In addition, as a last step, embryo development studies require evaluation of their capacity to develop full-term healthy offspring. Here, embryo transfer to the uterus is indispensable to perform any ARTs-related experiment.
The rabbit has been used as a model organism to study mammalian reproduction for over a century. In addition to its phylogenetic proximity to the human species and its small size and low maintenance cost, it has important reproductive characteristics such as induced ovulation, a chronology of early embryonic development similar to humans and a short gestation that allow us to study the consequences of ART application easily. Moreover, ARTs (such as intracytoplasmic sperm injection, embryo culture, or cryopreservation) are applied with suitable efficiency in this species.
Using the laparoscopic embryo transfer technique and the cryopreservation protocol presented in this article, we describe 1) how to transfer embryos through an easy, minimally invasive technique and 2) an effective protocol for long-term storage of rabbit embryos to provide time-flexible logistical capacities and the ability to transport the sample. The outcomes obtained after transferring rabbit embryos at different developmental stages indicate that morula is the ideal stage for rabbit embryo recovery and transfer. Thus, an oviductal embryo transfer is required, justifying the surgical procedure. Furthermore, rabbit morulae are successfully vitrified and laparoscopically transferred, proving the effectiveness of the described techniques.

Assisted reproductive techniques (ARTs) are in continuous evaluation to improve outcomes and reduce the associated risks. This manuscript describes a minimally invasive embryo transfer procedure with an efficient cryopreservation protocol that allows the use of rabbits as an ideal animal model of human reproduction.

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

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

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

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

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

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

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Microbead Implantation in the Zebrafish Embryo

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Imaging Subcellular Structures in the Living Zebrafish Embryo

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout (Oncorhynchus Mykiss) and Brown Trout (Salmo Trutta Morpha fario)

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

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines