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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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 membrane and soluble nuclear proteins6,10,12,15. It can be easily adapted for a more focused purification of proteins from other compartments, such as isolation of less soluble nuclear proteins using high salt extraction5. As mentioned in step 5.8, collection times and embryo aging periods can be set up in precise intervals to obtain different developmental stages.
It is often desirable to ascertain that the tagged protein of interest is expressed at levels closely matching the endogenous levels of expression, as we have shown using anti-total ERK antibody (Figure 2A). If the antibody against endogenous protein is not available, the functionality of the constructs can be verified by other assays, e.g. in a genetic rescue experiment. In most cases however, a mild overexpression of the tagged protein should still result in isolation of meaningful protein complexes, and may in fact facilitate extraction and purification of &#34;difficult&#34; proteins.
The main advantage of this protocol is its simplicity and a possibility to set up the whole procedure at moderate cost. An additional advantage of using smaller cages is that multiple transgenic lines can be set up and analyzed in parallel, which may not be feasible when using larger cages. The fly cages we describe here can be easily cleaned and reused for years. Instead of making 1-L cages from Nalgene jars as described, it is possible to use other available containers, such as a smaller version of the 5-L container.
We have successfully used this protocol (or its variations with small modifications) for purifying complexes containing various signaling proteins and associated components, followed by analysis by mass spectrometry6,10,15. These approaches have led to the identification of novel PPIs that were further validated in functional studies10,15. Variations of the protocol presented here have been previously used to analyze the phosphoproteome in Drosophila embryogenesis20, and to study the landscape of ubiquitination in neural development21,22. This procedure therefore represents a convenient gateway to analyzing PPIs in vivo, and is usable for other proteomics applications.

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><tbody><tr><td>Leaktite 5-Qt. Natural Multi Mix Container (Pack of 3)</td><td>Home Depot</td><td>209330</td><td>For making 5-L cages.</td></tr><tr><td>Leaktite 5-Qt. Natural Multi Mix Lid (Pack of 3)</td><td>Home Depot</td><td>209320</td><td>Diameter 8.37 in. Lids for 5-L cages.</td></tr><tr><td>Fisherbrand Petri dishes with clear lid, 100 x 15 mm</td><td>Fisher Scientific</td><td>FB0875712</td><td>Material: Polystyrene. To be used with 1-L cages.</td></tr><tr><td>Fisherbrand Petri dishes with clear lid, 150 x 15 mm</td><td>Fisher Scientific</td><td>FB0875714</td><td>Material: Polystyrene. To be used with 5-L cages.</td></tr><tr><td>Fisherbrand Petri dishes with clear lid, 60 x 15 mm</td><td>Fisher Scientific</td><td>FB0875713A</td><td>Material: Polystyrene. Used during embryo dechorionation.</td></tr><tr><td>Sefar NITEX nylon mesh</td><td>Sefar</td><td>03-180/44</td><td>180 micron, 44% open area, used for fly cages.</td></tr><tr><td>Milwaukee 3 in. Hole Dozer Hole Saw with Arbor</td><td>Home Depot</td><td>49-56-9670</td><td>3 inch cutting tool for making 1-L cages.</td></tr><tr><td>Nalgene Wide-Mouth Straight-Sided PMP Jars with White Polypropylene Screw Closure</td><td>Fisher Scientific</td><td>11-823-33</td><td>For making 1-L cages. Thermo Scientific cat # 21171000. Alternative 1-L containers can be used.</td></tr><tr><td>Red Star Active Dry Yeast, 2 pound pouch</td><td>Red Star&amp;nbsp;</td><td></td><td>can be purchased from Amazon.com or other suppliers.</td></tr><tr><td>Frozen 100% Apple Juice Concentrate&amp;nbsp;</td><td></td><td></td><td>can be purchased from a grocery store. Has to say &quot;100% apple juice&quot; on the can.</td></tr><tr><td>Methyl 4-hydroxybenzoate</td><td>Acros Organics</td><td>AC126965000</td><td>also known as methylparaben or Tegosept. Used as a preservative in AJ plates.</td></tr><tr><td>IGEPAL CA-630 for molecular biology, 100 ml</td><td>Sigma</td><td>I8896</td><td>used for preparing lysis buffer.</td></tr><tr><td>Nalgene Rapid-Flow Sterile Disposable Filter Units with CN Membrane&amp;nbsp;</td><td>Fisher Scientific</td><td>09-740-2A</td><td>0.2 &amp;mu;m pore size. Used for filtering 5x lysis buffer. Thermo Scientific cat # 1260020.</td></tr><tr><td>CO-RO ROCHE cOmplete Protease Inhibitor Cocktail</td><td>Sigma</td><td>11697498001</td><td>Vial of 20 tablets. Used for protease inhibition in lysis buffer.</td></tr><tr><td>Corning SFCA syringe filters&amp;nbsp;</td><td>Fisher Scientific</td><td>09-754-21</td><td>SFCA membrane, diameter 26 mm, pore size 0.45 &amp;mu;m. Used for filtering final extract samples.</td></tr><tr><td>BD&amp;nbsp;Luer-Lok Disposable Syringes without Needles</td><td>Fisher Scientific</td><td>14-823-2A</td><td>for filtering final extract samples. BD cat # 309604.</td></tr><tr><td>Bleach</td><td>Clorox</td><td></td><td>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><td>Corning Costar Netwell Plates</td><td>Fisher Scientific</td><td>07-200-213</td><td>mesh containers for embryo collections. 74 &amp;mu;m mesh size, 6-well.</td></tr><tr><td>Wheaton Dounce Tissue Grinders, capacity 15 ml</td><td>Fisher Scientific</td><td>06-435B</td><td>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 Drosophila embryogenesis19. Lysates were prepared using the described protocol, and the expression of protein from the arm-EGFP-ERK transgene was confirmed by western blotting for total ERK protein to simultaneously detect the levels of endogenous and transgenic EGFP-ERK, migrating at 42 kDa and 69 kDa, respectively (Figure 2A). A single band corresponding to untagged endogenous ERK (42 kDa) was detected in the y w control line. This result confirms a successful extraction of ERK and EGFP-ERK from embryos.
To test whether our extraction protocol is suitable for subsequent purification of a tagged protein, extracts from the y w and arm-EGFP-ERK flies were subjected to affinity purification using GFP-agarose beads following established protocols10,11. Silver staining of samples run on a gradient SDS-PAGE showed a successful purification of the bait protein, EGFP-ERK (Figure 2B). Besides EGFP-ERK itself, the experimental lane contained additional bands that were not observed in the control y w sample, suggesting that the purification procedure recovered potential ERK-interacting proteins.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55804/55804fig1.jpg" /> 
Figure 1: Preparation of Population Cages. (A-D) A 5-L cage made from a plastic container. (A) A starting container. (B) Bottom view of the 5 L cage. (C) Top view of 5 L cage with mesh attached. (D) Assembled 5 L cage with a 15 cm apple juice plate, bottom view. (E and F) A 3 inch (76 mm) drill bit used for drilling holes in the 1 L container. (G, H) A 1 L cage. (G) Top view of a 1 L cage with mesh attached. (H) Assembled 1 L cage with a 10 cm apple juice plate attached, bottom view. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55804/55804fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/55804/55804fig2.jpg" /> 
Figure 2: Extraction and Purification of EGFP-ERK from Embryos. (A) Western blot showing expression of EGFP-ERK and endogenous ERK in an extract prepared from the arm-EGFP-ERK transgenic line. The y w line was used as a control. Green, total ERK antibody; red, molecular weight marker. (B) Silver stained gel showing purified EGFP-ERK and associated proteins. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55804/55804fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

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

Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a protocol for performing the chromatin conformation capture technique in situ Hi-C on staged Drosophila melanogaster embryo populations. The result is a sequencing library that allows the mapping of all chromatin interactions that occur in the nucleus in a single experiment. Embryo sorting is done manually using a fluorescent stereo microscope and a transgenic fly line containing a nuclear marker. Using this technique, embryo populations from each nuclear division cycle, and with defined cell cycle status, can be obtained with very high purity. The protocol may also be adapted to sort older embryos beyond gastrulation. Sorted embryos are used as inputs for in situ Hi-C. All experiments, including sequencing library preparation, can be completed in five days. The protocol has low input requirements and works reliably using 20 blastoderm stage embryos as input material. The end result is a sequencing library for next generation sequencing. After sequencing, the data can be processed into genome-wide chromatin interaction maps that can be analyzed using a wide range of available tools to gain information about topologically associating domain (TAD) structure, chromatin loops, and chromatin compartments during Drosophila development.

Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

This work describes a protocol for the generation of high resolution in situ Hi-C libraries from tightly staged pre-gastrulation Drosophila melanogaster embryos.

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a ...

Genetics

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 ...

Biology

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 ...

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Calcium induced calcium release signaling (CICR) plays a critical role in many biological processes. Every cellular activity from cell proliferation and apoptosis, development and ageing, to neuronal synaptic plasticity and regeneration have been associated with Ryanodine receptors (RyRs). Despite the importance of calcium signaling, the exact mechanism of its function in early development is unclear. As an organism with a short gestational period, the embryos of Drosophila melanogaster are prime study subjects for investigating the distribution and localization of CICR associated proteins and their regulators during development. However, because of their lipid-rich embryos and chitin-rich chorion, their utility is limited by the difficulty of mounting embryos on glass surfaces. In this work, we introduce a practical protocol that significantly enhances the attachment of Drosophila embryo onto slides and detail methods for successful histochemistry, immunohistochemistry, and in-situ hybridization. The chrome alum gelatin slide-coating method and embryo pre-embedding method dramatically increases the yield in studying Drosophila embryo protein and RNA expression. To demonstrate this approach, we studied DmFKBP12/Calstabin, a well-known regulator of RyR during early embryonic development of Drosophila melanogaster. We identified DmFKBP12 in as early as the syncytial blastoderm stage and report the dynamic expression pattern of DmFKBP12 during development: initially as an evenly distributed protein in the syncytial blastoderm, then preliminarily localizing to the basement layer of the cortex during cellular blastoderm, before distributing in the primitive neuronal and digestion architecture during the three-gem layer phase in early gastrulation. This distribution may explain the critical role RyR plays in the vital organ systems that originate in from these layers: the suboesophageal and supraesophageal ganglion, ventral nervous system, and musculoskeletal system.

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Here, we describe a protocol for detection and localization of Drosophila embryo protein and RNA from collection to pre-embedding and embedding, immunostaining, and mRNA in situ hybridization.

Calcium induced calcium release signaling (CICR) plays a critical role in many biological processes. Every cellular activity from cell proliferation and ...

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged molecules within complex biological samples. While traditional confocal microscopes allow one to resolve two adjacent fluorescent point sources located a few hundred nanometers apart, observing the finer details of subcellular biology requires the ability to resolve signals in the order of tens of nanometers. Numerous hardware-based methods for super-resolution microscopy have been developed to allow researchers to sidestep such resolution limits, although these methods require specialized microscopes that are not available to all researchers. An alternative method for increasing resolving power is to isotropically enlarge the sample itself through a process known as expansion microscopy (ExM), which was first described by the Boyden group in 2015. ExM is not a type of microscopy per se but is rather a method for swelling a sample while preserving the relative spatial organization of its constituent molecules. The expanded sample can then be observed at an effectively increased resolution using a traditional confocal microscope. Here, we describe a protocol for implementing ExM in whole-mount Drosophila embryos, which is used to examine the localization of Par-3, myosin II, and mitochondria within the surface epithelial cells. This protocol yields an approximately four-fold increase in sample size, allowing for the detection of subcellular details that are not visible with conventional confocal microscopy. As proof of principle, an anti-GFP antibody is used to distinguish distinct pools of myosin-GFP between adjacent cell cortices, and fluorescently labeled streptavidin is used to detect endogenous biotinylated molecules to reveal the fine details of the mitochondrial network architecture. This protocol utilizes common antibodies and reagents for fluorescence labeling, and it should be compatible with many existing immunofluorescence protocols.

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

Here, a protocol for the implementation of expansion microscopy in early Drosophila embryos to achieve super-resolution imaging using a conventional laser-scanning confocal microscope is presented.

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged ...

Enhancing Protein Visualization with Antibody-Mediated Live Imaging

Visualizing Low-Abundance Proteins and Post-Translational Modifications in Living Drosophila Embryos via Fluorescent Antibody Injection

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein localization, dynamics, and function. Compared to immunofluorescence, live imaging more accurately reflects protein localization without potential artifacts arising from tissue fixation. Importantly, live imaging enables quantitative and temporal characterization of protein levels and localization, crucial for understanding dynamic biological processes such as cell movement or division. However, a major limitation of fluorescent tagging approaches is the need for sufficiently high protein expression levels to achieve successful visualization. Consequently, many endogenously tagged fluorescent proteins with relatively low expression levels cannot be detected. On the other hand, ectopic expression using viral promoters can sometimes lead to protein mislocalization or functional alterations in physiological contexts. To address these limitations, an approach is presented that utilizes highly sensitive antibody-mediated protein detection in living embryos, essentially performing immunofluorescence without the need for tissue fixation. As proof of principle, endogenously GFP-tagged Notch receptor that is barely detectable&#160;in living embryos can be successfully visualized after antibody injection. Furthermore, this approach was adapted to visualize post-translational modifications (PTMs) in living embryos, allowing the detection of temporal changes in tyrosine phosphorylation patterns during early embryogenesis and revealing a novel subpopulation of phosphotyrosine (p-Tyr) underneath apical membranes. This approach can be modified to accommodate other protein-specific, tag-specific, or PTM-specific antibodies and should be compatible with other injection-amenable model organisms or cell lines. This protocol opens new possibilities for live imaging of low-abundance proteins or PTMs that were previously challenging to detect using traditional fluorescent tagging methods.

Author Spotlight: Unveiling the Dynamics of Mechanical and Biochemical Signals in Animal Morphogenesis Research 

This protocol describes the customized antibody-based fluorescence labeling and injection into early Drosophila embryos to enable live imaging of low-abundance proteins or post-translational modifications that are challenging to detect using traditional GFP/mCherry-tag approaches.

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein ...

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons extend growth cones to navigate along specific pathways by responding to a large number of axon guidance cues that emanate from the surrounding extracellular environment. Growth cone target recognition also plays a critical role in neuromuscular specificity. This work presents a standard immunohistochemistry protocol to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. This protocol includes a few key steps, including a genotyping procedure, to sort the desired mutant embryos; an immunostaining procedure, to tag embryos with fasciclin II (FasII) antibody; and a dissection procedure, to generate filleted preparations from fixed embryos. Motor axon projections and muscle patterns in the periphery are much better visualized in flat preparations of filleted embryos than in whole-mount embryos. Therefore, the filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition, and it can also be applied to both loss-of-function and gain-of-function genetic screens.

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons ...

Neuroscience

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Assessing the expression pattern of a gene, as well as the subcellular localization properties of its transcribed RNA, are key features for understanding its biological function during development. RNA in situ hybridization (RNA-ISH) is a powerful method used for visualizing RNA distribution properties, be it at the organismal, cellular or subcellular levels 1. RNA-ISH is based on the hybridization of a labeled nucleic acid probe (e.g. antisense RNA, oligonucleotides) complementary to the sequence of an mRNA or a non-coding RNA target of interest 2. As the procedure requires primary sequence information alone to generate sequence-specific probes, it can be universally applied to a broad range of organisms and tissue specimens 3. Indeed, a number of large-scale ISH studies have been implemented to document gene expression and RNA localization dynamics in various model organisms, which has led to the establishment of important community resources 4-11. While a variety of probe labeling and detection strategies have been developed over the years, the combined usage of fluorescently-labeled detection reagents and enzymatic signal amplification steps offer significant enhancements in the sensitivity and resolution of the procedure 12. Here, we describe an optimized fluorescent in situ hybridization method (FISH) employing tyramide signal amplification (TSA) to visualize RNA expression and localization dynamics in staged Drosophila embryos. The procedure is carried out in 96-well PCR plate format, which greatly facilitates the simultaneous processing of large numbers of samples.

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Here we describe a whole-mount fluorescent in situ hybridization (FISH) protocol for determining the expression and localization properties of RNAs expressed during embryogenesis in the fruit fly, Drosophila melanogaster.

Assessing  the expression pattern of a gene, as well as the subcellular localization  properties of its transcribed RNA, are key features for ...

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 ...

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

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Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

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

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

Visualizing Low-Abundance Proteins and Post-Translational Modifications in Living Drosophila Embryos via Fluorescent Antibody Injection

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

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