We thank Susan Gerbi, Heidi Smith, and David Lambert for supplying material to test in-vivo centrifugation on Sciara coprophila and Ilyanassa obsoleta, respectively. We thank members of the Welte lab for comments on the technique and manuscript. Development and refinement of the in-vivo centrifugation assay was supported by NIGMS grant GM64687 to MAW. The images shown in Fig. 1 and Fig. 5E,F were previously published2 and are reprinted here with permission.

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	<li>Wieschaus, E., N&#252;sslein-Volhard, C., Roberts, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Looking+at+embryos.">Looking at embryos.</a> Drosophila: A Practical Approach. , 179-214 (1998).</li><li>Cermelli, S., Guo, Y., Gross, S. P., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+lipid+droplet+proteome+reveals+that+droplets+are+a+protein+storage+depot.">The lipid droplet proteome reveals that droplets are a protein storage depot.</a> Curr Biol. 16, 1783-1795 (2006).</li><li>Kiehart, D. P., Crawford, J. M., Montague, R. A., Sullivan, W., Ashburner, M., Hawley, R. 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T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Consequences+of+motor+copy+number+on+the+intracellular+transport+of+kinesin-1-driven+lipid+droplets.">Consequences of motor copy number on the intracellular transport of kinesin-1-driven lipid droplets.</a> Cell. 135, 1098-1107 (2008).</li><li>Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+lipid-droplet+transport+by+the+Perilipin+homologue+LSD2.">Regulation of lipid-droplet transport by the Perilipin homologue LSD2.</a> Curr. Biol. 15, 1266-1275 (2005).</li><li>Phadnis, N., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unpublished+Observations.">Unpublished Observations.</a> , .</li><li>Phadnis, N., Welte, M. 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No conflicts of interest declared.

Critical steps
Make sure that the agar concentration is high enough. If it is too low, the agar will disintegrate during centrifugation, and the embryos will burst or be randomly oriented. If the agar concentration is just slightly too low or if the agar is wet (insufficiently dried or insufficient removal of excess TSS in step 4.3), the embryos will float out of their holes during centrifugation and become improperly oriented. If the agar is poured too thinly, it will also collapse or crack during centrifugation.
Make sure that embryos have not advanced beyond cellularization stages. Otherwise, separation of organelles will not work (Fig. 5C) since organelles will become sequestered within thousands of individual cells. In younger embryos at the one cell stage, organelles are free to migrate large distances and accumulate according to their characteristic density. To avoid analyzing embryos that are too old, make sure to (i) include a pre-collection to allow females to lay old eggs (step 2.4), (ii) collect embryos for 3 hours or less (step 2.5), (iii) visually select embryos before cellularization stages for embedding into agar (step 4.2), (iv) keep embedding time short to prevent development through the end of cellularization (step 4.6), and (v) discard embryos that did not layer correctly (step 5.6).
Do not skimp on the centrifugation time. Although some of the embryos will show nice layering even after only 10 minutes of centrifugation, the separation is inconsistent from embryo to embryo. 30 min spins give highly consistent results. Also, the longer the centrifugation time, the longer the layers are stable after the centrifugation has ended. This is important for those applications in which it takes time to process the embryos further before use (e.g., if they need to be processed for transplantation experiments or mounted for confocal analysis).
Possible modifications
Egg collection and chorion removal Many different protocols exist for steps 2 and 3 1, 3, 9, and will work as well as the ones described here as long as the embryos used are young, egg yield is high, and the fraction of mis-staged embryos is minimized.
Anterior-posterior alignment The protocol orients all embryos such that the posterior end is buried in the agar and the anterior end is up (Fig. 3). This orientation is good for consistency, so that it is possible to use the micropyle as a landmark which end pointed up during centrifugation (Fig. 1, 5). However, with practice, it is easy to pick out the orientation of centrifuged embryos by the distinct appearance of the lipid-droplet and yolk layers by bright-field microscopy. Then it becomes possible to mount embryos with either end up; this flexibility speeds up the mounting process.
Unoriented centrifugation The most-time-consuming and technically challenging aspect of the protocol is mounting the embryos in agar (step 4). It requires touching each embryo twice (once to insert it into the hole and once to recover it after centrifugation). As an alternative, one can transfer embryos at step 4.1 into microcentrifuge tubes filled with TSS. When these tubes are centrifuged in a microcentrifuge (13,000 rpm, 10-30 min), embryos will also typically layer well 10-13. This variation allows one to process hundreds of embryos at the same time very quickly. However, layering is not as consistent; a fraction of the embryos does not develop distinct layers despite much higher centrifugation forces (~16,000 g) than applied in the protocol above. More challenging is the fact that the orientation of the embryos is variable, and one can observe separation at various angles to the major embryo axis. This variability requires the experimenter to sort out after the centrifugation how a particular embryo was arranged in the centrifuge tube. With the use of the yolk autofluorescence as a marker for the most-dense part of the embryo, this is often possible, though it requires much practice and more judgment to do it reliably. If there are sufficient embryos in the sample, one may be able to sort out those instances in which embryos happened to be arranged like in Fig. 1.
Centrifugation of embryos in their egg shell It is possible to embed embryos in agar for step 4 without removing the chorion first 2. In this approach, embryos on the collection plate are covered with halocarbon oil instead of 50% bleach in step 3.2 (halocarbon turns the chorion translucent). Embryos of appropriate stages are selected and transferred to a new agar plate using tweezers; by grabbing only the egg filaments with the tweezers, damage to the embryo proper is avoided. Embryos are embedded as in step 4.4 through 4.8, with the anterior egg filaments sticking up out of the hole in the agar. The egg shell is removed by 50% bleach treatment after the embryos have been centrifuged and dug out of the agar as in step 5.4. Centrifugation in the presence of the egg shell may improve the rate of devitellinization after fixation (step 6.2), but selection of the correct stages for centrifugation (step 4.2) is more challenging.
Applications
Embryo centrifugation is particularly useful for colocalization experiments, i.e., to test if a particular protein localizes to a certain major organelle. For example, the Klar protein is present on early embryonic lipid droplets, yet this localization is challenging to demonstrate in intact embryos. In part, this is because both Klar puncta and lipid droplets are abundant in the embryo periphery 10, making spurious colocalization hard to rule out. However, centrifugation concentrates lipid droplets into a distinct layer, and thus, colocalization with Klar signal becomes obvious 10.
Similar analysis has unambiguously demonstrated the localization of other proteins to lipid droplets 8, 13-15, including surprising examples like certain histones 2 and in cases where a protein is present ubiquitously but enriched on lipid droplets 8. As embryos can be selected visually by developmental stage (step 4.2), it is even possible to detect developmentally regulated recruitment of proteins to lipid droplets 8, 15. Testing for colocalization with lipid droplets is particularly easy since lipid droplets accumulate in a visually distinct "cap", so no further marker or stain is required to identify this layer. In fact, because of its ease and potentially striking visual results (Fig. 1, Fig. 5 D, E), our laboratory employs this technique now routinely as a first test whether a candidate protein is localized to lipid droplets. In principle, similar colocalization studies should work for the other organelles shown in Fig. 1, and likely for others (including intracellular parasites) whose distribution in centrifuged embryos has yet to be documented.
Since centrifugation concentrates organelles into tight bands, it facilitates isolation of a fraction enriched in these organelles. This approach has already been used to recover lipid droplets for transplantation into donor embryos 2. It may also be possible to biochemically analyze the enriched fraction (e.g., by cutting layers off embryos after fixation 15).
The method described here has applications beyond Drosophila embryos, as centrifugation can be used to stratify the eggs of many different species (including frogs, nematodes, tunicates, annelids, sea urchins, mollusks, and Cnidarians) 16. In our own laboratory, we have used the protocol above for embryos of the housefly Musca domestica 2 and have employed unoriented centrifugation in the analysis of embryos from the fungus gnat Sciara coprophila 17 and the snail Ilyanassa obsoleta 17. Finally, this method is not restricted to embryos: Centrifugation can also be useful to stratify other large cells, such as oocytes (e.g., in Drosophila 2, 18 and the milkweed bug Oncopeltus fasciatus 17). For different specimen, the exact centrifugation conditions need to be optimized: For example, 10 minutes at 3000 g is sufficient to efficiently stratify embryos of Musca domestica 2, and if centrifuged much longer, the embryos tend to break apart during fixation and immunostaining.

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		<th>Catalogue Number</th>
		<th>Comment</th>
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<td>60x15 mm Petri dishes</td>
<td>&nbsp;</td>
<td>Becton Dickinson</td>
<td>#35-1007</td>
<td>From Falcon</td>
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<tr>
<td>Wire mesh</td>
<td>&nbsp;</td>
<td>Small Parts</td>
<td>CX-0150</td>
<td>stainless steel type 304; mesh size #150 x150, wire diameter: 0.0026 inches</td>
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<td>Tungsten needles</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>10130-10</td>
<td>0.25 mm</td>
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<td>Moria Nickel Plated Pin/Needle Holder</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>26016-12</td>
<td>&nbsp;</td>
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<td>Fly Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific</td>
<td>59-100 or 59-101</td>
<td>For 65 mm or 100 mm diameter Petri dishes, respectively</td>
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<td>Halocarbon Oil 27</td>
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in-vivo centrifugation of drosophila embryos

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

Watch this Scientific Journal Video about In-vivo Centrifugation of Drosophila Embryos at JoVE.com

In-vivo Centrifugation of Drosophila Embryos

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

In-vivo Centrifugation of Drosophila Embryos

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

in-vivo-centrifugation-of-drosophila-embryos

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18.3K Views.  University of Rochester. We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.