We thank Yixian Zheng (Carnegie Institution of Washington, Baltimore, MD) for the original protocol and assistance in initial setup and members of the Lei laboratory for critical reading of the manuscript. This work was funded by the Intramural Research Program of the National Institute of Diabetes and Digestive and Kidney Diseases.

1. Starting the Cycle: Seeding Embryos

NOTE: The cycle starts with 1.5 g of collected material (a mixture composed of embryos and/or some first instar larvae) from the previous cycle. This material will be placed in a plastic container (see table of specific materials/equipment) with an active yeast mixture until the pupal phase. The container will be closed after the introduction of the biological mixture of embryos and larvae with the lid to avoid allowing the larvae to escape. It is necessary to make holes into the lid in order to allow air circulation. To avoid having larvae escape through the holes, foam plugs are used. Finally, It is recommended to wear gloves and lab coat, not only in this section but also in the whole protocol, to avoid getting dirty or staining clothes with bleach.

<ol>
	<li>Set up the plastic container.
	<ol>
		<li>Make three square holes with a razor blade in the plastic container lid of approximately 2.2 cm. Add tape (3/4 inch labeling tape) to the four corners of each hole to ensure a tight fit for the foam plugs (50 x 55 mm, d x l), and then place one foam plug in each hole.</li>
		<li>Cover the inside of the plastic container with plastic wrap. Over this film, add a layer of cotton to cover the bottom of the plastic container. Tear the cotton with your fingers.</li>
	</ol>
	</li>
	<li>Prepare the fly food.
	<ol>
		<li>Add 333 ml of deionized water in a 500 ml beaker. While stirring the beaker with a magnetic bar, add 167 &#956;l of propionic acid and 1.08 ml of phosphoric acid, from the stock solutions (99.96% and 85%, respectively).</li>
		<li>Slowly add 77.5 g of active dry yeast, avoiding big clumps. After dissolving the dry yeast, add 38.8 g of sucrose. 
		NOTE: Sucrose should be the last ingredient to be added because immediately after adding this component, fermentation will begin.</li>
	</ol>
	</li>
	<li>Immediately after dissolving the sucrose, pour the food over the cotton, and make sure to cover the cotton evenly. Close the plastic container with the lid to avoid escaped flies within the laboratory contaminating the food.</li>
	<li>Resuspend 1.5 g of the harvested embryos (not dechorionated) from the previous cycle with 5 ml of 70% ethanol. Cut in half two filter papers and distribute the biological mixture evenly over the 4 pieces using a spatula or a transfer pipet with a wide tip (cut if necessary). Lay the filter papers on top of the soaked cotton and close the lid. Finally, incubate the plastic container at RT (24&#186;C) and humidity (35%) until the pupa stage. 
	NOTE: The plastic container should not be placed in a humid chamber, otherwise this will encourage the growth of bacteria.</li>
</ol>
2. Continuation of the Cycle: From Embryos to Flies.
NOTE: This part of the cycle goes from the embryos placed in the plastic container on day 1 until 9 days&#160;later when the adult flies will emerge from the pupae. During these 8 days nothing has to be done but monitoring that the embryos progress correctly to the next stages of the Drosophila life cycle until eclosion. If larvae start dying and turning black during this time, check that the foam plugs are not too tight and that adequate ventilation is provided. This period is a good time to clean the fly population cage from the previous cycle.
<ol>
	<li>Observe the embryos become 1st instar larvae around 24 hr after the adult female deposited them into the fly food (see step 3). The resulting larvae will feed from the fly food prepared in step 1 during 4 days, growing and molting twice into 2nd and 3rd instar larvae.</li>
	<li>Clean the fly population cage.
	<ol>
		<li>Place the cage at 4 &#186;C for at least 30 min to slow down the activity of the flies. Remove the net that is covering one side of the cage, and with a fast movement, cap the open side with a large biohazard plastic bag. Pour the contents of the cage into the bag.</li>
		<li>With the biohazard bag still covering one side of the cage, position the cage vertically over a sink with the biohazard bag side down. Carefully remove the plastic bag to prevent flies escaping and discard it in a biohazard waste container.</li>
		<li>Carefully open the upper net enough to create a small hole, rinse the inside of the cage with water, which will wash the flies into the sink. Slowly increase the size of the hole and eliminate all the flies inside the cage.</li>
		<li>Clean the cage and both nets with water and soap. While the nets are still wet secure them on both sides of the dry clean cage, and finally put inside a lab soaker paper, which will prevent the plastic film prepared in step 1.1.2 from sticking to the cage. Now the cage is ready for the next cycle.</li>
	</ol>
	</li>
	<li>Four days after the initial setup, observe the larvae pupate. The larvae will remain in this stage for 4 more days. 
	NOTE: During this phase the pupae will cover the entire cotton surface inside the plastic container.</li>
	<li>During the pupal phase and before the first flies emerge, open the plastic container and place the plastic film containing the cotton with all the pupae over the lab soaker paper inside the clean population cage. Close the net with a double knot and clean the plastic container and the lid for the next cycle. 
	NOTE: Pupae affixed to the lid should be discarded and destroyed by autoclaving or freezing before eclosion.</li>
</ol>
3. Adult Flies.
NOTE: After the 4 days of the pupa stage the first flies will emerge from the pupae. Within 24 - 48 hr all flies should have eclosed. In this part of the cycle, it is important to provide them food with the aim of creating the right environment for reproduction.
<ol>
	<li>Prepare the molasses trays.
	<ol>
		<li>Combine in a 1 L flask 556.25 ml of deionized H2O, 90 ml of molasses, and 22 g of agar. Autoclave with a stir bar for 30 min, cool and add 9.25 ml of 10% Tegosept in 95% EtOH.</li>
		<li>Pour the mixture into a meat tray of 21 x 14.5 cm. Ensure that the volume is enough to cover 15 - 17 trays. Wait 10 - 20 min to solidify and cover each tray with a plastic film to avoid drying. Several flasks of 1 L can be prepared at the same time. Store the molasses trays at 4 &#186;C wrapped in plastic until required.</li>
	</ol>
	</li>
	<li>Add a molasses tray covered with wet yeast inside the cage with the adult flies.
	<ol>
		<li>Before adding the plate, place the cage at 4 &#186;C in the cold room for 30 min.</li>
		<li>Add 200 ml of deionized H2O into a beaker and slowly pour dry yeast while stirring with a spoon. Stop adding dry yeast when the mixture becomes approximately the consistency of peanut butter to prevent flies getting stuck into the food. 
		NOTE: It is important not to allow the food solution to solidify otherwise it will be very difficult to cover the molasses tray and extract the eggs during the harvesting process (see section 4.3).</li>
		<li>Remove the plastic film from one molasses tray and cover it with a layer of the fresh prepared wet yeast using a spoon or spatula. Be sure that the entire surface is covered.</li>
		<li>While the cage is still in the cold room, open the net and carefully add the molasses tray with wet yeast into the cage. Close the net and place the cage at RT. 
		NOTE: It may be useful to cover the plate with an empty meat tray during insertion, but be sure not to block access of the flies to the molasses tray.</li>
	</ol>
	</li>
	<li>Change the food plate every 24 - 48 hr to avoid the yeast to get too dry. Adult flies are very sensitive to desiccation.</li>
</ol>
4. End of the Cycle: Harvesting the Embryos
NOTE: The best fly fertility is 3 - 5 days after eclosion. Therefore, harvesting during this time will get the best embryo yield. After desired collections are completed, the cycle is over. The flies should be sacrificed and the cage cleaned as specified in step 2.2.
<ol>
	<li>Typically, 2 days after fly eclosion, introduce a new molasses tray with fresh wet food into the cage, as explained in section 3.2.5. To increase the yield of eggs deposition, with the help of the fingers, make the surface of the wet yeast as&#160;irregular as possible. If an old food tray is present in the cage, before adding the new plate, carefully remove it and discard it into a biohazard bag. Another tip for increasing the yield is to remove the cotton layer so all the eggs will be laid in the food. Maintain the cage at RT.</li>
	<li>Observe the embryos as small white dots. 
	NOTE: The eggs are ready to be collected. For routine maintenance of the fly cage, 2 day collection is the recommended time. This collection will contain mostly embryos and few 1st instar larvae. See representative results for typical yields of harvested embryos for various collection time points.</li>
	<li>Harvest the embryos
	<ol>
		<li>Place the cage in the cold room for about 30 min. In the mean time prepare the plastic container and the fly food for the next cycle as explained in section 1.</li>
		<li>In an 8 L autoclave tray add water to &#188; of the total capacity and then add 1 ml of 10% Triton X-100. While the cage is in the cold room remove carefully the molasses plate and place it inside a biohazard bag.</li>
		<li>Remove the tray from the bag and submerge it quickly (to prevent flies from escaping) in the detergent solution in the autoclave tray. Close the biohazard bag and put in a biohazard container. Flies will not drown in water that does not contain detergent.</li>
		<li>Use a large paintbrush and distilled water gun to wash embryos and yeast off of molasses. Discard plates and molasses into biohazard box.</li>
		<li>Pour solution carefully through a three sieve set (coarsest sieve 30 on top, 40 in the middle and 100 on bottom). Wash clumps on top level with distilled water gun until no clumps remain. 
		NOTE: The adult flies and 3rd instar larvae will be retained in the sieve 30.
		<ol>
			<li>Rinse the autoclave tray with distilled water and pour the rinse water through the sieves. Remove top sieve and repeat process if yeast clumps remain. Most of the 1st and all of the 2nd instar larvae will remain in the sieve 40.</li>
		</ol>
		</li>
		<li>Remove the second sieve. The yellowish material in the third (bottom) sieve is the mixture of embryos and small 1st instar larvae. Use distilled water gun to move all the eggs to one side of the sieve. Collect them with a spatula and weigh them.</li>
		<li>During these cycles the collection yield of embryos/larvae ranges between 7 and 13 g. Use only 1.5 g of this material to start a new cycle (section 1). Use the rest of the embryos for further processing or can be discarded.</li>
	</ol>
	</li>
</ol>
5. Further Processing
NOTE: The embryos not utilized for continuation of the population can be used immediately for experiments or alternatively can be frozen at -80 &#186;C. For both options, the embryos may need to&#160;first be dechorionated.
<ol>
	<li>For dechorionation, wash the embryos for 2 min in 50% bleach solution and rinse thoroughly with distilled water.</li>
	<li>To freeze the embryos, first dry by pressing firmly with a paper towel, and then weigh and place them, with the help of a spatula, into a 15 or 50 ml conical tube. Finally submerge the conical tube in liquid N2 for a few seconds.</li>
</ol>

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G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+interference+machinery+influences+the+nuclear+organization+of+a+chromatin+insulator.">RNA interference machinery influences the nuclear organization of a chromatin insulator.</a> Nat Genet. 38, 936-941 (2006).</li><li>Matzat, L. H., Dale, R. K., Moshkovich, N., 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=Tissue-specific+regulation+of+chromatin+insulator+function.">Tissue-specific regulation of chromatin insulator function.</a> PLoS Genet. 8, e1003069 (2012).</li><li>King, M. R., Matzat, L. H., Dale, R. K., Lim, S. J., Lei, E. 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The authors have nothing to disclose.

Starting with 1.5 g of material one can obtain a yield of collected embryos between 7 and 13 g per cycle. To get such an amount of material it is crucial to maintain the right culturing conditions for all the stages of the fly cycle.
The most important parameters are temperature and humidity, which should be 24 &#186;C and 35% respectively. If these two parameters cannot be held constant in the normal environment of the lab, one possibility would be to place the fly cage into a incubator or an environmental chamber. Other protocols recommend 70% humidity and also a constant 24 hr light-dark cycle to increase the yield of the produced eggs9,10. However keeping the humidity around 35% avoids bacterial contamination, and since the purpose of this protocol is only the maintenance of a population cage, the flies are kept in the normal light environment of the lab.
Another important point is to keep disturbances to the adult flies as low as possible. It may be advisable to keep the cage in a location separate from the fly room to avoid cross contamination from other flies.
The culturing of large populations of transgenic and mutant flies is not recommended, since it is very difficult to maintain their purity and they can exhibit pronounced abnormal mating behavior in large population cages14.
One possible problem while culturing Drosophila in large quantities is the presence of other organisms like mites and/or mold, which will compete for the food and therefore reduce the yield of produced eggs. To avoid this, it is very important to keep all the equipment clean, washing the cage, nets and fly boxes with water and soap, and discarding the disposable material (foam plugs) after every cycle. To reduce mold growing, propionic and phosphoric acid are added into the wet yeast when preparing the fly food in step 1.2 and Tegosept when preparing the molasses tray in step 3.1. It is sometimes helpful to place materials such as the plastic box or sieves in -20 &#730;C when time is short and materials cannot be cleaned right away.
A whole collection cycle occurs in 14 - 15 days, beginning when the embryos are seeded into the fly box and ending with the last collection day. During this period of time, it is recommended to organize a schedule in order to remember all the necessary steps for the maintenance of a population fly cage (Fig. 2), detailed in the protocol section. From the day that the eggs are seeded, until the adult flies emerge, it is necessary only to inspect the plastic box, and when pupation occurs, to place them inside the cage. After that, the flies have to be fed every 2 - 3 days until embryos are seeded for a new cycle. In the whole protocol, the longest day is during the collection and seeding of the embryos for starting a new cycle. As commented in the protocol, the best egg yield is 3 - 5 days after adult eclosion, and finally declines 2 days later. This gives us some flexibility in order to choose the day in which the harvest of the eggs will be performed.
If for any reason the yield of collected embryos is less than the desired starting amount (1.5 g), one always can add a new molasses tray and collect more eggs the next day. For constant collections, it is recommended to maintain 2 cages in parallel, and if higher quantities of embryos are required, it is also is possible to use bigger cages. In case of doing short time collections, one way to increase the yield is to take advantage of the egg-laying burst in the morning.
There are many advantages of collecting large amounts of various developmental stages. For example, collected embryos from population cages have been used very successfully in immunoprecipitation assays&#160;6-8, mass collection of larval tissues from dissociated larvae has demonstrated to be a very good source for 3C experiments4 and RNA preparations15, and heads from adult flies have been utilized for ChIP experiments16. In addition, adults are often needed to make fly extracts for tissue culture17.
One of the most promising applications of the cage is to provide material for high-throughput assays that allow the analysis and screening of genes, transcripts, proteins and metabolites in response to the exposure of pathogens, biological molecules, chemical substances and ionizing radiation. In these large-scale assays large numbers of individuals are required, and the fly population cage described here can be very helpful in order to obtain great quantities of material during the different phases of the Drosophila life cycle for their analysis and screening18.

The ability to combine genetic and biochemical approaches has made Drosophila a particularly suitable organism for biochemistry and molecular biology studies1-3. These studies often require large amounts of biological material, not only from adult flies, but also from larvae4, pupae5 and embryos6-8. To obtain large quantities of material, researchers have cultured flies using large containers known as fly &#34;population cages&#34;. These cages consist of a cylinder made of plastic covered by a net on both sides to allow the introduction of the food inside the cage without the flies escaping. These cages can be homemade9-11 or purchased from a company (see table of specific materials/equipment).
A major advantage of using this system to grow large numbers of flies is that the cycle of the fruit fly12 can be controlled in a way that all the flies develop in a relatively synchronized manner. This synchronization is achieved by seeding new embryos, feeding larvae/flies and sacrificing the adult flies at precise times. Using a synchronized fly population is particularly useful for developmental studies13.
The start of a new population cage from a few flies is a time-consuming process requiring many cycles of amplification 9-11. Even using bigger containers like fly culture bottles or minicages, the whole process can last for months. To avoid this time consuming step, many Drosophila laboratories regularly maintain such cages. It is most convenient to start a new cage starting from an embryo collection from an already established population cage. In general, most labs maintain wildtype population cages, such as Oregon R or Canton S. This manuscript presents a detailed protocol to maintain fly population cages.

<table>
	<tbody>
		<tr>
			<td>Bacto-Agar</td>
			<td>Beckton Dickinson</td>
			<td>214010</td>
			<td></td>
		</tr>
		<tr>
			<td>Curity practical cotton roll</td>
			<td>Kendall</td>
			<td>2287</td>
			<td></td>
		</tr>
		<tr>
			<td>Dry yeast</td>
			<td>Affymetrix</td>
			<td>23540</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>GE Healthcare Life Science</td>
			<td>1001-085</td>
			<td></td>
		</tr>
		<tr>
			<td>Foam tube plugs</td>
			<td>Jaece</td>
			<td>L800-D2</td>
			<td>50 mm Diameter x 55 mm Length</td>
		</tr>
		<tr>
			<td>Fly population cage</td>
			<td>Flystuff</td>
			<td>59-116</td>
			<td>9&#8243; Diameter x 14.4&#8243; Length. Includes the nets for the cage.</td>
		</tr>
		<tr>
			<td>Meat tray</td>
			<td>Genpak</td>
			<td>1002S (#2S)</td>
			<td>8.25 x 5.75 x 0.5 inches</td>
		</tr>
		<tr>
			<td>Molasses</td>
			<td>Grandma&#180;s</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic container</td>
			<td>Rubbermaid</td>
			<td>4022-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic film</td>
			<td>Glad</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphoric acid</td>
			<td>Fisher Scientific</td>
			<td>S 93326</td>
			<td>Toxic. Handle in Chemistry Hood</td>
		</tr>
		<tr>
			<td>Propionic acid</td>
			<td>Fisher Scientific</td>
			<td>A258-500</td>
			<td>Toxic. Handle in Chemistry Hood</td>
		</tr>
		<tr>
			<td>Stainless steel&#160; sieve #100</td>
			<td>VWR</td>
			<td>57324-400</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless steel&#160; sieve #40</td>
			<td>VWR</td>
			<td>57324-272</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless steel&#160; sieve #30</td>
			<td>VWR</td>
			<td>57324-240</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>MP</td>
			<td>152584</td>
			<td></td>
		</tr>
		<tr>
			<td>Tegasept</td>
			<td>LabScientific</td>
			<td>FLY5501</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton-X100</td>
			<td>Fisher Scientific</td>
			<td>BP151-500</td>
			<td></td>
		</tr>
	</tbody>
</table>

maintenance of a drosophila melanogaster population cage

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life cycle of Drosophila enables collection of large quantities of material from embryos, larvae, pupae and adult flies, in a synchronized way, at a low economic cost. A major strategy for propagating large numbers of flies is the use of a fly population cage. This useful and common tool in the Drososphila community is an efficient way to regularly produce milligrams to tens of grams of embryos, depending on uniformity of developmental stage desired. While a population cage can be time consuming to set up, maintaining a cage over months takes much less time and enables rapid collection of biological material in a short period. This paper describes a detailed and flexible protocol for the maintenance of a Drosophila melanogaster population cage, starting with 1.5 g of harvested material from the previous cycle.

The maintenance of a fly cage population is based on the fly life cycle. Therefore, after placing the initial biological mixture of embryos and larvae in the plastic container (Figure 1A) the fertilized eggs will become larvae in no more than 2 days and the larvae will grow for 4 days, relatively synchronized through the different instar larvae stages (see Figure 1B).
After the larvae have completed the 3rd instar larval stage they will pupate and cover the surface of the cotton inside the plastic container with some on the internal surface of the lid (Figure 1C). This pupation period will go for the next 4 days until the flies finish the metamorphosis process. During this period it is strongly recommended to transfer the flies into the cage. The first adult flies will start to eclose at day 9 - 10 after the initiation of the cycle (Figure 1D) and by day 11 all of them will have emerged from the pupae.
The best yield of the embryos collection is obtained 3 - 5 days after eclosion (day 13&#160;to 15) and finally declines in day 17 (7 days after eclosion). Therefore, it is recommended to add the last tray of fresh food at day 13 and collect the eggs at day 15 (Figure 1E and F). This will allow collection of the maximum number of embryos. Adding a third extra day before collecting may result in the food being too dry and consequently will diminish the yield of collected embryos. Table 1 shows the yield of 6 consecutive cycle collections, with a starting material of 1.5 g in each cycle, and Figure 2 is the recommended schedule for a whole cycle collection.
The material harvested in 2 day&#160;periods is mostly composed of embryos. However a small number of larvae are also present. For cage maintenance purposes these larvae are not a problem, since some degree of asynchronicity is expected. Nevertheless for some biochemistry experiments, the purity of these embryos is very important, but at the same time it is also necessary to collect large quantities of eggs to perform these experiments. For this reason several short consecutive collections were performed at 1 hr and 3 hr, after quickly adding at RT a new plate with food, to provide an estimation of the yield and purity that can be obtained with this method (Table 2 and Figure 3). The 1 hr consecutive collections started with a very low yield (14.5 mg) but then the quantities of embryos increased for each time collected but the last time. The low yield at first may be due to the flies being stressed during the process of changing the food but later acclimating. On the other hand, no larvae were observed in the five sequential collections (Figure 3A), showing that shorter collections yield higher purity. If cotton is not removed from the cage, the likelihood that larvae from earlier time points will remain and enter the fresh plate is increased. Even when cotton is removed, at times larvae will wander on the side of the cage and enter the food during a collection.
For the 3 hr consecutive collections, the yield ranged from 840 to 1250 mg, which is around 10 times more than the yields obtained in the 1 hr collections. The purity of these embryos was near 100% (Figure 3B). Occasional larvae were observed. Some developmental studies require a very strict synchronization for the embryos collected. To increase the synchronization purity, it is recommended to discard the first collection plate because if conditions are not ideal, females can retain more mature eggs and deposit them when conditions improve, such as with introduction of a fresh plate. Also it is important to know that older adult flies (&#62;6 days) produce less synchronized embryos. To verify the degree of synchronization with high accuracy, a DAPI staining of the collected embryos is recommended.
<img alt="Figure 1" src="/files/ftp_upload/53756/53756fig1.jpg" /> 
Figure 1. The Drosophila melanogaster&#160;Life Cycle in a Fly Population Cage.&#160;(A) Initiation of the cycle seeding 1.5 g of embryos into the plastic box container. As a reference, in the lower panel, the length of the filter paper is 8.5 cm. (B) Larvae growing relatively synchronized 5 days after the initiation of the cycle. (C) At day 8, most of the flies are in the pupa stage. The lower panels in B (B&#39;) and C (C&#39;) are magnifications of the indicated areas in the upper panels. (D) Adult flies emerged from the pupae 10 days after cycle initiation. (E) Adult flies inside the population cage before starting the process of collecting the embryos, at day 15. The yellowish material on the fly food are fertilized eggs. (F) Embryos (and a few larvae) collected at the bottom of the 100 coarsest sieve, after a whole cage cycle. <a href="https://www.jove.com/files/ftp_upload/53756/53756fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53756/53756fig2.jpg" /> 
Figure 2.&#160;Whole Cycle Collection Schedule. Figure 2 shows the recommended schedule for a complete cycle collection. <a href="https://www.jove.com/files/ftp_upload/53756/53756fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53756/53756fig3.jpg" /> 
Figure 3.&#160;Purity of 1 hr and 3 hr consecutive collections. Representative image of embryos collected at 1 hr (A) and 3 hr (B) after adding a new tray of food. <a href="https://www.jove.com/files/ftp_upload/53756/53756fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table>
	<tbody>
		<tr>
			<td>Cycle number</td>
			<td>Yield (g)</td>
		</tr>
		<tr>
			<td>1</td>
			<td>10.3</td>
		</tr>
		<tr>
			<td>2</td>
			<td>9.5</td>
		</tr>
		<tr>
			<td>3</td>
			<td>10.3</td>
		</tr>
		<tr>
			<td>4</td>
			<td>12.7</td>
		</tr>
		<tr>
			<td>5</td>
			<td>10</td>
		</tr>
		<tr>
			<td>6</td>
			<td>7.1</td>
		</tr>
	</tbody>
</table>
Table 1.&#160;Yield of Several Cycle Collections. Table 1 shows the yield of 6 consecutive cycle collections, with a starting material of 1.5 g. In each cycle, the last tray of food was added at day 13 and the harvest was done at day 15.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Collection #</td>
			<td>Length of collection (hr)&#160;</td>
			<td>Yield (mg)</td>
		</tr>
		<tr>
			<td>1</td>
			<td>1</td>
			<td>14.5</td>
		</tr>
		<tr>
			<td>2</td>
			<td>1</td>
			<td>21.6</td>
		</tr>
		<tr>
			<td>3</td>
			<td>1</td>
			<td>56.1</td>
		</tr>
		<tr>
			<td>4</td>
			<td>1</td>
			<td>160.1</td>
		</tr>
		<tr>
			<td>5</td>
			<td>1</td>
			<td>106.1</td>
		</tr>
		<tr>
			<td>1</td>
			<td>3</td>
			<td>960</td>
		</tr>
		<tr>
			<td>2</td>
			<td>3</td>
			<td>840</td>
		</tr>
		<tr>
			<td>3</td>
			<td>3</td>
			<td>1250</td>
		</tr>
	</tbody>
</table>
Table 2. Yield of 1 hr and 3 hr Consecutive Collections. Table 2 shows the yield of 5 consecutive collections 1 hr after adding new food and 3 consecutive collections 3 hr after placing a new molasses plate.

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Maintenance of a Drosophila melanogaster Population Cage

This manuscript reports a detailed protocol for culturing, on a regular basis, a population of Drosophila melanogaster using a fly population cage.

Maintenance of a Drosophila melanogaster Population Cage

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life ...

maintenance-of-a-drosophila-melanogaster-population-cage

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13.1K Views.  National Institutes of Health. This manuscript reports a detailed protocol for culturing, on a regular basis, a population of Drosophila melanogaster using a fly population cage.

Video: Maintenance of a Drosophila melanogaster Population Cage

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

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

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

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

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

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

Quantitative Comparison of cis-Regulatory Element (CRE) Activities in Transgenic Drosophila melanogaster

Gene expression patterns are specified by cis-regulatory element (CRE) sequences, which are also called enhancers or cis-regulatory modules. A typical CRE possesses an arrangement of binding sites for several transcription factor proteins that confer a regulatory logic specifying when, where, and at what level the regulated gene(s) is expressed. The full set of CREs within an animal genome encodes the organism&prime;s program for development1, and empirical as well as theoretical studies indicate that mutations in CREs played a prominent role in morphological evolution2-4. Moreover, human genome wide association studies indicate that genetic variation in CREs contribute substantially to phenotypic variation5,6. Thus, understanding regulatory logic and how mutations affect such logic is a central goal of genetics. 
Reporter transgenes provide a powerful method to study the in vivo function of CREs. Here a known or suspected CRE sequence is coupled to heterologous promoter and coding sequences for a reporter gene encoding an easily observable protein product. When a reporter transgene is inserted into a host organism, the CRE&prime;s activity becomes visible in the form of the encoded reporter protein. P-element mediated transgenesis in the fruit fly species Drosophila (D.) melanogaster7 has been used for decades to introduce reporter transgenes into this model organism, though the genomic placement of transgenes is random. Hence, reporter gene activity is strongly influenced by the local chromatin and gene environment, limiting CRE comparisons to being qualitative. In recent years, the phiC31 based integration system was adapted for use in D. melanogaster to insert transgenes into specific genome landing sites8-10. This capability has made the quantitative measurement of gene and, relevant here, CRE activity11-13 feasible. The production of transgenic fruit flies can be outsourced, including phiC31-based integration, eliminating the need to purchase expensive equipment and/or have proficiency at specialized transgene injection protocols. 
Here, we present a general protocol to quantitatively evaluate a CRE&prime;s activity, and show how this approach can be used to measure the effects of an introduced mutation on a CRE&prime;s activity and to compare the activities of orthologous CREs. Although the examples given are for a CRE active during fruit fly metamorphosis, the approach can be applied to other developmental stages, fruit fly species, or model organisms. Ultimately, a more widespread use of this approach to study CREs should advance an understanding of regulatory logic and how logic can vary and evolve.

Quantitative Comparison of cis-Regulatory Element CRE Activities in Transgenic Drosophila melanogaster

Phenotypic variation for traits can result from mutations in cis-regulatory element (CRE) sequences that control gene expression patterns. Methods derived for use in Drosophila melanogaster can quantitatively compare the levels of spatial and temporal patterns of gene expression mediated by modified or naturally occurring CRE variants.

Gene expression patterns are specified by cis-regulatory element (CRE) sequences, which are also called enhancers or cis-regulatory modules. A typical CRE ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called imaginal discs. These discs give rise to a high percentage of adult structures that are found within the adult fly. Here we describe a protocol that has been optimized to recover these discs and prepare them for analysis with antibodies, transcriptional reporters and protein traps. This procedure is best suited for thin tissues like imaginal discs, but can be easily modified for use with thicker tissues such as the larval brain and adult ovary. The written protocol and accompanying video will guide the reader/viewer through the dissection of third instar larvae, fixation of tissue, and treatment of imaginal discs with antibodies. The protocol can be used to dissect imaginal discs from younger first and second instar larvae as well. The advantage of this protocol is that it is relatively short and it has been optimized for the high quality preservation of the dissected tissue. Another advantage is that the fixation procedure that is employed works well with the overwhelming number of antibodies that recognize Drosophila proteins. In our experience, there is a very small number of sensitive antibodies that do not work well with this procedure. In these situations, the remedy appears to be to use an alternate fixation cocktail while continuing to follow the guidelines that we have set forth for the dissection steps and antibody incubations.

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

The adult structures of Drosophila are derived from sac-like structures called imaginal discs. Analysis of these discs provides insight into many developmental processes including tissue determination, compartment boundary establishment, cell proliferation, cell fate specification, and planar cell polarity. This protocol is used to prepare imaginal discs for light/fluorescent microscopy.

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called ...