1. Preparing Tools and Accessories
<ol>
	<li>Tip/funnel stoppers
	<ol>
		<li>Obtain two sponge plugs (the diameter of the plugs must be slightly larger than the internal diameter of the vials used to transfer flies). Make a hole in the centers of the sponge plugs with a heated electric soldering iron.</li>
		<li>Obtain two 1 mL pipette tips, cut one in half transversely with a sharp knife, and discard the pointed end. Then, cut 1.5 cm of the pointed end off the second pipette tip. Glue the remains of the two pipette tips together with an all-purpose adhesive to make an elongated pipette tip (Figure 1A).</li>
		<li>Insert a funnel and the elongated pipette tip into the sponge plugs to make a tip and funnel stopper (hereafter referred to as T- and F-stoppers) and cap the pipette tip with a 100 &#956;L microcentrifuge tube (Figure 1A). 
		NOTE: The length of the funnel stem must be greater than the height of the plug. If it is shorter than or equal to the height of the plug, then flies will escape from the stem opening. The end of the funnel stem should be situated at least 2 cm above the surface of the culture medium or the bottom of an empty vial. Small funnels (e.g., disk diameter &#60;60 mm) with small internal stem opening diameters (&#60;5 mm) are preferable. Either a glass or a plastic funnel can be used to make an F-stopper. However, plastic funnels are preferable for biology classes, as they break less readily than glass funnels.</li>
	</ol>
	</li>
	<li>Microdissecting needles 
	<ol>
		<li>Obtain mechanical pencils that feel comfortable in the hand and insect pins that match the diameters (e.g., 0.5 mm, 0.7 mm) of their lead refills.</li>
		<li>Cut the wide ends of the insect pins with a pair of pliers and file the cut flat. Replace the lead with the pins (Figure 1B). Press the click button and feed out 0.5&#8211;1 cm of a pin to conduct a dissection. Clean the pin and push it completely back into the pencil shaft after a dissection activity to make it safe for any person to handle. 
		NOTE: Microdissecting needles are useful not only in dissections of organs such as larval salivary glands but also in counting and sorting dead adult flies.</li>
	</ol>
	</li>
	<li>Hard icepacks 
	<ol>
		<li>Obtain several refreezable hard icepacks (large-sized icepacks are preferable). Figure 1C shows an icepacks that worked well, which measures 26.5 cm x 14.5 cm x 2.5 cm and has top and bottom sides that are completely flat.</li>
		<li>Cut medical gauze (nonsterile) into pieces that are slightly smaller than the cold surfaces of the icepacks they cover. For example, a piece of medical gauze slightly smaller than 26.5 cm x 14.5 cm is preferable to cover an icepack shown in Figure 1C. 
		NOTE: The necessary accessories for these chilling tools include: an ice box (we used a 25 cm x 15 cm x 15 cm foam box for one person and 37 cm x 28 cm x 20 cm box for more than one person), which is used to store crushed ice; a pair of fine-point tweezers, which are used to grab chilled flies by their wings and transfer them to a vial; a pair of protective work gloves, which are used to take chilled icepacks out of a -20 &#176;C freezer; and plastic film, which is used to cover the stage of a stereomicroscope.</li>
	</ol>
	</li>
	<li>Drosophila egg collection cage 
	NOTE: Ready-made Drosophila egg collection cages are available from many biotechnology companies11. Described here is a small acrylic bottle-shaped egg collection cage for 60 mm Petri dishes (Figure 1D left; the cage design is shown in the middle). It can be adapted for other Petri dish sizes (e.g., 100 mm, 35 mm). This allows the transfer of flies into or out of the cage with ease. A simple cage can be prepared as follows.
	<ol>
		<li>Use a snap cutter to cut a soft plastic drink bottle (500 mL, internal diameter ca. 65 mm) into an approximate 2:1 (pointed end:blunt end) ratio and discard the blunt end.</li>
		<li>Wrap a strip of card paper around an apple juice plate (internal diameter 60 mm) with adhesive tape [the apple juice plate is used to collect eggs (Figure 1E, right)].</li>
	</ol>
	</li>
	<li>Cordless tube brush driver
	<ol>
		<li>Obtain a cordless drill driver (max speed = 500 rpm).</li>
		<li>Obtain a tube brush that has bristles along its sides as well as its front. Ideally, the diameter of the brush should be slightly larger than the diameter of the culture vials that need to be cleaned. Cut the end of its handle so it can be inserted into the drill chuck (Figure 1D). 
		NOTE: The necessary accessories for these cleaning tools include stainless steel sponges and long cuff rubber gloves.</li>
	</ol>
	</li>
</ol>
2. Transferring Adult Flies from Vial A to Vial B
NOTE: Transferring adult flies from one vial to another is the most common practice conducted in Drosophila experiments [e.g., transferring flies from old culture (A) to fresh culture (B) or from a cross vial (A) to empty vial (B)] for anesthetizing. The protocol described here can be used for any adult fly transferring activities. Unless otherwise stated, this protocol is used to transfer flies from vial A to vial B throughout this paper.
<ol>
	<li>Check the stem of the funnel of an F-stopper and the pipette tip of a T-stopper carefully, then clear any flies that remain in the stoppers with a rubber air blower. This step is of paramount importance, especially when one set of T- and F-stoppers is used for the continuous transferring of different Drosophila lines.</li>
	<li>Tap down the flies in vial A and replace its plug with a T-stopper, then plug vial B with an F-stopper.</li>
	<li>Invert vial A over vial B, insert the pipette tip end of the T-stopper into the funnel opening of the F-stopper, knock the edge of inverted vial A to allow flies to slip out of the pipette tip and through the stem of the funnel, and drop into vial B. If any old food in vial A becomes less compact, it may drop when vial A is inverted and knocked. In such a situation, invert vial B over vial A and allow the flies to crawl up into vial B.</li>
	<li>Separate the T-stopper from the F-stopper. Cap the pipette tip end of the T-stopper with a 200 &#956;L microcentrifuge tube if the remaining flies in vial A need to be transferred to other vials momentarily; otherwise, remove the T-stopper and replug vial A. Remove the F-stopper and replug vial B.</li>
</ol>
3. Immobilizing Flies by Chilling
<ol>
	<li>Keep hard refreezable icepacks in a -20 &#176;C freezer for at least 24 h before use.</li>
	<li>Place a chilled, hard icepack at room temperature (RT) for 20 min. Slightly moisten a piece of non-aseptic medical gauze with some running water and allow it to closely cling to the surface of the icepack. The medical gauze can be reused in the next fly chilling. At the same time, chill an empty vial in crushed ice.</li>
	<li>Transfer adult flies that need to be immobilized into the chilled empty vial (CEV). When the two transfer vials are separated, cover the CEV with a Petri dish or a plug and knock the CEV against the crushed ice to tap all the flies in the CEV down to the bottom. Repeat this process several times until all flies are immobilized. The flies will be immobilized within 30 s. Next, place the CEV in the ice for 1 min. It is not advisable to transfer too many flies at one time for anesthetizing.</li>
	<li>Pour the chilled flies out onto the medical gauze that covers the ice pack. Spread out the overlapping flies with a paintbrush and make sure that each fly can be chilled by the cold surface of the icepack. If a chilled hard icepack swells slightly, place it on a towel and work on its flat side.</li>
	<li>Remove the stage clips from the stereomicroscope, cover the stage with a piece of plastic film, and put the icepack onto the stage. Turn on the top light (a cold light source is desirable), focus the stereomicroscope and move the icepack until the chilled flies can be seen clearly.</li>
</ol>
4. Killing Adult Flies for Counting, Sorting, or Discarding
<ol>
	<li>Transfer adult flies into an empty vial and cover it with a Petri dish.</li>
	<li>Invert the vial, heat it for 1 min + 20 s in a microwave oven, and allow the dead flies drop into the Petri dish.</li>
	<li>Put on protective work gloves and take the vial out of the microwave. Pour the dead flies onto a white paper card, count or examine the flies with a microdissecting needle under a stereomicroscope, and dispose of the fly bodies in a garbage can after observation.</li>
	<li>To kill unwanted flies, heat the flies for 2&#8211;3 min in a microwave oven, then tap the carcasses into a garbage can. 
	NOTE: It is not advisable to kill some wing mutant strains (e.g., wing length mutants) for examination, as it is difficult to judge from the carcasses if the wings extend beyond the tip of the abdomen, which is seen in wild-type flies.</li>
</ol>
5. Transferring Flies In/Out of Bottle-shaped Egg Collection Cage
NOTE: As mentioned above, T- and F-stoppers are used to transfer flies into and out of the egg collection cage. Flies do not need to be anesthetized throughout this process. Other details, such as preparing the apple juice medium, egg collection, and dechorionization, can be found in the literature12.
<ol>
	<li>Insert the egg collection cage into the apple juice plate or mount the apple juice plate to the cage made of a soft drink bottle. Seal the joint around the two components with a strip of paraffin film.</li>
	<li>Place as many flies as possible into the cage and replug the cage with a foam stopper after transferring the flies.</li>
	<li>To change the food for flies in the cage, transfer the flies in the cage to an empty vial.</li>
	<li>Replace the apple juice plate and reseal it, then transfer the flies from the vial back to the cage.</li>
	<li>When egg collection ends, transfer the flies into an empty vial and transfer them into culture vials.</li>
</ol>
6. Cleaning Glass Culture Vials
NOTE: Generally, an old culture vial contains live flies. In the protocol described here, these flies DO NOT need to be killed before cleaning unless they are transgenic flies.
<ol>
	<li>Remove any permanent marker ink from the glass culture vials with wet, stainless steel sponges.</li>
	<li>Soak the culture vials in running water.
	<ol>
		<li>Fill a laboratory sink with water, add liquid dishwashing soap into the water, and mix.</li>
		<li>Immerse the culture vials into the water, then remove the plug, allowing the water to run into the vial. The dish detergent in the water will make any remaining adult flies sink to the bottom and drown in the water.</li>
		<li>Soak the old culture vials in water for at least 30 min.</li>
	</ol>
	</li>
	<li>Loosen the chuck of the drill, insert the test tube brush and retighten the chuck. Check the direction of the rotation selector and ensure that the drill rotates clockwise. Adjust the speed trigger and ensure that the maximum speed is less than 500 rpm.</li>
	<li>Clean the culture vials.
	<ol>
		<li>Clean the culture vials roughly.
		<ol>
			<li>Put a long cuff rubber glove on the non-dominant hand and hold the vial in the water.</li>
			<li>Hold the cordless tube brush driver with the bare dominant hand, squeeze the brush into the culture vial, and squeeze the trigger. 
			NOTE: Do not dip the battery into the water. The rotating brush will break up old food, pupa, etc., and remove more than 95% of the waste.</li>
			<li>Dump the waste into a separate garbage can. Repeat this process until most of the waste in each vial has been cleaned.</li>
		</ol>
		</li>
		<li>Clean the culture vials thoroughly.
		<ol>
			<li>Clean the tube brush, drain and clean the sink, and refill it with clean water.</li>
			<li>Remove the remaining waste from each culture vial as described in section 6.4.1.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Jennings, B. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+&#8211;+a+versatile+model+in+biology+&#38;+medicine.">Drosophila &#8211; a versatile model in biology &#38; medicine.</a> Materials Today. 14 (5), 190-195 (2011).</li><li>JoVE Science Education Database. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Biology I: yeast, Drosophila and C. elegans. An Introduction to Drosophila melanogaster. , (2018).</li><li>Ashburner, M., Roote, J., Sullivan, W., Ashburner, M., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laboratory+Culture+of+Drosophila.">Laboratory Culture of Drosophila.</a> Drosophila Protocols. , (2000).</li><li>Greenspan, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fly+pushing:+The+theory+and+practice+of+Drosophila+genetics.">Fly pushing: The theory and practice of Drosophila genetics.</a> Cold Spring Harbor Laboratory Press. , (2004).</li><li>Ashburner, M., Thompson, J., Ashburner, M., Wright, T. R. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+laboratory+culture+of+Drosophila.">The laboratory culture of Drosophila.</a> The genetics and biology of Drosophila. 2a, 1-109 (1978).</li><li>Ratterman, D. M., O&#39;Donnell, 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=Eliminating+ether+by+using+ice+for+Drosophila+labs.">Eliminating ether by using ice for Drosophila labs.</a> Tested Studies For Laboratory Teaching. , 259-265 (2003).</li><li>. Culturing techniques for Drosophila Available from: <a target="_blank" href="https://www.ptbeach.com/cms/lib/NJ01000839/Centricity/Domain/113/ap%20biology%20Labs/Culturing%20techniques%20for%20Drosophila.pdf">https://www.ptbeach.com/cms/lib/NJ01000839/Centricity/Domain/113/ap%20biology%20Labs/Culturing%20techniques%20for%20Drosophila.pdf</a> (2019)</li><li>Markow, T. A., O&#39;Grady, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Drosophila: A Guide to Species Identification and Use. , (2006).</li><li>Artiss, T., Hughes, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Taking+the+Headaches+Out+of+Anesthetizing+Drosophila:+A+Cheap+&#38;+Easy+Method+of+Constructing+Carbon+Dioxide+Staging.">Taking the Headaches Out of Anesthetizing Drosophila: A Cheap &#38; Easy Method of Constructing Carbon Dioxide Staging.</a> The American Biology Teacher. 69 (8), e77-e80 (2007).</li><li>Qu, W. -. H., Zhu, T. -. B., Yang, D. -. X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Modified+Cooling+Method+and+its+Application+in+Drosophila+Experiments.">A Modified Cooling Method and its Application in Drosophila Experiments.</a> Journal Of Biological Education. 49 (3), 302-308 (2015).</li><li>. Egg-laying cages for drosophila Available from: <a target="_blank" href="https://www.kisker-biotech.com/frontoffice/product?produitId=0H-19-17">https://www.kisker-biotech.com/frontoffice/product?produitId=0H-19-17</a> (2018)</li><li>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=.">.</a> Drosophila: a practical approach. , (1998).</li><li>Tang, M., Peng, Q. -. F., Yang, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+devices+for+Drosophila+experiments+(in+Chinese).">Two devices for Drosophila experiments (in Chinese).</a> Bulletin of Biology. 45 (11), 49-50 (2010).</li><li>Zhou, T. -. y., Gan, J., Yang, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+sponge+plug+and+sponge+plug+based+fly+transferring+device+for+Drosophila+experiments+(in+Chinese).">Preparation of sponge plug and sponge plug based fly transferring device for Drosophila experiments (in Chinese).</a> Bulletin of Biology. 46 (6), 49-50 (2011).</li><li>Yang, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Genetics laboratory investigation. , (2016).</li><li>Yang, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Carnivory+in+the+larvae+of+Drosophila+melanogaster+and+other+Drosophila+species.">Carnivory in the larvae of Drosophila melanogaster and other Drosophila species.</a> Scientific Reports. 8, (2018).</li><li>Stocker, H., Gallant, P., Dahmann, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Getting+Started:+An+Overview+on+Raising+and+Handling+Drosophila.">Getting Started: An Overview on Raising and Handling Drosophila.</a> Drosophila: Methods and Protocols. , (2008).</li></ol>

The author have nothing to disclose.

Some homemade tools for handling basic activities involved in Drosophila rearing and experimentation are described in this paper. These tools are simple but rather effective. Virtually, any lab can make these tools with ease, and a research or a teaching laboratory does not need to find a ready-made alternative that is perhaps not available locally.
Fly transferring is the most common practice and a difficult task in Drosophila experiments. Unfortunately, until now, there hav...

The fruit fly, Drosophila melanogaster, is a model organism widely used in biological research and biology education to study a wide range of topics1,2. The basic problems of handling Drosophila are the transfer of adults from vial to vial and immobilization of the flies so they are easier to handle, as all adults (except for some mutants3,4) can fly.
Conventionally, a researcher transfers flies from one vial to another by holding two vials mouth-to-mouth, tapping the flies down or allowing flies to fly up into another vial, then separating and replugging both vials4. Obviously, this requires that the opening of two vials with the same diameter, and it is hard to control the quantity of flies transferred. Meanwhile, this requires quick hands to get the job done, and escaping stray flies can result in problems for the laboratory or classroom. Adding extra virgin flies or male flies to an already prepared cross is another routine task in Drosophila experiments. Conventionally, flies must be immobilized in the cross vial before the addition of extra flies.
Adult Drosophila are routinely anaesthetized by ether, CO2, or chilling5. Compared to ether and CO2 exposure, chilling is the most cost-efficient agent for immobilizing adult Drosophila and the least harmful to both the flies and researchers (especially young students)6,7. However, water that condenses continuously on the cold surface or chamber wets the flies. It is difficult to determine the phenotypes of wet flies, and they can easily become damaged during manipulation8,9. This has kept the chilling method from becoming more widely accepted.
Tools for fly transferal and a method for fly cooling have been previously described10. Herein, a modified chilling anesthesia technique is reported that is safe, reliable, and feasible for Drosophila experiments. Also described in this paper are 1) methods for killing adults for counting, sorting, or discarding, 2) labor-saving devices and protocols for cleaning glass culture vials, and 3) a simple cage for collecting eggs. The easily designed and cost-effective tools described here can be used to address the difficult issues of fly handling, and these methods have been tested and are proven to be robust, reliable, and easy-to-handle for experienced and novice researchers.

<table>
	<tbody>
		<tr>
			<td>a pair of pliers</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>cordless drill driver</td>
			<td></td>
			<td></td>
			<td>max speed: 500 rpm</td>
		</tr>
		<tr>
			<td>electric soldering iron</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>file</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>funnel</td>
			<td></td>
			<td></td>
			<td>diameter of disk&#60;60mm</td>
		</tr>
		<tr>
			<td>ice box</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>insect pins</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>infrared thermometer</td>
			<td></td>
			<td></td>
			<td>HCIYET HT-830</td>
		</tr>
		<tr>
			<td>long cuff rubber gloves</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>mechanical pencils</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>medical gauze</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>microcentrifuge tube</td>
			<td></td>
			<td></td>
			<td>100 ul</td>
		</tr>
		<tr>
			<td>microwave oven</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>peri dish</td>
			<td></td>
			<td></td>
			<td>internal diameter 60 mm</td>
		</tr>
		<tr>
			<td>pipette tips</td>
			<td></td>
			<td></td>
			<td>1 ml</td>
		</tr>
		<tr>
			<td>plastic film</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>plastic peri dish</td>
			<td></td>
			<td></td>
			<td>&#934;36 mm used to cover the empty vial</td>
		</tr>
		<tr>
			<td>point tweezers</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>protective work gloves</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>re-freezable hard icepacks</td>
			<td></td>
			<td></td>
			<td>26.5&#215;14.5&#215;2.5 cm or larger</td>
		</tr>
		<tr>
			<td>rubber air blower</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>snap cutter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>soft drink bottle</td>
			<td></td>
			<td></td>
			<td>500 ml, internal diameter c.a. 65 mm</td>
		</tr>
		<tr>
			<td>sponge stopper</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>stainless steel sponges</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>tube brush</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>vial</td>
			<td></td>
			<td></td>
			<td>&#934;34 mm &#215; 90 mm</td>
		</tr>
	</tbody>
</table>

simple homemade tools to handle fruit flies&#8212;drosophila melanogaster

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in practice, as adult flies fly. Demonstrated here is how to make some simple and cost-effective tools to address difficult issues in the handling of Drosophila. Holes in foam stoppers are made and pipette tips or funnels are inserted into the holes. Flies then move only in one direction into the pipette tip/funnel assemblage, allowing efficient control of the transfer of adult Drosophila into or out of a vial. Existing protocols have been modified for cool-anesthetizing flies by chilling in crushed ice and transferring them onto a cold, hard icepack surface. The icepack is covered with a piece of medical gauze that keeps immobilized flies from the condensed water when examined under a stereomicroscope. The flies are finally euthanized for counting and sorting or discarded by microwaving. A bottle-shaped cage has also been developed for collecting eggs, as well as a labor-saving device and accompanying protocol for cleaning glass culture vials.

The T- and F-stoppers were developed as a set of simple tools that can be adapted and used in any fly transferring activities. Transferring flies from an old culture into several fresh cultures involves removing the plugs of the fresh vials, replacing them with F-stoppers, then tapping down the flies in the old vial, quickly removing its plug, and replacing it with a T-stopper. If the old food is compact, then it is important to flip the old vial and insert the tip of T-stopper into the o...

Watch this Scientific Journal Video about Simple Homemade Tools to Handle Fruit FliesÃ¢â‚¬â€Drosophila melanogaster at JoVE.com

Simple Homemade Tools to Handle Fruit Flies&#8212;Drosophila melanogaster

Described here is the use of several homemade tools to transfer, chill, and kill adult Drosophila, as well as to clean glass culture vials and collect eggs. These tools are easy to make and are rather efficient in handling Drosophila.

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in ...

simple-homemade-tools-to-handle-fruit-fliesdrosophila-melanogaster

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JoVE Journal

Biology

15.8K Views.  China Agricultural University. Described here is the use of several homemade tools to transfer, chill, and kill adult Drosophila, as well as to clean glass culture vials and collect eggs. These tools are easy to make and are rather efficient in handling Drosophila.

Video: Simple Homemade Tools to Handle Fruit Flies—Drosophila melanogaster

Studying Aggression in Drosophila (fruit flies)

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

Studying Aggression in Drosophila fruit flies

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

Using Micro-Electro-Mechanical Systems (MEMS) to Develop Diagnostic Tools

Using Micro-Electro-Mechanical Systems MEMS to Develop Diagnostic Tools

Homemade Site Directed Mutagenesis of Whole Plasmids

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. With this method the cloned target gene can be altered by substitution, deletion or insertion of a few bases directly into a plasmid. It works by simply amplifying the whole plasmid, in a non PCR-based thermocycling reaction. During the reaction mutagenic primers, carrying the desired mutation, are integrated into the newly synthesized plasmid. In this video tutorial we demonstrate an easy and cost effective way to introduce base substitutions into a plasmid. The protocol works with standard reagents and is independent from commercial kits, which often are very expensive. Applying this protocol can reduce the total cost of a reaction to an eighth of what it costs using some of the commercial kits. In this video we also comment on critical steps during the process and give detailed instructions on how to design the mutagenic primers.

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. Here we demonstrate an easy and cost effective way to introduce base substitutions into a plasmid using standard reagents.

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. With this method the cloned ...

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 ...

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Conserved nutrient sensing mechanisms exist between mammal and fruit fly where peptides resembling mammalian insulin and glucagon, respectively function to maintain glucose homeostasis during developmental larval stages 1,2. Studies on largely post-mitotic adult flies have revealed perturbation of glucose homeostasis as the result of genetic ablation of insulin-like peptide (ILP) producing cells (IPCs) 3. Thus, adult fruit flies hold great promise as a suitable genetic model system for metabolic disorders including type II diabetes. To further develop the fruit fly system, comparable physiological assays used to measure glucose tolerance and insulin sensitivity in mammals must be established. To this end, we have recently described a novel procedure for measuring oral glucose tolerance response in the adult fly and demonstrated the importance of adult IPCs in maintaining glucose homeostasis 4,5. Here, we have modified a previously described procedure for insulin injection 6 and combined it with a novel hemolymph extraction method to measure peripheral insulin sensitivity in the adult fly. Uniquely, our protocol allows direct physiological measurements of the adult fly's ability to dispose of a peripheral glucose load upon insulin injection, a methodology that makes it feasible to characterize insulin signaling mutants and potential interventions affecting glucose tolerance and insulin sensitivity in the adult fly.

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Conserved insulin signaling pathways found in the fruit fly Drosophila melanogaster make this organism a potential tool for modeling metabolic disorders including type II diabetes. To this end, it is critical to establish physiological assays to effectively measure systemic insulin action in peripheral glucose disposal in the adult fly.

Conserved nutrient sensing mechanisms exist between mammal and fruit fly where peptides resembling mammalian insulin and glucagon, respectively function ...

Isolation and Biophysical Study of Fruit Cuticles

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and abiotic stresses and also regulates water flow from the external environment.1 A biopolyester (cutin) and long-chain fatty acids (waxes) form the principal structural framework of the cuticle; the functional integrity of the cuticular layer depends on the outer 'epicuticular' layer as well as the blend consisting of the cutin biopolymer and 'intracuticular' waxes.2 Herein, we describe a comprehensive protocol to extract waxes exhaustively from commercial tomato (Solanum lycopersicum) fruit cuticles or to remove epicuticular and intracuticular waxes sequentially and selectively from the cuticle composite. The method of Jetter and Sch&auml;ffer (2001) was adapted for the stepwise extraction of epicuticular and intracuticular waxes from the fruit cuticle.3,4 To monitor the process of sequential wax removal, solid-state cross-polarization magic-angle-spinning (CPMAS) 13C NMR spectroscopy was used in parallel with atomic force microscopy (AFM), providing molecular-level structural profiles of the bulk materials complemented by information on the microscale topography and roughness of the cuticular surfaces. To evaluate the cross-linking capabilities of dewaxed cuticles from cultivated wild-type and single-gene mutant tomato fruits, MAS 13C NMR was used to compare the relative proportions of oxygenated aliphatic (CHO and CH2O) chemical moieties.
Exhaustive dewaxing by stepwise Soxhlet extraction with a panel of solvents of varying polarity provides an effective means to isolate wax moieties based on the hydrophobic characteristics of their aliphatic and aromatic constituents, while preserving the chemical structure of the cutin biopolyester. The mechanical extraction of epicuticular waxes and selective removal of intracuticular waxes, when monitored by complementary physical methodologies, provides an unprecedented means to investigate the cuticle assembly: this approach reveals the supramolecular organization and structural integration of various types of waxes, the architecture of the cutin-wax matrix, and the chemical composition of each constituent. In addition, solid-state 13C NMR reveals differences in the relative numbers of CHO and CH2O chemical moieties for wild-type and mutant red ripe tomato fruits. The NMR techniques offer exceptional tools to fingerprint the molecular structure of cuticular materials that are insoluble, amorphous, and chemically heterogeneous. As a noninvasive surface-selective imaging technique, AFM furnishes an effective and direct means to probe the structural organization of the cuticular assembly on the nm-&mu;m length scale.

Aerial plant organs are protected by the cuticle, a supramolecular biopolyester-wax assembly. We present protocols to monitor selective removal of epi- and intracuticular waxes from tomato fruit cuticles on molecular and micro scales by solid-state NMR and atomic force microscopy, respectively, and to assess the cross-linking capacity of engineered cuticular biopolyesters.

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and ...

Fruit Volatile Analysis Using an Electronic Nose

Numerous and diverse physiological changes occur during fruit ripening, including the development of a specific volatile blend that characterizes fruit aroma. Maturity at harvest is one of the key factors influencing the flavor quality of fruits and vegetables1. The validation of robust methods that rapidly assess fruit maturity and aroma quality would allow improved management of advanced breeding programs, production practices and postharvest handling. 
Over the last three decades, much research has been conducted to develop so-called electronic noses, which are devices able to rapidly detect odors and flavors2-4. Currently there are several commercially available electronic noses able to perform volatile analysis, based on different technologies. The electronic nose used in our work (zNose, EST, Newbury Park, CA, USA), consists of ultra-fast gas chromatography coupled with a surface acoustic wave sensor (UFGC-SAW). This technology has already been tested for its ability to monitor quality of various commodities, including detection of deterioration in apple5; ripeness and rot evaluation in mango6; aroma profiling of thymus species7; C6 volatile compounds in grape berries8; characterization of vegetable oil9 and detection of adulterants in virgin coconut oil10. 
This system can perform the three major steps of aroma analysis: headspace sampling, separation of volatile compounds, and detection. In about one minute, the output, a chromatogram, is produced and, after a purging cycle, the instrument is ready for further analysis. The results obtained with the zNose can be compared to those of other gas-chromatographic systems by calculation of Kovats Indices (KI). Once the instrument has been tuned with an alkane standard solution, the retention times are automatically converted into KIs. However, slight changes in temperature and flow rate are expected to occur over time, causing retention times to drift. Also, depending on the polarity of the column stationary phase, the reproducibility of KI calculations can vary by several index units11. A series of programs and graphical interfaces were therefore developed to compare calculated KIs among samples in a semi-automated fashion. These programs reduce the time required for chromatogram analysis of large data sets and minimize the potential for misinterpretation of the data when chromatograms are not perfectly aligned.
We present a method for rapid volatile compound analysis in fruit. Sample preparation, data acquisition and handling procedures are also discussed.

A rapid method for volatile compound analysis in fruit is described. The volatile compounds present in the headspace of a homogenate of the sample are rapidly separated and detected with ultra-fast gas chromatography (GC) coupled with a surface acoustic wave (SAW) sensor. A procedure for data handling and analysis is also discussed.

Numerous and diverse physiological changes occur during fruit ripening, including the development of a specific volatile blend that characterizes fruit ...

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 ...

High-throughput Functional Screening using a Homemade Dual-glow Luciferase Assay

We present a rapid and inexpensive high-throughput screening protocol to identify transcriptional regulators of alpha-synuclein, a gene associated with Parkinson&#39;s disease. 293T cells are transiently transfected with plasmids from an arrayed ORF expression library, together with luciferase reporter plasmids, in a one-gene-per-well microplate format. Firefly luciferase activity is assayed after 48 hr to determine the effects of each library gene upon alpha-synuclein transcription, normalized to expression from an internal control construct (a hCMV promoter directing Renilla luciferase). This protocol is facilitated by a bench-top robot enclosed in a biosafety cabinet, which performs aseptic liquid handling in 96-well format. Our automated transfection protocol is readily adaptable to high-throughput lentiviral library production or other functional screening protocols requiring triple-transfections of large numbers of unique library plasmids in conjunction with a common set of helper plasmids. We also present an inexpensive and validated alternative to commercially-available, dual luciferase reagents which employs PTC124, EDTA, and pyrophosphate to suppress firefly luciferase activity prior to measurement of Renilla luciferase. Using these methods, we screened 7,670 human genes and identified 68 regulators of alpha-synuclein. This protocol is easily modifiable to target other genes of interest.

We present a rapid and inexpensive screening method for identifying transcriptional regulators using high-throughput robotic transfections and a homemade dual-glow luciferase assay. This protocol rapidly generates direct side-by-side functional data for thousands of genes and is easily modifiable to target any gene of interest.

We present a rapid and inexpensive high-throughput screening protocol to identify transcriptional regulators of alpha-synuclein, a gene associated with ...