Simple Homemade Tools to Handle Fruit Flies—Drosophila melanogaster

Summary

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.

Abstract

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.

Introduction

The fruit fly, Drosophila melanogaster, is a model organism widely used in biological research and biology education to study a wide range of topics^1,2. 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 mutants^3,4) 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 manipulation^8,9. 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.

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Protocol

1. Preparing Tools and Accessories

1. Tip/funnel stoppers
   1. 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.
   2. 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).
   3. 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 μL 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 mm) with small internal stem opening diameters (<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.
2. Microdissecting needles
   1. 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.
   2. 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 0.5–1 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.
3. Hard icepacks
   1. 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.
   2. 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 °C freezer; and plastic film, which is used to cover the stage of a stereomicroscope.
4. 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.
   1. 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.
   2. 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)].
5. Cordless tube brush driver
   1. Obtain a cordless drill driver (max speed = 500 rpm).
   2. 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.

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.

1. 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.
2. Tap down the flies in vial A and replace its plug with a T-stopper, then plug vial B with an F-stopper.
3. 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.
4. Separate the T-stopper from the F-stopper. Cap the pipette tip end of the T-stopper with a 200 μL 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.

3. Immobilizing Flies by Chilling

1. Keep hard refreezable icepacks in a -20 °C freezer for at least 24 h before use.
2. 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.
3. 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.
4. 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.
5. 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.

4. Killing Adult Flies for Counting, Sorting, or Discarding

1. Transfer adult flies into an empty vial and cover it with a Petri dish.
2. Invert the vial, heat it for 1 min + 20 s in a microwave oven, and allow the dead flies drop into the Petri dish.
3. 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.
4. To kill unwanted flies, heat the flies for 2–3 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.

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.

1. 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.
2. Place as many flies as possible into the cage and replug the cage with a foam stopper after transferring the flies.
3. To change the food for flies in the cage, transfer the flies in the cage to an empty vial.
4. Replace the apple juice plate and reseal it, then transfer the flies from the vial back to the cage.
5. When egg collection ends, transfer the flies into an empty vial and transfer them into culture vials.

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.

1. Remove any permanent marker ink from the glass culture vials with wet, stainless steel sponges.
2. Soak the culture vials in running water.
   1. Fill a laboratory sink with water, add liquid dishwashing soap into the water, and mix.
   2. 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.
   3. Soak the old culture vials in water for at least 30 min.
3. 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.
4. Clean the culture vials.
   1. Clean the culture vials roughly.
      1. Put a long cuff rubber glove on the non-dominant hand and hold the vial in the water.
      2. 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.
      3. Dump the waste into a separate garbage can. Repeat this process until most of the waste in each vial has been cleaned.
   2. Clean the culture vials thoroughly.
      1. Clean the tube brush, drain and clean the sink, and refill it with clean water.
      2. Remove the remaining waste from each culture vial as described in section 6.4.1.

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Results

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...

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Discussion

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...

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Materials

Name	Company	Catalog Number	Comments
A pair of pliers
Cordless drill driver			max speed: 500 rpm
Electric soldering iron
File
Funnel			diameter of disk<60mm
Ice box
Insect pins
Infrared thermometer			HCIYET HT-830
Long cuff rubber gloves
Mechanical pencils
Medical gauze
Microcentrifuge tube			100 ul
Microwave oven
Parafilm
Peri dish			internal diameter 60 mm
Pipette tips			1 ml
Plastic film
Plastic Peri dish			Φ36 mm used to cover the empty vial
Point tweezers
Protective work gloves
Re-freezable hard icepacks			26.5×14.5×2.5 cm or larger
Rubber air blower
Snap cutter
Soft drink bottle			500 ml, internal diameter c.a. 65 mm
Sponge stopper
Stainless steel sponges
Tube brush
Vial			Φ34 mm × 90 mm