Laboratory Maintenance of the Lower Dipteran Fly Bradysia (Sciara) coprophila: A New/Old Emerging Model Organism

The protocols described here represent a century of experience from the Bradysia (Sciara) stock centers overseen sequentially by Charles Metz, Helen Crouse, and Susan Gerbi as well as input from others.

1. Mating crosses

1. Use one adult female and two adult males per 28 mm diameter glass vial. Once 100% success is achieved in recognizing wing phenotypes for mothers who will have just daughters or sons, use two females and two males per vial to increase the number of larvae/vial. Add females to each vial before adding the males who wake up faster from the anesthesia than the females.
   NOTE: The description below presumes that you have CO₂; alternatively, ether can be used to anesthetize the adult flies.
2. If using ether to anesthetize the adult flies, tape a pad of folded laboratory wipes (e.g., kimwipes) to the inside of a round glass lid of a Coplin jar and use a dropper to transfer some ether from the bottle to moisten the laboratory wipe pad (but not saturated and so wet that ether liquid will fall off it and risk drowning the flies). For step 1.6 below, place the moistened ether pad on top of the opened vial for ~1 min until the adults stop moving. Once the adults have been transferred to a white ceramic tile plate (used instead of the white fly-pad), periodically (when the flies' legs begin to twitch) hold the pad that has been freshly ether-moistened over (not touching) the flies on the plate for ~1 min.
   CAUTION: Ether is flammable and should be stored in a ventilation hood and not in a refrigerator.
3. Arrange within arm's reach a tray with vials of adult females, a tray with adult males, and a tray of empty vials containing 2.2% (wt/vol) agar. On the lab bench, place placard notes stating "female" or "male" and the wing phenotype of the mother so that the vial with adults selected for the cross will be placed into the correct group, where all vials in one group will have just male progeny and all vials in the other group will have just female progeny.
   NOTE: Be sure that no condensation droplets are within the vial as the adults will stick to the droplets. If the vials were stored in a plastic box, put the vials on a lab bench at room temperature for at least 1 h before use to let the condensation evaporate.
4. Turn on the CO₂ gas and the lamp for the dissecting microscope.
   NOTE: A light source with fiber optics is preferred as it emits less heat, which causes the anesthetized flies to wake up faster.
5. Vigorously tap the vial with adults on a rubber pad so that the adults fall to the bottom of the vial and remove the plug; insert the CO₂ gun nozzle and add back the plug.
6. Depress the nozzle trigger so that CO₂ flows into the vial for ~1 min to anesthetize the adults.
7. Put a foot on the foot pedal so that CO₂ will flow onto the white fly-pad rather than the gun nozzle. Keep the foot pedal depressed the entire time that flies are on the white fly-pad (or else intermittently depress the foot pedal whenever the flies' legs begin to twitch).
8. Remove the nozzle and plug from the vial and invert the vial over a white fly-pad under a dissecting microscope.
9. Tap the bottom of the inverted vial against the microscope so that the adults fall from the vial onto the white fly-pad.
10. Select the fattest adults (recently eclosed with white abdomens) and use fine-tipped forceps to gently pick up the adult by its middle or hind leg. Do not injure the front legs that are used for the mating dance. Do not use adults that have just eclosed and whose bodies are fully white and not yet turned black because their wings will be short and not fully developed so that the wing phenotype cannot be scored. Adults with slimmer abdomens can still be used, though they have reduced fertility. Do not use skinny adults whose wings are raised vertically away from their body, since they are dead.
11. With the other hand, remove the plug from the vial with 2.2% (wt/vol) agar. With the hand holding the forceps with the adult, tap the forceps vigorously against the top inside wall of the vial so that the adult falls to the bottom of the vial. Replace the plug into the vial.
12. Repeat steps 1.5-1.11 above to set up each vial containing female adults. Use a paintbrush to sweep the unused flies from the white fly-pad back into their parent vial. Put a checkmark on the label of the parent vial to indicate that it has been used for mating (though it can be used again if needed).
13. For routine stock maintenance, set up 6-8 vials with female-producer mothers and 6-8 vials of male-producer mothers (Figure 4). Set up half of the vials with mothers (or fathers) from one adult vial and the other half of the new vials using a different adult vial to minimize genetic bottlenecks.
    1. Occasionally, a mis-segregation event will produce an exceptional male in female-producing vials. If a vial with adult females has an exceptional male, remove and squish to kill that male. If possible, discard all females from that vial and wait a few days until more adult females eclose and can be safely used for crosses.
       NOTE: It is preferable to not use the females in that vial for crosses because they may have mated with the exceptional male and not produce a fertile cross.
14. Once the adult females have been added to all the vials, repeat steps 1.5-1.11 to add two adult males (Figure 4) to each vial already containing female flies. Tap the vial with females on a rubber pad so that they do not escape when the two males are added sequentially.
    NOTE: Work quickly and do not overanesthetize the adult flies as that will kill them.
15. Add a label to each vial stating the stock, the cross (for female or male progeny), whether the mother came from adult vial #1 or #2, and the date of mating. Also enter the information above into a notebook and state the number of vials set up for each cross.
    NOTE: It is convenient to add a paper towel to separate the male-producing vials from the female-producing vials in the tray.
16. Leave the vials undisturbed on the countertop for ~15 min to be sure that the adults wake up and fly around. Observe if they are mating (very soon after they wake up) where the female and male are oriented posterior to posterior (the male clasper will grab the female's pointed ovipositor) (Figure 4, bottom).
    NOTE: An adult female will accept an adult male just once, so do not jostle the vials after mating as this could separate the male from the female while in the process of mating and that female will not re-mate.
17. Place the tray with mated vials in the incubator (e.g., 21 °C). Label the tray with the name of the stock (e.g., HoLo2) and the week of the 5 week cycle (weeks 1, 2, 3, 4, or 5).
    NOTE: Keep the tray with just-mated flies in a separate part of the incubator or another incubator as a reminder to not feed the vials until the larvae have emerged (described below).

2. Mass mating

NOTE: Typically, a single B. coprophila mother will have 60 offspring in her brood. When a larger number of offspring is required for experiments, a mass mating can be done instead of the single-pair mating described above. The mass mating can be done in the standard 28 cm diameter glass vials when the mothers will be collected a day later for induced egg-laying and embryo collection. However, if a larger number of larvae are needed, the mass mating is done in a jar with a larger surface to prevent overcrowding. Pierce several small holes in the lid of the jar so that the larvae can get some air to breathe.

1. Follow the steps above in section 1 but use fine-tipped forceps to move all female-producer mothers (or all male-producer mothers) to a front corner of the white fly-pad. Use 10-15 anesthetized fat adult females and sweep this group with a paintbrush into the vial or jar with small holes pierced in the lid.
   NOTE: Perform steps 2.1 and 2.2. quickly for mass mating to avoid over-anesthesia that would prevent fertile crosses.
2. Select 20-25 anesthetized fat adult males and move them with fine-tipped forceps to the front corner of the white fly-pad.
3. Vigorously tap the vial or jar with the females on a rubber pad so that they do not escape when the plug or lid is removed to sweep in the anesthetized cluster of adult males from the white fly-pad using a paintbrush.

3. Embryo collection after mass mating

1. One day (24 h) after mating, perform steps 1.5-1.11 to anesthetize the adult flies (mixture of females and males) and transfer them to a white fly-pad.
   NOTE: Oogenesis is not yet completed when the adult flies mate, which triggers finishing meiosis; the mature eggs are fertilized by sperm stored in the spermatheca when the eggs are discharged ²⁹. Completion of oogenesis can take 1-2 days, which is why 1 day is allowed after mating before egg-laying is induced.
2. With fine-tipped forceps, pick up an adult female by her wings and place her on a 100 mm diameter Petri dish containing 2.2% (wt/vol) agar, with her wings inserted into the agar. Repeat this step sequentially for each adult female on the white fly-pad. Discard the adult males on the fly-pad.
3. Once all the female flies are impaled on the agar, induce egg-laying by gently squeezing the head with forceps until the female fly has seizure-like movements. Alternatively, gently squeeze her thorax. She will then lay a cluster of fertilized eggs within 30-60 min. Cover the Petri dish with its lid that is moistened with a water-dampened laboratory wipe to prevent electrostatic attraction of the eggs to the lid.

4. Checking "hatch" of larvae

1. Check for "hatch" of larvae 1 week after mating; remove the plug from the vial and use a dissecting microscope to score for larvae. Their black jaw will open and close repeatedly and they will slowly move forward. If there are only a few larvae, write "few" on the vial label, as a reminder to feed that vial less. Examine each vial in the tray with that cross and enter the number of vials with larvae in the notebook in a column next to the number of vials that were set up in that mating.
   NOTE: Deep yellow eggs will not develop. White eggs are likely to develop and 1 day before the larvae emerge, black pigment (the future jaw) will develop at the anterior end of the egg. Do not mistake mold with a white filament ending in a black sphere at its end for larvae-the mold will not move, unlike the larvae that crawl forward on the agar.
2. Add a small bit of straw (just once) to each vial with larvae to control excess humidity and provide a hiding place of cover for the larvae.
3. Move the tray to the incubator (e.g., 21 °C) with trays of larvae from previous weeks' mating crosses and begin to feed the vials with newly emerged larvae (see section below on feeding).
   NOTE: Generally, the larvae will be in a cluster near their dead mother and they will begin to eat her; presumably this transfers the yeast from the mother's gut to the gut of the larvae. You can begin feeding on the day that the larvae emerge or within 2 days thereafter. CAUTION: If feeding is delayed, the larvae will eat each other, with only one fat larva remaining per vial!
4. Continue to check vials that did not yet have larvae for 3 days per week. If no larvae have emerged after 7-10 days, discard the vial or store it for vial washing.

5. Making the food

NOTE: All the food ingredients should be pesticide-free!

1. Use a tablespoon to measure the following ingredients by volume and deposit them into a metal or glass pan (e.g., an 8 inch x 8 inch metal baking pan): 4 parts oat straw (8 Tbsp), 2 parts Shitake mushroom powder (4 Tbsp), 1 part spinach powder (2 Tbsp), 1 part nettle powder (2 Tbsp). Use the tablespoon to mix the ingredients well in the pan.
   NOTE: Alternatively, 2 parts of just spinach powder or just nettle powder can be used instead of 1 part of each.
2. Cover the pan with aluminum foil and autoclave on a dry cycle for 20-30 min. Let it cool to room temperature overnight or longer.
3. Remove the foil from the pan and break up the caked sterilized food mixture, using a grinding motion with the tablespoon to create a powdery mixture.
4. Add 1 part (2 heaping Tbsp) of Brewer's yeast and mix well into the autoclaved food mixture.
   NOTE: The Brewer's yeast is not autoclaved as that would kill the yeast.
5. Transfer the food mixture to a sterile capped jar.
   NOTE: The same type of 240 mL jar as used for mass mating can be used, and the recipe above will fill the jar. Similarly, the same type of sterilized capped jar should be filled with just straw that was autoclaved in a foil-covered pan.

6. Feeding

NOTE: Adjust the amount of food given according to the age and amount of larvae. Give a lot of food to jars with many larvae from a mass mating. Give just a light sprinkle of food to Petri plates with larvae that are being stored for a few days for developmental staging. The feeding method described below was developed in the lab of Charles Metz ²⁹, and it has been used by his lab and by Helen Crouse, Susan Gerbi, and others successfully for a century.

1. Wash hands and rinse well to remove soap.
   NOTE: Gloves are not recommended as they diminish the feel of your fingers for regulating the amount of food given to each vial.
2. Store a capped sterile jar with ground straw and a sterile jar with food in the incubator (e.g., 21 °C) where the larvae are kept. Remove the lid from the jar and pour some food into a clean bowl (e.g., a candy dish) for easier access; replace the lid on the jar containing the remaining food. Keep the open bowl with food in a metal or glass small tray; this will inhibit mites or other insects from crawling over the wall of the tray to enter the bowl with food.
3. Pick up some food between the second and third fingers (or between the thumb and second finger). With the other hand, remove a vial from the tray and remove the plug, holding it with fingers while feeding. Examine the vial for the age and number of larvae and deposit the appropriate amount of food into the vial by rotating the two fingers holding the food against one another. Vials with newly emerged larvae just need a few grains of food. Vials with older larvae should have a thin layer of food covering the top of the agar.
4. If there is white mold in the vial, use 70% ethanol sprayed on a laboratory wipe to wipe clean a long metal probe (e.g., with a wooden handle), remove the plug, and insert the clean probe into the vial to tap down the mold on the surface of the agar. If there is a lot of mold, swirl the probe to wrap the mold around the probe to remove it from the vial. Do not disturb the top of the agar as larvae are living there; add just a small amount of food and replace the plug in the vial. Wipe the probe clean with a laboratory wipe dampened with 70% ethanol before storing the probe or using it to clean another vial.
5. Once all vials in a tray have been fed, replace the tray in the incubator and remove the next tray for feeding as in step 6.3.
6. After feeding is completed, pour the remaining food from the bowl into the previously sterilized jar, cap the jar, and store it in the incubator.

7. Collection of larvae or pupae on Petri plates

1. For small numbers of larvae, use a metal probe or spatula that has been wiped with 70% ethanol to dig into the top layer of agar in a vial to transfer the adhering agar with some larvae to a 100 mm diameter sterile Petri dish half-filled with 2.2% (wt/vol) agar.
2. For larger numbers of larvae, insert a spatula along the wall at the bottom of the vial to ease the agar plug out of the vial and deposit it face-side up in an empty Petri dish. Use a dissecting microscope to find the larvae at the top of the agar plug and transfer them with fine-tipped forceps to a 100 mm diameter sterile Petri dish half-filled with 2.2% (wt/vol) agar.
3. Use a dissecting microscope to sort the larvae with fine-tipped forceps into groups of the same developmental stage.
   NOTE: Due to slight developmental asynchrony, there will be several different clusters of sorted larvae on the Petri plate with 2.2% (wt/vol) agar.
4. Add a tiny sprinkle of food and place the Petri dish with larvae in the incubator. Remove it daily for observation with the dissecting microscope for selection of the desired developmental stage.
   NOTE: For DNA puff studies, early eyespot larvae proceed through 10×5, 12×6, 14×7, and edge eye/drop-jaw stages with ~1 day at each of these stages ^30,31. Choose pupae whose eyes are ¼ to ½ filled with pigment to have meiosis I and II stages in the pupal testes.

8. Cold room storage of larvae

1. As a backup, keep in the cold room about four vials of female larvae and four vials of male larvae from 2 consecutive weeks of crosses.
2. For the backup storage, feed larvae that are early 4^th instar (pre-eyespot stage) and put them in an open tray on a shelf in the cold room; feed these vials just once a month.
3. Remove the 16 vials from the 2 consecutive weeks of mating crosses from the cold after 2-3 months (mark this on a calendar as a reminder); place them in an incubator (e.g., 21 °C) to feed them normally and let them develop into adults to use for crosses.
4. When the vials are removed from the cold room, put a fresh set of vials with early 4^th instar larvae in the cold room so that these vials are always available as a backup for stocks.
   NOTE: Viability after cold room storage has not been tested systematically for different developmental stages, but our experience reveals that cold storage of early 4^th instar larvae works well.

9. Vial washing

1. After the adults have died (~2-3 weeks after eclosion), remove the vials from the incubator and place them at 60-70 °C for 1 hour to kill any remaining organisms.
2. Remove the cotton plugs and store them in a plastic box.
3. Use a spatula to scrape the agar plug from the vial into a waste can.
4. Soak the vials overnight or longer submerged in water in a dishpan.
5. Use a test tube brush saved just for this purpose (never used for vessels with chemicals nor with soap) and scrub up and down in the vial under running tap water. Place the cleaned vial open side down in a metal basket. Fill the basket with cleaned vials.
   NOTE: Never use soap in the vials as any residual soap could kill B. coprophila.
6. Add a wire mesh lid to the basket, and while holding the lid in place, invert the basket to fill all the vials with deionized water. While still holding the lid in place, invert the basket to empty the deionized water from the vials. Repeat the deionized water rinses 4x.
7. Place the baskets with cleaned vials on paper towels or an open mesh platform (e.g., lab cart) to dry overnight or longer. Store the dry vials in a drawer.
   NOTE: Do not let vials with dead adults remain too long before washing them as it could invite an infestation of mites.

10. Pouring agar

1. Fill a large container (metal basket or metal tray) with clean vials and add a cotton plug to each vial. Reuse the plugs taken from old vials when the vials were washed, and use the plugs repeatedly until they degrade and fall apart and should be discarded. Sterilize the container with the plugged vials on a dry cycle for ~30 min and let them cool to room temperature to store them in the autoclave container. Always keep two containers (one actively in use and one spare) with autoclaved plugged vials on hand.
2. Place 11.0 g of agar powder in a 1 L Erlenmeyer flask and add 500 mL of distilled water to make a 2.2% (wt/vol) agar solution sufficient to pour ~24 vials (~21 mL/vial). Swirl the flask and place it in a microwave oven.
   NOTE: From here on, use a heat-resistant autoclave glove to handle the flask. The same 2.2% (wt/vol) agar solution can be poured into Petri plates (for picked larvae) or jars (for mass matings) if these are needed.
3. Microwave for 1 min, remove the flask to swirl it, replace it in the microwave oven, and microwave it again for 1 min. Repeat this several times.
4. Watch the flask through the glass door of the microwave oven. When the agar solution begins to foam and boil up, immediately use the manual stop button. Do not swirl the flask at this point (it could boil over), and carefully remove it from the oven to the counter and let it rest for a few minutes.
5. Remove the plug from a sterilized vial and hold it with one hand; use the other hand to pour ~2.5 cm high 2.2% agar into the sterilized vial. Then, replace the plug into the vial. Repeat this until all the agar has been poured.
   NOTE: If too little agar is poured into the vial, it will dry up more quickly during use and shrink away from the walls of the vial. If too much agar is poured into the vial, it is hard to focus on the top layer when scoring the "baby-hatch" of larvae with a dissecting microscope.
6. Let the poured vials remain on the counter at room temperature for 1-2 days so that the agar hardens and the moisture evaporates completely. Use the vials for mating crosses or store them lying on their side in a plastic box with a tight-fitting lid; place a paper towel dampened with water on top of the vials before placing the lid on the box (this will prevent the agar from drying out and shrinking away from the wall of the vial). Store the box with poured vials at room temperature for a few days or in a 4 °C refrigerator or cold room for up to 2 weeks. Before using the stored vials, remove them from the box and leave them on the counter at room temperature for an hour or two to let the condensation evaporate from inside the vials (the adult flies would stick to the condensate and drown).

11. Making plugs for vials

1. Place a clean vial in a styrofoam 50 mL test-tube holder (force the tube in so it stands up straight). Cut a square of cheesecloth (2-4 layers thick) and place it on top of the vial.
2. Place scraps of cheesecloth from previously made plugs on top of the cheesecloth to create a thicker layer. Place cotton on top of the cheesecloth and press down into the vial, making sure to tightly fill the top inch or so of the vial.
3. Hold the top of the ends of the cheesecloth together and tie it with string. Cut the excess string and excess cheesecloth (leave enough tails of the string to keep it tied and enough cheesecloth to comfortably grasp the plug).
   NOTE: Fit plugs so that they are snug. You should be able to pull them out with your index and forefinger while you hold them in one hand. If you pick up the vial by the plug and it falls out, it is too loose. If it makes a loud popping sound when pulled out, it may be too tight. If too tight or if the plug has a layer of hard, dry agar on it, the flies may suffocate. If the plug is a little too loose or slipping down the sides of the vial, it can be tapped on the countertop to squash it into a slightly wider size.

12. Typical weekly schedule (for greatest efficiency, perform tasks in the order listed)

1. Monday (~30 min)
   1. Feed vials with larvae.
   2. Pour agar into sterile clean vials with plugs.
2. Wednesday (~2 h)
   1. Place vials with dead adults in an oven for 1 h and store for washing.
   2. Check for larvae "baby-hatch".
   3. Feed vials with larvae.
   4. Perform weekly mating crosses.
   5. Other duties as needed: autoclave clean vials with plugs to have a reserve supply as needed; make new food and autoclave straw as needed; autoclave jars as needed for food and straw.
3. Friday (~30 min)
   1. Check for larvae "baby-hatch".
   2. Feed vials with larvae.