Equipment#

• Dissection-suitable microscope

• Forceps:

  • Fine, sharp
  • Thicker, blunt
  • ‘Grippy’, fine but blunted

• Small electric heater

• Hypodermic needle

• UV glue + gun

• Optional: custom hook

• Fly holder

  • Recommended: cone-shaped holders for walking behavior and visual field-of-view
    • The design files for the standard, T shaped, and circular (C) cone-holders are attached below
    • Depending on the average size of your fly, you can adjust the slot size by adjusting the thickness parameters
    • Ordering: CNC Machined from Protolabs out of black delrin
    • Note: the very original cone-shaped holder design was provided by the Maimon Lab. They’ve been acknowledged in Fisher et al. If you revert to the original Maimon lab holder design, they should be acknowledged.

Design Files:

Standard holder for small/medium flies

📎 Coneholder_MC_v5.stp

Standard holder for large flies

📎 Coneholder_MC_v5_large.stp

T shaped platform w/magnet slots

📎 Coneholder_MC_v5T.stp

Circular platform

📎 Coneholder_MC_v5C.stp

Previous versions

📎 cone_shaped_holder_for_perfusion.ipt

📎 cone_shaped_holder.STEP

Detailed Procedure#

1. Perform the final cross that gives you your experimental flies. For optimal culture density, there are multiple methods that work:
2. Cross at low density (3♀x 3♂) in molasses food (or german food is the cross isn’t taking well) to encourage the development of large, healthy progeny. Wait ~10 days for the pupae to mature, remove the parents at 25ºC/77ºF.
3. An alternative crossing method is to have (8♀x 3♂) and to flip the vial onto new food every 2-3 days until the parental flies are too old or unproductive.
4. For very specific age-staging, low genotype yields, or less healthy crosses, use 15♀x 10-12♂, and flip onto fresh food after 2 days, or once eggs have started filing the bottle. Flipping a second time is possible. The large number of larvae makes the food easier to grow on, but to prevent overcrowding the cross must be cleared daily or multiple times daily once adults eclose (Stephen’s method for normal flies).
5. Fly selection and preparation:
6. Select virgin females from crosses. If waiting to age further, keep in fresh molasses vials at low density, e.g. 3x flies per vial. We do not want flies laying eggs on the ball, and we would prefer larger animals, so virgin females are the best choice if sex is not an experimental factor. Typically we want experimental flies to be about 1-day old post-eclosion.
7. If applicable, starve the flies before the experiment. Perform starvation in an empty plastic vial with a moist kimwipe as a humidifier, keeping 3-4 flies per vial. The optimal starvation time might depend on the genotype you are using and the weather. 5-8 h starvation might be best for the colder/drier days, where fly stocks can be a bit more unhealthy, and overnight starvation might be best for warmer months.
8. Position fly in holder
9. Anesthetize flies on ice (in a vial).
10. Transfer fly into the holder with blunt forceps. The holder is placed ventral-side-up and the fly is inserted with its dorsal-side-down, legs in the air. Some people place the fly’s head and thorax into the holder; others place the fly thorax-only and then readjust the fly after flipping the holder.
11. Flip holder. The fly should be wedged in place, and so should not fall out. If it does, your fly may be too small or you may have placed the fly too far into the holder initially.
12. Position the head and thorax and then glue the fly into place. Details: 1. Depending on your experiment, you may want different head angles. For calcium imaging of the central complex, for example, you will want a more posterior prep. Electrophysiologists place the head position for optimal access to the cell body. 1. Use UV glue to glue the head and thorax into place. Fill in the space between the fly’s neck area and the sides of the teardrop. Apply glue slowly to prevent seepage. Some people use pipettes to deliver small amounts of glue. Another technique is to let a fly walk on a small Kimwipe piece while applying glue. 1. Optionally remove the wings using fine forceps. Wing removal may assist walking as the fly is better fit into the teardrop and wing motion does not occur.
13. Removing proboscis movement
14. Anesthetize fly using a small piece of ice on the dorsal part of the holder. This is important if you need walking behavior.
15. Remove proboscis movement. There are tradeoffs for each method, but generally severing the lateral muscles of the proboscis and/or removing it completely will reduce brain movement but worsen behavior: 1. Proboscis muscle clipping. Use your fine forceps or a fine gauge needle to clip proboscis muscles until the fly can no longer use it.
    1. Downside: a drooping proboscis may be a walking hazard in some set ups. Cutting the lateral muscles can also worsen behavior
    2. Upside: dramatically reduces proboscis motion (entirely if done perfectly) and is less invasive than proboscis removal.
    3. Note: a video called “proboscis_immobilization_procedure.mp4” demonstrating this technique can be found on the shared lab server in the “manuals, protocols, and databases” directory. 1. Proboscis removal. Use your blunt forceps to pull out the proboscis by the labellum. Use fine forceps to clip away the soft tissue connecting the proboscis to the head. Pull the proboscis away, letting the remaining soft tissue seal the hole in the fly’s face.
    4. Upside: is easier than muscle clipping and removes proboscis motion entirely
    5. Downside: highly invasive, will typically result in worse behavior over time 1. RECOMENDED: Proboscis gluing, using either hot wax or UV glue. Upside: not as invasive as surgery. Extend the proboscis using your blunt forceps, and glue onto the holder surface in front of the fly. Pull the proboscis towards the fly’s abdomen and glue in place, careful not to catch the legs. Glue the proboscis without extension. This requires glue at each point where the proboscis can move.
    6. Downside: can risk getting glue on the eyes and legs. some proboscis motion may still occur, and it my be hard to do this with certain head angles.
    7. Upside: easy and causes no tissue. 1. RECOMENDED: Cut and glue proboscis (combining above methods)
    8. Upside: redundant method to ensure minimal to no brain movement, especially helpful for electrophysiology behavior experiments.
    9. Downside: severing lateral muscles can worsen behavior.
16. Flip the holder so that it is dorsal-side-up and remove ice. Add carbogenated saline into holder, on top of the fly’s head. Saline can be warmed.
17. Remove cuticle
18. If doing a posterior prep, at the posterior of the fly’s head is a ‘dark square’. This is the entry hatch to the fly’s brain! With your hypodermic needle first cut the cuticle along the ventral-most edge, then the sides, then with your fine forceps, lift the hatch off and away.
19. Clear away any internal fat and trachea obscuring the brain. You should have a clean, gently pulsing brain when you finish.
20. Clip muscle 16, (also see here) which causes high frequency brain oscillations. You can either:
21. Take a custom-made ‘hook’ and insert it round the anterior side of the brain. Angle it, and push the brain with it gently to reveal the muscle. Once you see it, hook the hook around the muscle fiber and yank it free. Be aware that to either side of muscle 16 are the antennal nerves.
22. At a posterior head angle, gently tug the esophagus with medium-fine forceps. Muscle 16 is directly connected to the esophagus on the proboscis-side of the head. Beware that the VNC runs directly underneath the esophagus.
23. Avoid damaging the antennal lobe connections that run on either side of the esophagus! Damaging these will often result in poor behavior. This is perhaps more relevant for folks who do anterior preps.
24. If you are not sure what these connections look like, you should be able to tell because any movement inflicted onto the connection tract should also move the corresponding antennal lobe of the brain.
25. If doing electrophysiology, use a very fine pair of grippy forceps to de-sheath where you expect to find the cell body.
26. Ask someone if you don’t already know how to do this step.
27. Your fly is now ready. You can let the fly ‘walk’ on a piece of tissue, and observe that it spins the tissue or shifts it about a little, and is therefore still active.