There are many different ways to fix Drosophila embryos that are often specific for different downstream applications. While most applications use Formaldehyde or Para-formaldehyde - other methods like methanol or heat fixation can also be used. Here, I consolidate several fixation protocols that have been used successfully in the Eisen lab.
All fixation protocols have three parts. Part 1 is the fixation. Part 2 is the devitilination. Part 3 consists of the final washes and long term storage. Many of these protocols are fairly similar to one another but have been optimized for various purposes. Like virtually all protocols, not all of the differences between methods have been thoroughly tested. My aim here is to present the various flavors of fixation methods that have been explored by the lab.
*** WARNING: Formaldehyde, Heptane, hexane, and methanol are very hazardous and should not be used outside the hood. Please read the SDS for each chemical thoroughly (I AM NOT KIDDING - DO IT!!!) before using. If any is spilled, notify a lab member immediately and do not panic. Always wear gloves, safety glasses, and a lab coat and change gloves often. While wearing gloves, do not touch anything that you would normally touch without gloves on (like your phone!!!).
This protocol is based off of this protocol adapted for use in our lab by Ashley Albright (09/20/18). She used this fixation method for RNA in situ hybridizations.
- 1.5ml eppendorf tubes
- Drosophila cages with appropriate plates
- embryo collection sieve
- embryo sieve
- paintbrush
- rotating shaker
- water-tight scintillation vials
- 37% formaldehyde
- heptane
- bleach
- 1.3x PBS - made from 10x PBS
- methanol
- Collect embryos in choice of cage. We have used medium cages, small cages (we call them baby cages), apple juice "cup" cages (via Colleen Hannon), or a population cage. Age embryos to the desired stage.
- When ready to collect, depending on the plate used, add 50% bleach to the embryos for 2-3 minutes.
- For baby cages with grape plates - rinse embryos directly into a collection sieve with water. Then add bleach to a deep petri dish or a pipette tip box and submerge the sieve into the bleach.
- for molasses plates or apple juice plates with a descent holding capacity - add 50% bleach straight to the plate. Use a paintbrush to dislodge the embryos and dissolve the yeast into solution.
- Pour bleach solution containing embryos and yeast into the collection sieve.
- Rinse embryos with water for several minutes until the basket no longer smells like bleach.
- Use a paintbrush to transfer embryos to a labeled scintillation vial containing 3 mL 1.3 X PBS.
- In a chemical hood, add 4 mL heptane and 1 mL of 37% formaldehyde to the vial.
- Shake at an angle on a shaker for 20 minutes, ~150 RPM (time sensitive).
- Devitellinize embryos by adding 10 mL methanol and vortexing for ~30 seconds. The embryos should fall to the bottom of the tube. The ones that float were either not devitellinized or did not fix correctly. Sometimes this is a sizeable percentage - I have never been able to figure out why.
- Take off the methanol and floating embryos, being careful to leave all the embryos that fell to the bottom of the vial. 10. Add 1 ml of methanol and pipette embryos with a cut 1000ml tip to a labeled eppendorf tube.
- Was 3x with 1ml of methanol.
- Store at -20˚C in methanol. Embryos can keep for > 1 year stored this way.
This protocol was developed for IF and DNA FISH where small numbers of embryos are collected for each condition. Ampules of 16% Paraformaldehyde were used instead of the bottle of 37% formaldehyde. Paraformaldehyde is essentially the powdered polymer that, in solution, breaks down into the monomeric formaldehyde (more on the differences here). Formaldehyde can be substituted. I performed DNA FISH with this protocol substituting Formaldehyde for the PFA and saw no difference.This protocol also uses triton in the fixation mixture so that the embryos don't stick to the sides of the tube. Therefore, this fixation needs to be gently rocked. Some antibodies stain better in formaldehyde fix or methanol-only fixes depending on the antibody.
- 1.5ml eppendorf tubes
- Drosophila cages with appropriate plates
- embryo collection sieve
- embryo sieve
- paintbrush
- rocker plate - example
- 16% methanol-free paraformaldehyde
- heptane
- bleach
- 10x PBS
- Triton X-100
- methanol
| Final conc | stock conc | for 1ml | |
|---|---|---|---|
| PBS | 1x | 10x | 100ul |
| Triton X | 0.1% | 100% | 1ul |
| Paraformaldehyde | 4% | 16% | 250ul |
| H20 | 640ul |
- Clear for <30 minutes. Embryos collection for 2:00 and aged for 1:20. These times are to collect mostly embryos before nuclear cycle 14.
- Embryos are bleached for 2-3 minutes in 50% Bleach to dechorionate. Rinse well with water. Pat dry on towel.
- Rinse with 1xPBS + 0.5% Triton.
- Take a large brushful of embryos and place them in a 1.5ml tube filled with 1xPBS + 0.5% Triton.
- Take off and wash with H20.
- Take off supernatant and wash with 100% isopropanol.
- Add 500ul of Heptane + 500ul of fixative solution in an eppi
- Fix for 30 minutes with gentle rocking on the rotator plate.
- Remove the lower aqueous layer and replace with 750ul of methanol and vortex for exactly 30 seconds to remove the vitelline membrane.
- TIP: Embryos that sink have their vitelline membrane removed; those that stay at the interface still have their vitelline membranes.
- Remove as much heptane and methanol as possible. Replace with fresh methanol and wash rapidly (10 seconds).
- Add 1ml of fresh methanol and keep O/N at 4˚C.
This fix protocol was used for fixing large batches of embryos for ChIP-seq by Xiao-Yong. Importantly, this protocol does not have a methanol devitalination step (probably because embryos are going to be homogenized for chromatin immunoprecipitation)
- 250ml bottle
- Stir bar
- 50 ml conical
- collection sieve
- Drosophila cages/collection plates
- Hexanes
- 37% formaldehyde
- 10x PBS
- Triton X-100
- Isopropanol
- Shaker
| Final conc | stock conc | for 10mls of top layer | |
|---|---|---|---|
| PBS | 1x | 10x | 27.5ml |
| Formaldehyde | 37% | 37ml | |
| Hexane | 210.5ml |
- Prepare the crosslinking solution by stirring with a stir bar for 30 minutes. Take off the top layer for crosslinking.
- Collect embryos into sieve. Wash with water.
- Dechorionate for 2 minutes in 50% bleach
- Completely immerse the embryos in a beaker of 100% isopropanol and shake back and forth to disperse clumps. Remove cup and dry embryos as much as possible with paper towels beneath the mesh.
- Transfer embryos with a paintbrush to a 50ml Falcon tube containing 10 mls formaldehyde saturated hexanes per gram of embryos (I often used 30mls when using population cages). Shake vigorously for 5 minutes at room temperature. Allow the embryos to settle and pour off as much of the solution as possible (in the hood).
- Resuspend embryos in 40mls of 1X PBS, 0.5% Triton X-100 and shake vigorously at room temp for 5 minutes.
- Pour embryos onto small nylon mesh tube and repeat with fresh buffer. Embryos should clump at first but then eventually become monodispersed.
- Pour all of the embryos in PBST into a small mesh to collect them, then use a paintbrush to transfer them to an Eppendorf tube containing 1ml of 1X PBS, 0.5% Triton X-100 - these can be kept for about a week or so at 4˚C. Alternatively, you can transfer embryos to a tube (dry) and then flash freeze in liquid nitrogen and keep at -80˚C.