200424-IFprotocol.md

Title : Immunostaining in Drosophila embryos

Author: Jenna Haines

Date: 07/16/18

Introduction:

This protocol is adapted from Brand 1999, Drosophila Protocols (CSH) 2000, and from Colleen Hannon for Double staining. I have included notes from an example experiment for reference. The fixation protocol is three days long (not including the collection and fixation). Days 2 and 3 can be combined but that would be a very long day. All washes are at room temperature except overnight washes or incubations which are at 4˚C. This protocol includes a fix with paraformaldehyde in a solution containing Triton. The Triton makes it easier to use plastic tubes but experiments comparing paraformaldehyde, formaldehyde, and heat fixation showed little difference for both the anti-Zelda and anti-Bicoid antibody so any other preferred fixation method is fine.

If performing a double staining, make sure that primary antibodies are from different host species so that the secondary antibodies do not detect the wrong antibody. Do single stainings of each primary before or concurrently with the immunostaining.

Materials

• 1.5ml eppendorf tubes
• embryo seive
• paintbrush
• rotator plate (rocker)
• Primary and Secondary antibody
• confocal microscope
• aluminum foil
• clear, fast-drying nail polish
• sissors
• cover slips and glass slides for microscopy

Solutions

Fixing solution

	Final conc	stock conc	for 1ml
PBS	1x	10x	100ul
Triton X	0.1%	100%	1ul
Paraformaldehyde	4%	16%	250ul
H20			640ul

PBT - wash buffer

	Final conc	stock conc	for 500 mls
PBS	1x	10x	50ml
Triton X	0.1%	100%	500ul
H20			450ml

PBTB - blocking buffer

	Final conc	stock conc	for 5 mls
PBT			5ml
BSA	1%	100%	50mg
H20			450ml
** make fresh every time and keep at 4˚C when not using

Protocol

Collection.

1. 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.
2. Embryos are bleached for 2-3 minutes in 50% Bleach to dechorionate. Rinse well with water. Pat dry on towel.
3. Rinse with 1xPBS + 0.5% Triton.
4. Take a large brushful of embryos and place them in a 1.5ml tube filled with 1xPBS + 0.5% Triton.
5. Take off and wash with H20.
6. Take off supernatant and wash with 100% isopropanol.

Fix.

7. Add 500ul of Heptane + 500ul of fixative solution in an eppi
8. Fix for 30 minutes with gentle rocking on the rotator plate.
9. 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.
10. Remove as much heptane and methanol as possible. Replace with fresh methanol and wash rapidly (10 seconds).
11. Add 1ml of fresh methanol and keep O/N at 4˚C.

Immunofluorescence

IF day 1.

12. Remove methanol and rehydrate in PBT (PBS + 0.1 Triton).
13. Wash 1x with PBT
14. Wash 2x5 minutes with 1ml PBT at RT on a rocker
15. Wash1x30 minutes in 1ml PBT at RT on a rocker
16. Resuspend in 100ul of PBTB + Block for 30 minutes at room temperature on a rotor.
17. Divide into tubes per sample.
18. Add primary antibody(S) to each sample (see below). Rock overnight (O/N) at 4˚C.

IF Day 2.

19. Rinse embryos 3x with 1 ml PBT.

    • TIP: the primary antibody used for the incubation can be recovered and saved for further use. Antibody can be used several times.

20. Wash 6x20 min in 1 ml PBT

21. Block for 30 min at room temperature in 100 μl PBTB

22. Add diluted secondary antibody- wrap tube with alumina foil, rock slowly at room temperature for 2 hrs.

23. wash embryos three times for 5 min each with PBT

24. Wash the embryos six times for 20 min (or 4x30 min) each with PBT. * Usually I do the last wash O/N at 4˚C

IF Day 3. Counterstain with DAPI and mount

25. Add 1ul of Dapi to the 1ml PBT.
26. Incubate for an 5 minutes.
27. Wash 6x in PBT
28. Add 100ul of Dabco and let incubate at RT (STANDING) for at least 30 minutes out of the light.
29. Mount slides. Keep at -20.
30. When ready to mount embryos, bring the embryos to room temperature. Transfer 25-30 ul of embryos to a clean slide, place two coverslips on both sides of drop, about 1 cm apart, place a third on top.
    • TIP: Alternatively, to image with the 63x objective on the confocal simply place one coverslip over about 25ul of dabco/embryos and then seal the edges with clear quick-dry nail polish. Make sure to use an eyedropper to drop nailpolish around the edges (not brush) so that there is a thick seal.
31. To save the slide, completely fill the space underneath the coverslip with glycerol and seal all the edges of the coverslips with fingernail polish. The slide can be kept at -20˚C in the dark for many months.

Image on confocal

Conclusion

I used this protocol for double and single antibody experiments. To perform double staining, add both antibodies at the same time in the incubation steps. When doing this, it is important to make sure that there isn't any cross-reactivity between antibodies even if they are derived from different species. To do this, first perform staining with each antibody alone to find the right concentration. Then it is important to have a control where the secondaries are swapped between the two primary antibodies (this should be negative).

Antibodies are notorious for failing to stain properly with different conditions. Therefore, it is important to optimize a few key steps of the protocol for each antibody. Conditions to optimize are 1) fixative used (I usually use PFA because that is compatible with many different antibodies), 2) Primary antibody concentration and incubation period, 3) secondary concentration and incubation period, 4) Wash number and duration (generally more shorter washes are better than few long washes).

A shorter version of this protocol has worked well for me. I used primarily anti-Zelda antibodies.

Alternative protocol from Colleen

A shorter version of this protocol has worked well for me. I used primarily anti-Zelda antibodies.

IF Day 1. Primary antibody + wash

1. Block embryos in PBTB for 30 minutes at RT in 100ul PBTB.
2. Add primary antibody(S) to each sample (see below). Rock for 2 hours at 4˚C The following are examples: a. Added 1ul of anti-Zld : 4ul PBTB – 1:500 b. Added 1ul anti-Bcd : 4ul PBTB – 1:500 c. Added 1ul anti-lam : 4ul PBTB – 1:500
3. Wash embryos 3x with 1ml PBS.
4. Rinse embryos 4x with 1 ml PBT for 15 minutes each. Typically, I do the last wash overnight, rocking at 4˚C
   • TIP: the primary antibody used for the incubation can be recovered and saved for further use. Antibody can be used several times.

IF Day 2. Secondary antibody + wash + Counterstain

5. Block for 30 min at room temperature in 100 μl PBTB
6. Add diluted secondary antibody- wrap tube with alumina foil, rock slowly at room temperature for 1.5 hrs.
   • added 2ul of secondary stock to 8ul blocking buffer. –1:5 and then added 1ul of that. (1:500)
7. Wash embryos 3x with 1ml PBS.
8. Rinse embryos 4x with 1 ml PBT for 15 minutes each.
9. Add 1ul of Dapi to the 1ml PBT.
10. Incubate for an 5 minutes.
11. Wash 6x in PBT
12. Add 100ul of Dabco and let incubate at RT (STANDING) for at least 30 minutes out of the light.
13. Mount slides. Keep at -20˚C.
14. When ready to mount embryos, bring the embryos to room temperature. Transfer 25-30 ul of embryos to a clean slide, place two coverslips on both sides of drop, about 1 cm apart, place a third on top.
    • TIP: Alternatively, to image with the 63x objective on the confocal simply place one coverslip over about 25ul of dabco/embryos and then seal the edges with clear quick-dry nail polish. Make sure to use an eyedropper to drop nailpolish around the edges (not brush) so that there is a thick seal.
15. To save the slide, completely fill the space underneath the coverslip with glycerol and seal all the edges of the coverslips with fingernail polish. The slide can be kept at -20˚C in the dark for many months.

Example Results:

Blue- Dapi/Nuclear stain, Red - anti-Bicoid, Green- anti-GFP (AVES-GFP-1010)