C. elegans Immunohistochemistry

Sandhya Koushika and Michael Nonet, Washington University

 

Grow worms on a 10 cm E. coli seeded plate to young adult (or other stage if desired). Late adult animals with lots of eggs generally do not give good morphology.

1.        Wash worms off with 1X PBS

2.        Spin down worms in 12ml glass conical tubes. Wash 3-4 times with PBS until all E. coli are gone (clear). Make sure that you resuspend all worms thoroughly each time.

3.        Aspirate as much of the 1X PBS as possible. Precooling of worms will yield a milder fixation and often helps immunostaining of finicky antisera [precool eppendorf tube, fix and tube of worms on ice].

4.        Add a mix of 400µl Bouin’s +400µl MeOH + 10µl ßME. Alternately 400 µl of 4% paraformadehyde +400µl MeOH + 10µl ßME depending on the Ab. For a small plate of worms 200+200+5 is enough.

5.        Transfer worms to a 1.5 ml centrifuge tube with a glass pipette. For sensitive Abs.

6.        Rock tubes 15 min in rotary shaker at RT or 4°C depending on Ab. For sensitive Abs fix only 7-10 min.

7.        Quick freeze in liquid N₂ and keep at –80°C for minimum 2 hours (can be kept for months with little problem). For problem Abs I have had best results with O/N at –80°C.

8.        Quick Thaw under running hot H₂O until it melts (but before it is room temp.)

9.        Rock tubes 15 min (10 min at 4°C for problem Abs).

10. Do 4 to 5 BT washes. 1^st BT wash is a rinse and all subsequent ones are 3-5 min. washes. Typically I do spin at 1-2 K for 30 seconds.

11. Make BTB: 1x Borate Buffer, 0.5% Triton X-100, 2% BME. Avoid using if over 2 weeks old.

12. Spin, aspirate BT

13. Add BTB, rock a few seconds to mix

14. Repeat steps 11-12 once more

15. Resuspend worms in BTB and incubate for 1 hr with rotation/rocking (best is 30 min x 2 and then every hour for 2-4 hours.)

16. Wash again w/ BTB and incubate 2-3hrs. If permeablization is a problem do a harsh spin i.e. at 14k for 10-15 seconds coming down slowly to 1-2 K.

17. Quick rinse with AbA or PBST

18. Block for 30 min- 1 hr. with PBST/AbA. Remove as much of soln and add Primary Ab

19. Incubate in 4°C o/n, no shaking needed. Some Abs only work if you do 3 o/n at 4°C and on the last day a few hours at room temp. before beginning washes.

20. Wash 5x for 5-10 min in AbA. For better staining do 6-8 changes in 3-6 hours.

21. Remove most of the AbA

22. Add Secondary Ab (We recommend Alexa-conjugated secondary antibodies from Molecular Probes. Use at 1:500 for most applications)

23. Incubate 2-3 hr at RT (4°C o/n will yield better staining).

24. Wash 4x 5-10min in AbA. For better staining do 6-8 changes in 3-6 hours.

25. Mount on slides with 1 µl mounting medium.

 

Solutions

4% paraformaldehyde:

4 g paraformadehyde in 50 ml water. Keep at 60 C. Add few drops of 1M NaOH to help it dissolve. Allow solution to cool to room temp. Add 50 ml 0.2 M Na/K PO4 pH 7.2 (8.06 g KH₂PO₄, 37.75 gms Na₂HPO₄^.7H₂O in 100ml H₂O pH adjusted to 7.2). Filter and cool to 4°C before using.

 

10x PBS:

80 g NaCl

2 g KCl

14.4 g Na₂HPO₄^.7H₂O

2.4 g KH₂PO₄ pH 7.4

fill to 1 liter.

 

Bouin’s:

75 ml saturated picric acid

25 ml formalin

5 ml glacial acetic acid

 

 

50x Borate buffer:

1 M H₃BO₃

0.5 M NaOH

(pH should be ~9.5 after NaOH addition)

 

BTB:

1X Borate Buffer

0.5% Triton X-100

2% BME

 

AbA (PBST):

1X PBS

0.5% BSA

0.5% Triton X-100

10mM azide (optional)

Tween 20 can also be used in place of Triton X-100

 

Mounting medium:

0.1 g n-propyl gallate

3.5 ml glycerol

1.5 ml 100mM Tris pH 9.5

10 µl 1mg/ml DAPI (optional)

Last modified 4/25/05