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Drosophila TEM/LM Protocol

Protocol for Patrick Dolph's lab, Biology Dept.

  • NOTE: All steps preceding the 100% ethanol step should be carried out on ice.
Materials and methods.
Incubate with slow rocking

1) Dissect heads by cutting them in half and taking the larger half (or both halves) or use whole heads but cut off the probusci - Place in formaldehyde/glutaraldehyde fix in glass scintillating vials and shake overnight on ice. If shaking vials sideways, make sure to remove paper on inside of plastic lid. Try to get heads to sink into the fix so that they don’t just float on the top.

Use fresh paraforaldehyde & glutaraldehyde each time. Can store opened glutaraldehyde in a closed/capped scintillating vial at 4oC for a maximum of 3 weeks.

To make up 8% paraformaldehyde: (10 ml) ****Make up fresh each time****

0.8g of paraformaldehyde

bring up to 9 ml with dH20

pH to 9-10 with 8 ?l of 10 N NaOH

heat on a hot plate (or boiling water bath) until paraformaldehyde goes into solution

bring pH to 7 by adding 7 ?l of conc. HCl

add 1 ml of 10X PBS

cool on ice and check pH with paper (pH should be 7)

Fix: To make up 8 ml Final

2 ml 0.4M PO4 buffer, pH7.4 0.1 M PO4 Buffer

2 ml 8% Paraformaldehyde 2% Paraformaldehyde

4 ml 8% Glutaraldehyde 4% Glutaraldehyde (fresh ampule)

0.15g Sucrose 2% Sucrose

dH20 =8ml

Rotate overnight on a circular rotator at 4oC (cold room).

  • Note: Tanya’s protocol uses 2% glutaraldehyde (no paraformaldehyde) and does the glut/ OsO4 steps together.

2) Rinse 1 hour with several changes of 0.1M PO4 buffer, pH 7.4, 3% Sucrose. Change buffer every 15 minutes and shake on ice.

Rinse Buffer

10 ml 0.5M PO4 buffer

1.5g Sucrose

dH20 = 50ml

3) Fix heads overnight in 1% OsO4/2% Sucrose in 0.1M PO4 buffer, pH 7.4. Again, make sure that the heads sink in the fix.

Use fresh OsO4 each time ? do not store unused OsO4 as it is a very nasty fixative (just throw away). ****Always do OsO4 steps in the hood as it is a very volatile compound****.


-2 ml of 4% OsO4

-1.5 ml of 0.4M PO4 buffer

-3 ml of 4% sucrose in dH20

6.5ml total (aliquot to number of vials, i.e., 1 ml per vial with 4 samples)

  • NOTE: Rotate overnight on a circular rotator at 4 C (cold room).

4) Rinse samples twice in dH20 - Can store indefinitely at 4 C.

5) Dehydrate with EtOH series, i.e., 10%, 25%, 50%, 75%, 100% (all at 4 C).

-remove old solution

-add new EtOH solution, make sure walls are rinsed

-withdraw solution

-swirl solution to make sure it is mixed

-incubate on ice for 10 minutes while shaking

Note: Leaving enough EtOH in vials to keep eyes covered prevents cuticle from lifting away from the retina.

6) Do final EtOH (100%) rinse and move samples to room temp. Let the samples warm to room temp. (approx. 20 minutes).

7) Equilibrate the eyes in propylene oxide with 2 X 10 minute incubations. ****Always do propylene oxide steps inthe hood as it is a very volatile compound****. ****Use nitrile gloves (not latex) when handling propylene oxide****.

8) Add to vial
75% propylene oxide
25% Spurs

9) Rotate samples on a shaker (8 hours to overnight).

10) Place in 50% Spurs (50% propylene oxide) (8 hours to overnight).

11) Place in 75% Spurs (25% propylene oxide) for 6 hours.

12) Place in 100% Spurs for 24 hours, changing Spurs every 6 hours or so (switch Spurs as often as possible). Have samples rotating on a shaker in the fume hood with the caps off.

13) Remove heads with a pasteur pipet to molds filled with 100% Spurs.

14) Orient heads.

15) Bake samples in the plastic caps at 65 C in a vacuum oven overnight (set oven to 15 psi). Make sure to check orientation of heads after an hour or so.

For Light Microscopy:

Cut 0.5 ? sections and stain with Methylene Blue/Azure II.

A. 1% Methylene Blue in 1% Na Borate (Borax)

B. 1% Azure II in H2O

Mix fresh A + B 1:1

stain section, heat, rinse and dry.

For Transmission Electron Microscopy:

Cut 70-90nm sections and mount on 400HH Cu grids

stain sections with 2%UAmeoh for 20' and Reynold's LC for 3-5'.

Last Updated: 10/2/08