EM Fixation of Embryos

 

EM Fixation of Embryos

Adapted following advice from Andy Fire and Jim Priess

1. Mount embryos on slides as if for DIC, except agar has fixative.

Agarose Fixative

3ml 2% Agarose : 1.5% LGT Agarose + .05% HGT Agarose

5ul 1M MgCl2

400ul 0.2 M NaCacodylate (pH 7.4)

400ul sucrose (68.46g/100ml)

500ul 25% Glutaraldehyde

Make up above mixture immediately before mounting embryos. The pad should be thick, so use triple thickness tape when preparing slide. Transfer the eggs to pads with minimal fluid (to preserve osmolarity). Transfer with a pick or mouth pipette. Use an eyebrow hair to push the eggs to the center and arrange them so that they are close to each other without touching.

2. Overlay a small amount of slightly cooled agarose mix on slide and place a coverslip [#1 thickness] on top. I found that an 18mm coverslip was best for Scott Emmon's laser, but that the small 12mm coverslips worked best on Andy Fire's laser.

3. Make a map of the embryos - their arrangement on the pad, their orientation, and their relative stages. Mount a bunch of 1, 2, and 4 cell embryos on a pad, then watch them go through landmark events, like gastrulation, the migration of the dorsal hyp nuclei, or cell deaths to stage them.

4. When an embryo reaches the stage you want, permeabilize the egg shell and vitelline membrane with a laser. Shoot for edge of shell. You can see the break in the shell and the membrane. The laser needs to be at full power. On Andy's [Olympus] scope, we use a 40x oil objective. Also kill (explode) any embryos that you don't want and record their positions on your "map". Use a few embryos to pretest the fix solution for osmolarity [see below].

5. Place the slide in a humidified chamber at room temperature for 2 hours to allow fixation.

6. Remove the coverslip by pipetting 25mM NaHEPES (pH 7.5) around the base of the pad, then gently slide the coverslip off. Eggs will usally stay in place.

7. Begin dehydrations and washes. Either leave the group in their original agar pads, or cut up the pad and place indivdual eggs in separate wells of a 9-well glass plate. Either way, trim the agarose with a clean new razor blade.

8. Rinse in NaHEPES:

2X30"

1X2'

9. Wash in 0.2M Na Cacodylate pH 7.4

2X30"

1X5'

1x10'

2X1 hr

10. Post-Fix in 1% OsO4 +1%KFe(CN)6 in .2M NaCac -1 to 2 hours.

11. Wash in 0.1M NaCac

2X30"

1X5'

2X10'

12. Post-Fix in 0.2% tannic acid in 0.1M Cac - 15'

13. Wash in 0.1M NaCac

2X30"

1X5'

2X10'

Wash in 0.1M NaAcetate

2X30"

1X5'

1X10'

1X30'

14. Stain 1% Uranyl Acetate in 0.1M NaAcetate - 3 hours.

15. Wash in in NaAcetate

2X30"

2X10'

Wash in 0.1M HEPES

1X30'

2X5'

16. Before the dehydration series, remount the embryos into large chunks (~1 cc) of agar (2%) and then cut each block into a distinct shape so that every embryo can be distinguished from all the other embryos. The larger blocks help protect the embryos as you go thru all the steps and they are also much easier to see ( and therefore to avoid sucking into your pipette and discarding). The distinct shapes allow you to dehydrate all the embryos in the same vial without confusing them.

17. Rinse the blocks in dH20 and transfer to a vial. I used a scintillation vial.

18. Dehydration:

6' 30% EtOH

10' 50% EtOH

10' 70% EtOH

~undefined .. 70% EtOH [if you need to take a break, this is a safe place to halt processing overnight]

6' 90% EtOH

6' 95% EtOH

3X20' 100% EtOH

20' 1:1 Propylene Oxide:EtOH

2X8' Propylene Oxide

30' Propylene Oxide:Araldite 2:1

60' Propylene Oxide:Araldite 1:2

2X30' Araldite Room Temp

Trim the blocks and return embryos to separate wells of a 9 well plate.

24hrs Araldite RT

Transfer blocks to whatever mold you will use to hold the embryo and try your best to orient embryo for easiest trimming and cutting

3 days Araldite @60 degrees C

Buffer Recipes

0.2M NaCacodylate

A 42.8g Na(CH3)2As02 3H2O

1000ml dH2O

B 0.2MHCl [10ml conc HCl + 603ml dH2O]

50ml A + B /18.3 -> pH/6.4 Q.S. to 200ml

Pretest the osmolarity

Carolyn Norris' recent update to this method [during the June 4 workshop in Madison] stresses the importance of osmolarity trials to discover the correct amount of sucrose in the fixative to match the embryo's inherent osmolarity:

Before fixing the final specimens, she recommends doing several trial embryos with differing amounts of sucrose in the final fixative. While watching under the microscope, she shoots several laser holes in the eggshell. If the embryo collapses rapidly (implodes), then the sucrose concentration is too high. If the embryo blows up rapidly (explodes), then the sucrose is too low. Under optimal conditions, the embryo will be seen to swell in volume very slowly after a laser hole is put through the eggshell. Thus the final fixative should be tested empirically on the day of fixation, to optimize the exact osmolarity.