2.5% glutaraldehyde in YFB (Your Favorite Buffer) preserves specimens exceptionally well; however, it is more viscous than formaldehyde. If tissue is thick, penetration of the fixative will be a problem. You can overcome this in part by using...

1/2 strength Karnovsky’s fixative:

2% prepared formaldehyde*

2.5% glutaraldehyde

in YFB

(some add 5 mM CaCl₂)
 
 

*To make an 8% stock, add 8 g of the HIGHEST quality paraformaldehyde you can find (EM Sciences, Polysciences, maybe Sigma) to about 90 ml of water. The paraformaldehyde will not dissolve. Place on a stirring hot plate in a fume hood, stir slowly and heat to 70-80ºC (steam should be visible, but not boiling). Carefully add 1 N NaOH, drop by drop (usually 2-3 drops suffice) until the solution clears. Give each drop a chance to work. Cool. Dilute to 100 ml, and filter through Whatman paper). Store at -20ºC. Defrost in 37ºC water bath--let it warm all the way before agitating the tube to get the formaldehyde back into solution. Note that if you are having trouble realizing adequate fixation, you might want to try making the formaldehyde fresh for every preservation.

Buffer choices:

A phosphate buffer. A good choice, but beware that phosphate crystals can precipitate during sample preparation, which will then be usable, but not adequate for publication at high magnifications. This precipitation is exacerbated by the presence of Ca2+. However, many great images have been published using phosphate.

If you want to eliminate that variable, then sodium cacodylate buffer is a great choice. Beware: it is toxic. Use at 0.1-0.2 M sodium cacodylate (MW 214); pH to 7.4 with HCl.

2. Fix for a minimum of two hours (pistils) at room temperature; all day is optimal. If you want to keep the samples for awhile before further processing, store at 4ºC.

3. Wash 3X with YFB. (NB: If you have "complex" tissues, each wash should last anywhere from 5-15 minutes).

4. Osmicate whenever lipid preservation is desired. IN A HOOD, postfix with 1-2% OsO₄ diluted in YFB. Ampules of stock osmium solutions are most convenient; store at -20ºC when not in use. Minimize the glassware, plastic, etc., that will come into contact with osmium. Since it is not so critical to get the concentration exact, you can dilute the solution directly on the specimen, using a Pasteur pipet. Put YFB in first, however. You can store left-over osmium wrapped with Parafilm. However, when ready to re-use it, check to make sure that the osmium has not sublimed by assuring that the solution is still a dull yellow. Postfix for 2 hr at RT. NO LONGER.

5. Wash as in step 3.

6. Dehydration: Using ethanol stored at -20ºC, dehydrate the specimens, 2X in each ethanol concentration (this can be done on the bench; just keep the alcohols stored cold) for 5-15 minutes, depending on the specimen thickness. An example of a successful series is: 50%, 70%, 80%, 90%, 95%, 100%. After the 100%, do two more washes with room RT 100% (if you can, open a fresh bottle for this step). Finally, do another wash with the RT 100%, but let the specimen incubate 20-30 minutes.

7. Wash 2X for 10 minutes each with amyl acetate. Assure that you are doing this in an amyl-acetate-resistant container (such as polypropylene).

8. Critical point drying:

Transfer specimens to holders.

Place holders into chamber of drying apparatus

Place chamber in ice water

Open liquid carbon dioxide; let gas flow through apparatus.

After 10-15 minutes, when the smell of amyl acetate has dissipated (it smells like bananas), close the gas intake to the chamber; shut off the tank of carbon dioxide.

Exchange ice water to hot (just boiled works well) water.

Let pressure go up; after it has reached a maximum, slowly release pressure.

Carefully remove holders and specimens. Keep them dry!

9. Sputter coating--8 nm gold.