Glutaraldehyde/KMnO4 protocol

(Many of the suggestions in protocol described below relate to the problem that KMnO4 tends to precipitate in tissues that have been previously fixed with aldehydes.  This seems to be caused not only by residual free aldehydes in the tissue, but also by residual Hepes buffer in the tissue.)

(Note that cells or tissues are never cooled below room temperature at any time during this workup.)

  1. Primary fixation of culture in 2% glutaraldehyde in "NaHCa" buffer*, for no more than 1 hour at RT*.
  2. Wash in "NaHCa" buffer 2x, 5min each
  3. Wash for 15 min in "quenching solution" (like one would use for immunocytchemistry) (This is to eliminate free aldehyde groups and general stickiness.) (It consists of 50mM glycine, 50mM lysine, and 50mM NH4Cl,all dissolved in the usual "NaHCa" buffer.)
  4. Wash again in "NaHCa" buffer, 5x, 5min each

*Aldehyde fixaton should always be done at room temperature (RT), never in the cold or in the ‘frig, since it’s not fixing the lipids in biological membranes, and chilling lipids promotes artifactual phase-separations.

*"NaHCa" buffer consists of 30 mM HEPES buffer, brought to pH 7.4 with NaOH (not KOH!), with 100 mM NaCl present to make it roughly isotonic and 2 mM CaCl2 present to preserve cell membranes.
(P.S. For virus experiments, 10mM MgCl2 is also added to this fixative, to improve the preservation of delicate & stressed cytoplasmic proteins & cytoplasmic polynucleotides.)
(In making up the buffer solution, to which the glutaraldehyde will be added later, be sure not to add the CaCl2 until after pH'ing the buffer to 7.4, or else the calcium will precipitate when the NaOH hits the surface of the solution.) (We usually add the CaCl2 from an unbuffered 1M stock in distilled water.)
(When adding the fixative itself to this buffer, use an "EM grade" of glutaraldehyde, usually supplied by commercial companies as a concentrated solution (25-70%) in a sealed 2ml or 10ml vial over argon or another inert gas.)

  1. Wash in 0.1M NaCl, 2x,10min each (This is to eliminate all traces of Hepes buffer from the tissue, otherwise left over from the "NaHCa" buffer)
  2. Postfix in 0.5% KMnO4 freshly dissolved in 0.1M NaCl (with a bit of bath-sonication to get it dissolved) (for exactly 15min, again at RT)
  3. Wash two more times in 0.1M NaCl, 5min each, to rinse away any residual KMnO4
  4. Dehydrate through 25%, 50%, 75%, 95%, and 100% ethanol (20 min total for this)**
  5. Embed in epoxy resin according to the following patented “protocol for fast embedding”, which involves diping the coverslips sequentially into nine small beakers containing the following:
    1. 100% Ethanol (really dry!)
    2. 100% Ethanol, again
    3. 100% Ethanol, again
    4. 100% propylene oxide
    5. 100% propylene oxide, again
    6. 50/50 (V/V) propylene oxide and epoxy resin*
    7. 50/50 (V/V) propylene oxide and epoxy resin, again
    8. 100% epoxy resin
    9. 100% epoxy resin, again (The coverslips should of course be held with forceps
      and should be ‘swished’ around in each beaker for just about 30 seconds)
    (*The full mixture of epoxy resin, including the accelerator, is made up fresh right before use, and is outgassed by repeated vacuum-application many times, until all the bubbles come out of it.  This we do with the vacuum-oven that will be used for the final epoxy polymerization, but with the heater turned off, so the epoxy doesn’t get heated and start to polymerize.  This the most tedious step of the whole procedure..You’ll see!))

The coverslips are then set in aluminum dishes filled with a ~1mm deep layer of epoxy resin and placed in the vacuum oven (now warmed up to 65degC), and evacuated two or three times, to draw off any residual propylene oxide.

Finally, the coverslips are inverted over  “Beem” capsules (or large gelatin pills), held perfectly upright somehow, and filled to the very top with the same fresh epoxy resin (trap no bubbles!), and placed in the heated 65degC vacuum-oven for polymerization. (Don’t apply any vaccum at this point, though, or bubbles might get trapped under the coverslip.)

After polymerization, the coverslips are “popped” off the Beem capsules by the usual technique: touching the glass to a copper or brass block partially immersed in LN2, and then pulling it off the epoxy.

(Actually, we abhor this technique for separating glass coverslips from polymerized epoxy, and have switched completely to growing all our monolayer cultures on Thermanox® coverslips (15mm diameter circles).  These have adhesive properties almost identical to Falcon tissue culture dishes (much better than glass), and much more importantly, they peel off the “Beem” capsules without the slightest problem, and without all the LN2 fuss....so long as one removes the sample from the 65degC oven and peels the plastic coverslip off the capsule immediately, e.g., while it is still hot!!

If you don’t know about these Thermanox coverslips, you can read up on them at the following links:

http://www.nunc.de/page.aspx?ID=226

http://www.emsdiasum.com/microscopy/technical/datasheet/72274.aspx
 
The only problem with Thermanox coverslips is that even though they’re clear and transparent, they have very poor optical properties, so you cannot image cells on them by looking through them,  but must use a water-immersion objective in the LM, so as to image cells on them from above.