Prepping of samples on 'Nevada' glass
For electron microscopy, cells are grown on 3x3mm pieces of glass generated by scoring and breaking standard #1 glass coverslips. (The tiny size of these coverslips is to facilitate their later quick-freezing.)

(These coverslips are generally cut into the shape of the US State of Nevada, to permit ready determination of which side of the glass the cells are attached to.)

These tiny coverslips you will receive from us, in a 20ml glass scintillation-bottle.  We will have cleaned them with chromic acid after cutting them down to size, and we will have rinsed & sonicated them umpty-ump times in Milli-Q water to get rid of any traces of chromate.

  • Upon receipt, you have to do all the following steps in a sterile hood, with a 1L bottle of sterile distilled water (autoclaved) ready to use:
  • To retrieve the coverslips from the scintilation-bottle, swirl it a bit and abruptly pour it out into a sterile p60 culture dish.  (Have more distilled water ready to add to the bottle and repeat, if any pieces of glass remain stuck inside it.)
  • Now to sterilize the coverslips, fill four more p60 culture dishes with sterile water and pass each coverslip individually through each dish (with only a single "stablike" immersion in each dish, plus a couple of seconds of swirling-motion under the surface).
  • To hold onto the coverslips while doing this, grab 'em on the very corner with sterile forceps; curved Dumont #7's are best for this.
  • The goal here is to NOT transfer whatever bit of oil that might inadvertently have gotten on the surface of the water in these dishes onto your coverslips, or your cells won't stick to 'em properly.
  • End up by placing each coverslip in a sterile p35 culture dish that is empty (no water).
  • Before the coverslips dry out, add your cells, already in full medium and at the concentration you would usually plate them at, right into the p35.

(Repeat: don't let the water-rinsed coverslips dry out before plating cells onto them, or the cells won’t attach properly.)

At the appropriate point of growth & differentiation, the coverslips should be removed from culture medium, rinsed 3x10 sec in three separate p60 culture dishes containing Ringer’s solution (1, below) and warmed to 37 deg in a open waterbath. 

Immediately thereafter, they should be fixed in 2% glutaraldehyde freshly dissolved in a proper vehicle (2, below) from a concentrated stock of EM-grade glutaraldehyde. (Fixation is at room temperature, and thereafter, the fixed cells are never cooled in a ‘frig...it promotes membrane phase-separations.)   

Each tiny coverslip should then be placed individually into a 1.5ml Eppendorf tube, (e.g., one coverslip per tube, so the cells can’t get rubbed off), and all  the tubes sent to us by FedEx, in a Sigma-type styrofoam box filled with something that will insulate them from getting cold in the belly of an airplane.

As these tubes are prepared, it’s very important to make sure that they are filled completely with fixative and no air-bubbles at all are left inside, or the air bubble will come to cover the cell-associated surface of the coverslip during the tumbling of FedEx baggage, and all will be lost.

(1) Mammalian "Ringer's" solution:

155 mM NaCl, 3 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 3 mM NaH2PO4, and 5 mM HEPES brought to pH 7.4 with NaOH, plus 10mM glucose (for maintaining vertebrate cells in a healthy, living condition during light microscopy or during any physiological or pharmacological manipulation(s) in preparation for EM:)

In making this up this solution, be sure not to add the CaCl2 until after pH'ing the whole mixture of other salts 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.)

For long term light microscopic recording of living cells we commonly add 1% BSA to this solution, so long as it will not adversly affect any drug-tratments we are trying (e.g., complex the drugs or something), or we even add 10% fetal calf serum (minus neutral red pH-indictor), if we want to keep physiological functions "revved up" to the max.

(2) For fixing whole, living cells, we use 2% glutaraldehyde in what we call "NaHCa" buffer.

This 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.
(Note:  These days, we typically add 150mM sucrose to this fixative-buffer, as we are having trouble with cells “blowing out”, even though the fixative is supposedly isotonic.)

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.

WHAT WE DO, ONCE THE COVERSLIPS REACH US:

Upon receipt, the coverslips are picked out of the fixative with fine forceps and washed by brief immersion in several different dishes of distilled water, using extreme care to insure that during this step, no oil films are generated on the dishes of water and transferred onto the coverslips.  (The reason for this 30 to 90 sec water-wash - the most problematic step in the whole procedure - is that ANY residual salts or organic molecules left on the coverslips at the time of freezing will appear on their surfaces as an unattractive “scum” after freeze drying.)

Next, without allowing any time for air- drying, each water-washed coverslip is mounted on a 3x3mm slab of aldehyde-fixed and water-washed rabbit lung (0.8mm thick) that serves as a “cushion” for the next step. Then it is quick-frozen by abrupt impact against an ultrapure copper block cooled to 4 degrees above absolute zero by a spray of liquid helium.  Thereafter, the coverslip is stored in liquid nitrogen until mounting in a Balzers’ Model 301 vacuum-evaporator.  In this device, it is next freeze-dried by warming it to minus 80 degrees Celsius for 15 min, and then is rotary-replicated with a thin (~2nm) film of Pt evaporated over 5-10 sec from an electron beam gun mounted 15-20 degrees above the horizontal, all while the coverslip is rotated at 5Hz.  This Pt ‘replica’ of the coverslip and the cells remaining attached to it is then immediately supported or “backed” by evaporating ~10nm of pure carbon onto it, using a standard carbon-arc supply mounted some 10 degrees off the vertical.  This takes approx. 4 sec. and is done while the coverslip continues to rotate, to insure the generation of a strong uniform film of carbon.

Next, the coverslip is removed from the Balzers device, allowed to thaw, and the replica is floated off by immersing it at a ~45 degree angle into full strength (47%) hydrofluoric acid (HF).  Immediately thereafter, the replica is picked up off the surface of the HF with a glass rod and transferred via the same rod through several washes of distilled water, then a brief cleaning by floating on standard household bleach, and then more distilled water washes, before being picked up on a 75 mesh formvar-coated EM grid.  For electron microscopy, the grid is mounted in a eucentric side-entry goniometer stage of a JEOL 200CX electron microscope, imaged at 30-70K magnification, and photographed in stereo at +/- 10 degrees of tilt off the vertical axis.

For the production of final “anaglyph” stereo images, the two stereo micrographs representing each field are placed in proper register on a Bessler copy stand and photographed at an additional 3-6x magnification with a Kodak 520 digital camera, producing a 2000x1200 pixel, ~`1.5MB B&W  JPEG file for each view.  The files are next sorted into left and right views by direct inspection on the computer screen, and then using Adobe Photoshop, the right view is converted to a pure red-channel RGB image and the left view to a blue+green-channel RGB image.  Next, either one of these colored images is copied directly onto the other, and the two are imaged simultaneously by selecting the  “screen” command in the “Layers” menu of Photoshop.  This creates an anaglyph stereo image of the original field, in a roughly 3-4MB  JPEG file.  The anaglyph is finally brought into perfect alignment by using the “free translate” command in Photoshop on one of the two layers.  (This operation in Photoshop even allows for correction of slight mismatches in magnification between the two original electron micrographs.)

The final digital anaglyph stereo image is then transferred to a standard dye-sublimation printer for publication.