0.1 M PBS: Weigh 80 g NaCl, 2 g KCl, 14.4 g Na₂HPO₄⋅2H₂O, and 2.3 g NaH₂PO₄⋅H₂O, and add distilled water (Milli Q) up to 1,000 ml. Dilute 1:10 with distilled water prior to use.

4% Paraformaldehyde: Dissolve 4 g of granular parafolmaldehyde in 100 ml of PBS. This solution must be prepared under the chemical fume hood. Pre-heat the PBS to 60°C to allow the parafolmaldehyde to completely dissolve, then add 50 μl of 1N NaOH while stirring.

4% Paraformaldehyde/0.4% glutaraldehyde: Mix 70% glutaraldehyde and 4% paraformaldehyde in a 1:175 v:v ratio.

0.15% Glycine: To 30 mg of glycine, add PBS up to a 20 ml final volume.

2.3 M sucrose: Dissolve 78.73 g of sucrose in 50 ml of PBS and stir until the sucrose is completely dissolved. Add PBS to a 100 ml final volume and store 50 ml aliquots at 4°C.

2% Methylcellulose: Add 4 g of methylcellulose to 196 ml of distilled water while stirring at 90°C. Rapidly cool to 10°C on ice while stirring, and then add water to a final volume of 200 ml. Seal with parafilm and stir overnight at 4°C. Store methylcellulose at 4°C in the dark, for up to 3 months.

12% Gelatin: Dissolve 12 g of gelatin powder (local food brand is satisfactory) in 75 ml of PBS by first stirring the solution for 10 min at room temperature and then for 4–6 h at 60°C. When the gelatin is completely dissolved, cool the solution to 37°C, add 200 μl of a 10% sodium azide solution, and finally add PBS to a 100 ml final volume. Aliquot and store at 4°C, up to several months.

10% BSA stock solution: Add 10 g of serum albumin in 100 ml of distilled water (Milli Q) and gently stir overnight, to prevent foaming, at 4°C. Set the pH to 7.4 with 1N NaOH, and then add sodium azide to a 0.02% final concentration. Centrifuge the solution for 1 h at 100,000  ×  g, retrieve the supernatant, and store in small aliquots at 4°C.

4% Uranyl acetate: Dissolve 4 g of uranyl acetate in 100 ml of distilled water by stirring at room temperature.

0.3 M oxalic acid: Dissolve 3.78 g of oxalic acid in 100 ml of distilled water by stirring at room temperature.

2% Uranyl acetate/0.15 M oxalic acid: Mix 4% uranyl acetate and 0.3 M oxalic acid in a 1:1 v:v ratio. Adjust the pH to 7–8 by adding drops of 25% ammonium hydroxide while stirring.

0.4% Uranyl acetate/1.8% 25 ctp methylcellulose: Gently mix 1 ml of 4% uranyl acetate to 9 ml of 2% methylcellulose solution. Store the solution at 4°C in the dark, for up to 3 months.

50% Glycerol in water: Add 100 ml of glycerol to 100 ml of deionized water while stirring. Aliquot to 100 ml bottles and autoclave.

Formvar: Weigh 1.1 g of formvar and place in a 100 ml volumetric flask. Add chloroform to a 75 ml volume, and stir until dissolved. Subsequently, add more chloroform to a 100 ml final volume and let rest overnight at room temperature (see Note 6). Store formvar solution in a clean bottle for up to 3 months.

Fix cells by adding a mixture of 4% paraformaldehyde/0.4% glutaraldehyde to an equal volume of culture medium for 2 h at room temperature (see Note 7).

Quench free aldehyde groups by incubating the cells for 10 min in 0.15% glycine at room temperature.

Add a small volume of 1% gelatin in PBS. Scrape off the cells from the tissue culture dishes with a cell scraper.

Transfer the cells to a 1.5 ml microcentrifuge tube, pellet the cells for 1 min at 13,000 rpm (16,100´g) and remove the supernatant.

Resuspend the cells with 1 ml of 12% gelatin for 10 min at 37°C.

Pellet the cells for 1 min at 13,000 rpm and carefully remove the supernatant (gelatin 12%).

Resuspend the pellet in 1 ml of 12% gelatin.

Repeat the centrifugation step and remove the supernatant.

Transfer the pellet to 4°C for 15 min.

Carefully cut the microcentrifuge tube, at the level of the pellet, using a razor blade and remove the solidified gelatin/cell pellet from the microcentrifuge tube.

Using a sharp razor blade cut small (∼ 1 mm³) gelatin blocks with the aid of a stereo-microscope (see Note 8) while taking good care not to dry the sample during the cutting procedure. Rapidly proceed to the next step.

Submerge the small blocks in 2.3 M sucrose solution, and slowly rotate tubes overnight at 4°C (see Note 9).

Carefully retrieve the specimen blocks from the 2.3 M sucrose solution, using stainless biology tweezers # 5, and transfer to the top of the aluminum pins.

Remove the excess sucrose by gently touching with a piece of filter paper while leaving enough sucrose to glue the block to the pin.

Position the specimen in the best orientation for sectioning (see Note 10).

Rapidly freeze pins by dipping in liquid nitrogen and store in liquid nitrogen (see Note 11).

The formvar film is made on a microscope slide. The surface is cleaned by wiping the slide with a chamois and next with lens paper. The Formvar solution is poured gently without forming air bubbles in an adapted separating funnel provided with two stop-cock valves in its outlet. The upper one is used to regulate the flow, the second one to switch on or off. The level of the Formvar solution in the funnel is about 2/3 of the length of the slide. The slide is carefully placed in the funnel with a pair of tweezers and the top of the funnel is covered to reduce evaporation of the chloroform. Then the lower valve is opened with one turn. The upper valve is set in such a way that the Formvar/chloroform solution flows out of the funnel in 10s. Remove the slide from the funnel after 30 s and loosen the film at the edges of the slide by cutting the film 0.5 mm from the edges with a razor blade. Fill a beaker with clean distilled water and slowly bring the slide in vertical position into the water. The film will float on the water surface. The thickness of the film should be about 50 nm, which will give it a silver-white appearance on the water surface. The grids can be placed on the film, avoiding irregularities in the film. Once the film is covered with grids, a microscope slide provided with a sticker is gently pressed on the film into the water. The film with grids will attach to the sticker. Be sure to submerge the slide far enough into the water so that the film is totally submerged before the slide can be removed from the water and dried.

Cut semithin (200–300 nm) or ultrathin cryosections (60–65 nm) with a diamond knife using the cryo-ultramicrotome. The temperature of sample, knife and chamber are set at −100°C for semithin sections, and −120°C for ultrathin cryosections, respectively. The cutting speed is set at 1–2.5 mm/s.

Collect ribbons of flat cryosections on a droplet of a 1:1 mixture of 2.5 M sucrose and 2% methylcellulose and transfer to formvar film-coated, carbon stabilized, gold finder EM grids.

Store the cryosections at 4°C, for up to 1 month.