Resin Embedding

Basic Processing Protocol for Tissue Culture Cells Grown in Plastic Dishes.

This method can easily be applied most biological samples without problem.

2). Carefully scrape the cells from the surface of the plastic, preferably using a wedge-shaped piece of teflon held in artery forceps, transfer to Eppendorf tubes and spin them down into a pellet. The presence of the protein in the buffer will stop the cells from sticking to the sides of the centrifuge tube.
Cells may be scraped as soon as the aldehyde is added and the cells left to fix as a pellet. This may cause the cell pellet to stay together during processing.

3). If the pellet of cells will hold together, remove them from the Eppendorf tube and transfer them to glass vials for further processing.
If the cell pellet is too small or fragile for this, leave in the Eppendorf tube. Large pellets cannot be easily processed in the bottom of these tubes. If the pellets are too large but also fragile, the cells can be split into smaller pellets and processed more easily in their tubes.

4). Wash pellets with 200 mM sodium cacodylate buffer (3 x 5').
At this stage, the samples will be fixed so can be washed in any sutable buffer, and even water. Sodium cacodylate is suggested because it is often the buffer used during the first aldehyde fixation.

5) Post fix in 2% osmium tetroxide in 100 mM cacodylate buffer (1 hr on ice).
Take care with the osmium tetroxide, even the fumes are dangerous.

6). Wash in water (3 x 5').

7). Wash in 50 mM sodium maleate pH 5.2 (3 x 5').

8). en bloc stain in 1% uranyl acetate in maleate buffer pH 5.2 (1 hr RT).
Using uranyl acetate at low pH will improve membrane contrast. Analternative contrasting method, which adds even more membrane contrast, is to soak the samples iovernight at 4°C in a saturated solution of uranyl acetate in70% methanol . If using this method omit the next water wash and continue with the dehydration steps.

9). Wash in water (3 x 5').

10). Dehydrate: 70% ethanol (3 x 5'). 95% ethanol (3 x 5'). 100% ethanol (3 x 10').

11). Replace the ethanol with propylene oxide (2 x 5'). Cells that have not been scraped from the culture dishes and pelleted can be removed from the plastic surface of the dish at this stage.
The propylene oxide will dissolve the plastic and the cell layer will float off. Remove the cells quickly because the propylene oxide continues to dissolve the plastic. Transfer the cell layer, in propylene oxide, to Eppendorf tubes, pellet them down and continue. These pellets do not hold together very well so tke care that they do not fall apart in the resin.

12). Leave in fresh propylene oxide for 20' then in propylene oxide/Spurr's resin (1:1 mix) overnight. Mixing during the incubation will help infiltrate the Epon into the pellet.
A slow turning wheel will achieve the correct amount of mixing.

13). Remove the cap from the tubes and leave open for 2 hr. Remove Spurr's resin and replace with fresh resin, return the sealed container to the mixing wheel and leave for 4 hr in resin.

14). Transfer the cell pellets to fresh Spurr's resin in molds, label the blocks with paper labels and leave at 60°C overnight. After one night in the oven, the pellets should be hard enough to trim and section.

If you are impatient for your results and find that this method is too long. Embedding using microwave ovens is a useful alternative.

1). Wash cells with PBS twice. Remove the PBS and replace it with 3% gluteraldehyde in 100 mM sodium cacodylate (pH 7.4). After 1 hr, remove the fixative and replace with buffer containing either 1% BSA or 1% FCS.
The cells can be washed with 100 mM sodium cacodylate buffer instead of PBS. This may alter the morphology of the mitochondria.