Exercises:
Stain mitochondria in living cultured cells
Materials:
1. HeLa cells or other similar cultured cells, grown on cover slips.
2. DiOC6(3) 0.5 mg/ml in ethanol, keep protected from light at room temperature.
3. Silicon rubber for making observation chambers
4. Forceps, microscope slides, kimwipes

Procedure
Make an 0.1 ug/ml solution of DiOC6(3) in growth media (1:5000 dilution of stock dye). This can be used for a few hrs, but not overnight.
Make silicon rubber chamber: cut out a small square so that the cover slip will fit over it.
Press the silicon rubber chamber down on a microscope slide.
Fill the chamber with dye solution, place the cover slip on top and press down carefully to make a good seal.
Wipe excess fluid with kimwipe so that the outside of the cover slip and chamber is dry.
Observe in fluorescence scope using fluorescein filters.
If fluorescence staining of mitochondria is dim, try 0.2 ug/ml or higher.

Stain ER in living cultured cells
Procedure
Make an 0.5 ug/ml solution of DiOC6(3) in growth media (1:1000 dilution of stock dye). This can be used for a few hrs, but not overnight
Make and set up silicon rubber chamber as above.
Mount the cover slip as above, observe in fluorescence scope using a 63x or 100 x objective lens.
It may take 5-10 minutes for ER staining to develop. If only mitochondria are stained, try 1.0 ug/ml DiOC6(3) staining solution.
The mitochondria are usually swollen at dye concentrations that stain the ER. This indicates that the dye concentration is toxic to them.

Stain ER in living onion cells.
This is a very good preparation for doing time lapse of ER movmenets or real time movies of mitochondrial movements.
Additional materials to above:
1. An onion, preferably fresh with no blemishes
2. A razor blade
3. Water, either tap or spring, but not distilled or deionized)

Procedure
Make an 0.5 ug/ml solution of DiOC6(3) in water.
Cut out a small square (0.5") from an onion using a razor blade. Use forceps to pull off the thin epithelium from the inner-facing side of the square. Place the epithelium flat on a microscope slide. Put about 20 ul of dye solution on top of it, then put a 22 x 22 mm cover slip on top of that.
Observe fluorescence using a 63x or 100x objective lens.

Stain ER in fixed cells
Additional materials:
1. Sucrose cacodylate buffer: 100 mM sucrose, 100 mM sodium cacodylate, pH 7.4
2. Glutaraldehyde; 0.25% in the sucrose cacodylate buffer. Make it in a capped culture flask or scintillation vial. For these experiments, it can be kept in the refrigerator for a week or two, though you should not keep it for so long for electron microscopy. Note - glutaraldehyde is a fixative: Do not drink or mouth pipet! Wash it off your hands! It is not good to breathe in the fumes for long periods of time.
3. Parafilm, lid top of a 100 mm petri dish.
4. Scintillation vial for glutaraldehyde waste.
Procedure
Make a staining solution of 2.5 ug/ml DiOC6(3) in sucrose cacodylate buffer.
Cut a square of parafilm that will fit over the lid top of a 100 mm petri dish. With the paper still on, put the parafilm side against the inside of the lid top, then press it so that the parafilm is snug against the lid top. Remove the paper.
Put a cover slip cell side up on the parafilm (remove the cover slip from the culture dish vertically so that there is little medium on it). Fix the cells by putting on a few drops of 0.25% glutaraldehyde solution for 3-5 min.
Remove the fixative from the cover slip. Put on a few drops of DiOC6(3) staining solution
After 10 sec of staining, wash the cover slip once or twice with sucrose cacodylate solution, then mount the cover slip in a silicon rubber chamber in sucrose cacodylate.
Observe fluorescence using 63x or 100x objective lens.
Note: fluorescence is best in the first 10 min after staining. The dye starts to accumulate in lysosomes afterwards. With longer periods of time, autofluorescence from glutaraldehyde begins to build up. Staining of fixed cells has the advantage that mitochondrial morphology is preserved.

Further exercises for the interested:
Effects of fixation, permeabilization, tonicity
Devise a live cell chamber which allows perfusion
Stain living cells for ER with DiOC6(3)
Perfuse through:
1) glutaraldehyde (does glutaraldehyde fixation preserve ER morphology?)
2) formaldehyde (3.7% in sucrose cacodylate; dilute from 37% formalin) (does formaldehyde fixation preserve ER morphology?)
3) formaldehyde then triton x-100 (0.1%) (what does detergent do to the ER?; how good an image then, would you expect from immunofluorescence labeling of ER?)
4) glutaraldehyde then 1:1 hypotonic sucrose cacodylate; 1:3 hypotonic sucrose cacodylate (is tonicity important for preserving structure even in fixed cells?)
5) triton x-100 (what does detergent do to the ER?)
6) saponin (permeabilized cell models of Balch et al.) (is saponin more gentle on ER structure than triton X-100?)

Staining of cell free preparations
Make staining solution of 0.5 µg/ml.
Add to cell free preparations.
Observe.