Cell Culture and Transfection
Day 1: Plating Cells
1. Add an acid-washed, sterile glass cover slip to each well of 6-well dish.
2. Dilute poly-lysine solution (Sigma P8920, 0.1%) 1:10 in sterile water. Add 0.5 ml of diluted solution on to each cover slip. Incubate at RT for 3-5 minutes; remove lysine solution, wash cover slips with 2 ml of PBS.
3. Plate HeLa cells at a seeding density of ~2.5 × 105 cells/ml in 2 ml of growth medium in six-well dishes containing poly-lysine–treated glass cover slips. Want cells to be at 50 to 80% confluency in ~24–48 hours for transfection.
Day 2: Transfection
1. Mix 150 ul DMEM (without FBS) and 6 ul TRANSIT reagent in a microfuge tube. Vortex 5 seconds to mix, and let sit at RT for 5 minutes.
2. Add 2 ug of plasmid DNA to the mixture, pipetting up/down to mix, and let sit for 10 minutes.
3. Add the DNA mixture to the cells, noting what DNA went into which well. Gently swirl a couple of times.
4. Add 20 ul of 0.1X MONSTER Reagent (1:10 dilution in sterile water) to each well.
5. Return cells to growth chamber.
6. Fix cells ~20 - 30 hours later.
Day 3: Fixing cells and mounting the cover slips
1. Remove the growth medium and add 2 ml PBS (37°C).
2. Remove the PBS and add 2 ml of 4% paraformaldehyde/PBS; incubate 20 minutes at 4 C (place dish in refrigerator)
3. Remove fixative and gently wash cells 2 times with 2 ml PBS
4. Wash cells twice with 2 ml of deionized water.
5. Add a drop of Gel Mount to a microscope slide and carefully transfer the cover glass to the slide with the cells down in the Gel Mount solution. Make sure cell side of cover is down. Let slides dry overnight in a dark drawer.
6. Let slides dry for >16 hours before imaging (mounted slides may be stored at room temperature, protected from light, for months)
4% paraformaldehyde fixative
Combine sodium phosphate salts in 50ml of water until dissolved, add 10 ml formalin with stirring; add NaOH or acidic acid if necessary to get pH 7.0. Make up to 100 ml with remaining water. Fixative is usually stable for 30 days if stored at 4 degrees C. Make fresh if any doubt. Old fixative will mess up your results. Note: Formaldehyde is a carcinogen and is toxic by inhalation. It may cause very serious irreversible effects through inhalation and skin contact.
Acid washing coverslips
Most coverslip manufacturers apply a coating of silicone to glass slides to prevent them from sticking together, this silicone can prevent some cell types from adhering to the glass. Acid washing removes the silicone and micro-etches the surface of the glass to give adhesion similar to plastic without affecting fluorescent light.
1. Make-up 100 ml of 2 parts Nitric Acid (66ml)- 1 part HCl (33ml) in a glass beaker in the hood. Solution will turn orange-red. Wear acid gloves and avoid acid vapors -this is an extremely dangerous solution!
2. Place #1 coverslips into solution a few at a time. Let sit for about 2 hrs. with occasional swirl.
3. Decant acid into waste bottle carefully or flush with water down sink.
4. Wash extensively in deionized water until pH of water is back up to 5.5 -6.0.
5. Store in 70% EtOH in closed jar.