Protocol

Prepare a 6-well plate by adding 2 ml of Raw Cell media per well. If coverslips are required, add one coverslip/well prior to adding the media.
Wash the cells pre-cultured in a T25 flask with 5 ml of PBS and trypsinize with 1.5 ml of trypsin for five minutes. After adding 3.5 ml of fresh media, determine the cell concentration using a hemocytometer.
Add the cells to the six well plate at the following concentration:
- 500,000 cells/well for subsequent microscopy analysis of transfected cells grown on the coverslips => ~35-40 cells/field-of-view at 63x, with ~20% transfected
- 1 million cells/well for subsequent parachuting of cells onto coverslips
Rock the plate sideways to evenly distribute the cells.
Culture the cells overnight at 37^oC.

Thaw the DNA to be used for transfection.
Set up a reaction for each well in a microcentifuge tube:
- Add 3.3 µg of DNA to 150 µl of serum-free media. Vortex the tube for 1-2 seconds to mix.
- Add the Fugene-HD Transfection Reagent at a ratio of 3:1 of Fugene (µl) :DNA (µg). Flick the tube 15-20 times to mix. Note: The Tranfection Reagent is ethanol-based and evaporates easily. Keep the Transfection Reagent in the fridge until its addition and do not leave uncapped. Re-wrap in parafilm after use.
Let the reaction sit for 15 min at Rm temp in the tissue culture hood, for the DNA-containing micelles to form.
Add the total reaction volume of each microcentrifuge tube to its respective cell well, drop-by-drop. Shake the plate sideways to mix.
Incubate the transfecting cells overnight at 37^oC.

Aspirate the media. Wash the cells twice with non-sterile PBS, at 2mL/well. Note: Be careful not to knock the cells off the coverslips during the wash steps.
Set up a batch reaction. For six wells, combine:
- 1.2 ml 10X PBS
- 3 ml 18% PFA
- 10.8 ml ddH₂O
Add 2mL of reaction/well. Let the plate sit at Rm temp for 15 min. Note: Keep in the dark if transfected with a fluorophore.
Aspirate. Wash once-twice with PBS and cover with 2 ml PBS/well.

Raw Cell Transfection