I'm using ~P5 fast-dividing (time to double ~ 1 day) HEK cells grown to about 90% confluence in a T75 flask. Cells were kept in DMEM (+high glucose, +glutamine) + 10% HI-FBS + Anti-Anti until the day of transfection (1 day after plating from thawed vial). On day of transfection, I changed to Anti-free media a few hours before transfection.
Transfection: Lipo 3000 protocol with 24ug total DNA (VSV-G, Pax2, and transfer plasmid expressing GFP and GOI, 2nd gen lenti system). Media was harvested about 48hrs after transfection. (Cells were bright green under fluorescence after 1 day, and media turned yellow around the harvest time). Media was spun at low-speed and supernatant (13mL total) was filtered (0.45 uM syringe filter) and spun in UC (using SW41Ti rotor x 25,000rpm, polyallomer UC tube) for 1 hour, 45 min. A very small yellowish pellet was observed. Media was decanted into bleach and tube held upside down onto a kimwipe for 3 minutes. I added 200uL of sterile PBS to the pellet and triturated gently - the pellet disappeared quickly. The PBS was left overnight at 4 degrees to complete resuspension. The PBS turned a little red because of residual media on the side of the tube.
Titer was still only about 1 x 10^7 whereas I want 10^8 - 10^9 as most protocols seem to promise.
Any ideas? Should I switch to the Roche Fusion HD reagent? Should I use serum-free media during transfection?