Thanks to L. Wirtz and G. Cross for these protocols. Click here to go to their web site.
The important thing to bear in mind is that procyclic forms show density dependent growth. Cells at low density actually double more slowly than cells at high density, and harsh dilution can kill a culture. This is especially important to remember when scaling up nascent clones (see below) or when adapting established cell lines to new serum/culture conditions. To adapt procyclics to new culture conditions, maintain them at densities between 5 and 10 x 106/ml. Cultures will enjoy an initial honeymoon phase of normal growth (generation time about 12 hrs) but within a week or 10 days, will almost inevitably undergo a crash. If the crash occurs at sufficiently high density, an adapted culture will emerge which grows well in your serum. Be sure to freeze cells before the crisis. No need to do any transfections until you have an adapted line.
For stable transfections, we use 1-3 x 107 cells (in 0.5ml) and 5-10 µg of DNA per cuvette. Cells are harvested from a log phase culture (4-8 x 106/ml) with a gentle 4o spin (<2000rpm in a tabletop), washed once in ice-cold ZPFM, and resuspended at 1-3 x 107/0.5ml. Cell suspension and cuvettes are kept on ice. Settings for electroporation on the BTX apparatus are peak discharge of 1.6 kV, 2.5 kV/resistance, and resistance timing set on R2. A single pulse is used; the time constant should be about 0.3. Cells are zapped then transferred to 10 ml of SDM medium with appropriate drug(s) according to host cell background, and incubated overnight. Selection is applied the following day (2.5 µg/ml of Phleo for Lew79 derivatives, G418 at15 µg/ml for Lew13 derivatives, and Hygro at 50µg/ml for Lew90 derivatives). At this time we go ahead and dilute the cells serially in 24-well microtitre plates, using conditioned medium. This speeds the cloning process, and this way, the clones are truly independent. We usually put 1 ml of undiluted culture into the first row of wells and then make a 1:1 dilution (0.5 ml original culture plus 0.5 ml conditioned SDM) in the 2nd row. The 3rd and 4th rows just contain serial dilutions of the second row (1:1 or 1:2), again in conditioned SDM. To prepare conditioned medium, pellet cells from a late log culture (7-9 x 106/ml) and sterile-filter the culture supernatant. The filtered conditioned medium can be stored a few weeks at 4o or frozen. Don't forget to add Hygro, G418 and phleo to the conditioned medium when doing the cloning.
The Phleo will wipe out the untransformed background in about 7 days. By about 6 days, you'll be able to see a pronounced difference in numbers of viable cells between transfected and negative controls. You can use Lew79 as a positive control. Let positive wells become very dense before feeding. They will look very yellow and ugly, but it's imperative to resist the urge to feed them. Dilution has killed more transfectants than drug around here. Only when wells are literally teeming with tryps @>4 x 106/ml (about day 9/10 for Lew79 transformants), should you feed. Then start scaling the volume in the well up by no more than 1:1 dilutions. When you have 2 ml, transfer to a standing flask and continue with the gentle dilution. If you take these nascent procyclic clones below about 3 x 106/ml, you'll really slow them down, if not lose them.
Once they're established you can induce anything linked to the Tc-responsive parp promoter by adding Tetracycline. For maximal induction use Tetracycline @ 0.1-1µg/ml. We measure about a thousand-fold increase in luciferase by 12 hrs and levels max out by 48 (slower than with single promoter constructs probably because of differences in the luciferase 3'UTRs). For induction to Pol II level transcription, use about 5-10ng/ml Tc.
Procedure is essentially same as for procyclics except that cells are handled at 37 or room temp. Use 107 cells and 5-10 µg of DNA per 0.5 ml per cuvette. Note that log phase for BFs is in the range of 2 x 106/ml. Cytomix can be used instead of ZPFM. BTX settings are as for procyclics. Cells are zapped, transferred to 10 ml of HMI 9, and surviving cells counted. 50,000- 1,000,000 living cells (depending on the efficiency for different constructs) are then distributed in approx. 0.5 ml volumes into the wells of a microtitre plate. Selection is applied the following day by addition of an equal volume of HMI9 plus 2x drug, to each of the wells. It takes about 5 days to distinguish positive wells, which can be scaled up immediately.
We use Promega's luciferase assay system (cat #'s E1500, E1483, E1501 & E1531), basically following the protocol in their Technical Bulletin # TB161. For luciferase assays from stable expressors, I pellet 1-2 million cells in an eppendorf (30 sec), lyse them with 100µl of the Promega buffer (or 0.2% Triton in PBS) on ice, then pellet the cell debris by a 3-5 minute microfuge spin at 4·. I add 5 µl of the supernatant to 45 µl of the Promega assay buffer, mix and read immediately in the Turner luminometer.
For transient assays, 5-10µg of DNA is used per 1-2x 107 cells per cuvette. Competent cells are prepared and zapped as for stable transfection, then transferred to 5 ml of medium and incubated 12-20 hrs. The entire culture is harvested (4o C, gentle spin) and the pellet lysed as above. The lysate is cleared, and 5 µl added to 45 µl of assay buffer (room temp), mixed and read as above.