I bound competent cells (H. influenzae and B. subtilis, separately) onto cover slips in chambers, and then washed in medium containing genetically marked DNA. After 15 minutes at 37 °C I washed in some medium with DNase I and then medium containing 0.5% low-melt agarose, with and without either nalidixic acid (to select the H. influenzae nalR allele) or tryptophan (to select the B. subtilis trp+ allele). Then I sealed the chambers' ends with wax from a candle (clumsily) and incubated them overnight.
The results were a bit messy. The H. influenzae cells started out quite dense (1000-5000 per 40X field of view) and without nalidixic acid they grew into nice microcolonies. With nalidixic acid they instead formed filaments (i.e. grew without dividing, and then stopped growing). At the edges of the chamber some large and well-defined microcolonies were present, but these were absent from the central part of the chamber. So transformation is clearly happening, but I can't estimate a frequency. The distribution of the NalR microcolonies suggests that oxygen might affect resistance, but I don't know enough about how the drug acts (a gyrase inhibitor) to guess why that would be the case.
The B. subtilis results were worse. The chambers with and without tryptophan had similar numbers of cells. Both had substantially more cells than had originally been attached, so there must have been some growth. But there were no obvious microcolonies. I might have made an error with the medium. But many of the cells were moving around, so I suspect that I also need to use a higher concentration of low-melt agarose to block their strong motility.
On a separate topic, a reader suggested using polyethylene glycol (PEG) to block the poly-L-lysine coating and prevent beads from sticking to it once the cells had been bound. After checking that PEG isn't toxic to cells, I tried washing my chambers with a 1% solution of the PEG type we had on the shelf (PEG 3350, which I think is a moderate chain length for PEG), washing it out, and then washing in some beads and washing them out after 10 minutes. Result - no significant difference between PEG and no PEG in the numbers of beads bound to the coverslip. I could try a higher concentration of PEG, but maybe the problem is washing out the PEG before adding the beads. I'll try adding PEG to the beads as well as to the washing solution. If this does prevent beads sticking, I'll next need to test whether PEG inhibits transformation.
While I was at it I also tested whether the 16% glycerol that's mixed with the frozen competent cells inhibits cell binding to the coverslips. I had been conscientiously pelleting the thawed cells and resuspending them in glycerol-free medium before adding them to the chambers, but now I know that cells bind just fine in the presence of glycerol. The glycerol is subsequently washed out of the chamber along with the non-attached cells, so I won't bother with the centrifuging step any more.
And I tested alternatives to sealing the chamber ends with wax, which is difficult to apply and tends to form big lumps rather than a smooth layer. (I don't like the risk of getting wax on the microscope lens.) First I tried a better way of applying the melted wax - rather than using a glass Pasteur pipette with a rubber bulb, I tried using an old Pipetman p200 with a snipped-off tip. This gave better control, but the wax still formed a big lump when it met the glass. Parafilm didn't work - a film of liquid quickly seeped under the parafilm. Melting the parafilm with a heated spatula didn't help. I also tried using paint from a paint-Sharpie; this only sort-of worked, perhaps because it's a water-based paint. (But it was cool to look at the paint droplets under the microscope.)
9 hours ago in The Phytophactor