I need to determine the number of integration sites of the plasmid DNA into the genome of a CHO cell line. After selection, the population of zeocin resistant transfectants were cloning by single cell and clones verified by Southern blotting.
Developing X-ray film revealed single-site, two-site and more, but each band shines as 50-100 copies, compared with the positive control - plasmid DNA, corresponding to one copy per genome.
Any suggestions as to what I should do?
Have you tried a more precise method like Real Time Quantitative PCR on your genomic DNA? You can design primers for the zeocin resistance marker and compare it to a single copy gene such as beta actin or 36b4.
Given that your dilution row and everything is OK, the differences you see may be due to different efficiencies at which your fragments are transferred to the membrane (generally, larger fragments are harder to blot). Are your integration fragments much smaller than those deriving from your plasmid DNA?
If possible at all, do qPCR (you don't need to do absolute quantitation, only relative quantitation against a reference single copy gene). Quantitation via Southern blotting is too tricky, as you have to account for differences in transfer efficiency, non-linearity if using x-ray films, etc. and is too laborious if you have to process many samples.
I am not sure I totally understand the question, but if you want to know the number of integration sites by Southern blotting it is important that you use restriction enzymes which don't cut the plasmid sequence (probably try 6 & 8 cutters). Otherwise, if you have multiple copies at an insertion site you will get multiple bands.
The plasmid you have loaded is just to check that the labelled probe is working which you can do a dilution as Andrei suggests. Alternatively, you could carry out a fluorescent in situ hybridization which is a lot easier!
- Badges
- Science topic