Oligo insert cloning - (Mar/06/2013 )
I've been trying for months now to insert a 122bp ds oligo into a 8000bp vector but it just doesn't work. I tried vector:insert ratios from 10:1 to 1:5, different ligases/buffers and also changed the time for ligation (2h - over night at RT, 4°C and 16°C). But nothing worked so far. So now I think there might be something wrong with my oligo design.
The setup was as follows:
2 single strand oligos, annealing: 10 nmol each, 5min 99°C, then let cool down to RT (4h). I used annealing buffer (
The oligo contains compatible overhangs for BamHI and Pst1 but as you can see both strands were designed 5' to 3' (manufacturers data sheet) so is that the reason why the ligation cannot take place?
The oligos are not phosphorylated but the cut vector should have PO4 groups from restriction digest. Can I do something to use both oligos at the same time or do I have to design at least one with reverse direction (3' - 5')?
Anyway, is there any method to check whether the annealing has worked or not - how can I distuinguish if I have single strand or double strand oligo after annealing?
Thanks in advance!
You can't have double stranded DNA with paralel strands (like 5' end on one side and 3' and on the other as in your picture). The strands would not anneal the way you draw it.
If you planned to have this cassete, you designed it wrong. Oligos are always synthetised (and put in data sheets) in 5'->3' orientation, so you have to reverse the second strand before ordering it.
You needed to order sequence like this to anneal with your first strand:
OK thanks, that was also my thought about this issue. I'm very new to cloning and not a biologist so retrospectively spoken this was a bad design... I'll order a new one.
But still - anyone ideas about how to check wheter after annealing I have still single or double strand oligos?
Intercalator dyes like SYBR Green I binds double strand much more efficiently than single strand. By running a melting curve you could see the differences in fluorescence. But you would need something to compare with.