the long story short, I needed a detection assay for a certain mutation, preferably as a commercial kit, and the only cost-worthy for this matter were finally ABI Custom SNP Assays (I know they're for genotyping, not a mutation detection but given the problem set-up, I didn't needed too much sensitivity, either there would be a mutation in majority of cells within sample, or I don't care).
Now, if you don't know it designing a custom assay for somethig other than SNP on the ABI pages just suck. No, SUCK. But anyway, I needed an assay to distiguish a one-base deletion.
I also designed other two custom SNP assays this way, for C>T substitutions and they work rather fine. This one does not.
ABI SNP assays have two allel-specific Taqman probes with MGB so the point is to get "high VIC-low FAM" for wild-type samples and "probably some VIC - high FAM" on the mutated ones. For two-colored genotyping there is a nice plot, that software then clusters.
It should look somehow like this:
It doesn't. It looks like this:
(they have both high fluorescence for the allele, that should not be there, it even called the "100%" mutant as a heterozygote)
If you draw line there you see both "extremes" are too close. I would have a real difficulty to tell any sample that will fall between 100 % wt and 100% mutant, and if there is one possitive, it surely will.
From the look at the amplification plots, it's obvious that in contrast to the first assay, where in the corresponding channels the florescence of the "unmatched" allele reached plateau in much lower fluorescence level (so increasing/decreasing cycles would have no further effect), in this assay they seem to have a similar efficiency to the "matched" allele.
Also, FAM and VIC channels crosstalk, but the graph is designed to take care of that, because it displays fluorescence from both channels in kind of visual ratio, so this cannot be the sole effect of channel crosstalk.
It seems to me, that the probes are just not specific enough, and that it's something I can't really affect much.
I have definitelly ideas how this could be done really right, and is not, but due to the situation won't be and it's not even worth it really (since it is part of the I will never ever again... project) so I want to get quicky over that or just leave it.
For example, I can't be sure, if my standard is really 100% mutant, it only theoretically should be as it was created by OE-PCR mutagenesis and diluted more than 1000x (no cloning of such useless thing, really..), but of course I'm not sure. But from the graph, it seems in general, that samples have to high fluorescence of the "unmatched" allele, than they should have. (I have no access to patient sample with this mutation to test)
So, since the whole amplicon looks like this:
(yes, this short)
orange - primers, blue - probe(s), red - the C, that get's deleted
I wondered, if the stretch of Gs could do some harm, either to assay and the OE-PCR before, so I'm now running it side by side with 3% and 5% DMSO (still don't have betaine) to see if it helps anything, but then I have no further ideas. Of course, the main reason could be that the design doesn't work (and they only tested that it somehow runs on wildtype samples) and I can't do anything, but if any of you have some idea how to help it, that's not lenghty I would appreciate it.