We used CRIPSR editing to make a 2 nt change in a mouse genome, and identified a positive pup by PCR on tail-snip DNA, using a primer that ended in the 2 nt mutation, which would not amplify wild-type DNA. We bred this mouse expecting to get heterozygotes, but when screening ear punch DNA cannot find any positives by PCR. We amplified the region containing the mutation, isolated a single PCR amplimer, and used Sanger sequencing to screen the amplifiers. Using the forward primer for sequencing all pups read unambiguous wild type. However, using the reverse primer we see a divergence at the mutation site, where we see traces of equal height for the mutation and the wild type sequence, as you might expect for a heterozygote. I cannot figure out why one strand looks wild-type and one appears to contain the mutation. The primers used to generate the amplimer do not match any other position in the mouse genome. Any theories for this apparent discrepancy between strands?
Does this work?
The forward primer for the amplimer containing the deletion position anneals with its 3'end or nearby base over a natural polymorphism. This polymorphism is on the mutated allele. Under the pcr conditions you amplify both alleles hence the reverse sequencing primer sequences both alleles. Under sequencing cycle conditions, however, the primer does not anneal properly over the polymorphism so only the wild type is sequenced. The poor annealing of the primer on the polymorphism if the mutation is on that allele is present but cannot sequence at your annealing temperature. Clone and sequence or design a new forward sequencing primer
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