I did a bulk mRNA-seq experiment where I treated human cells in vitro with control of drug. I am only interested in determining differential gene expression at the gene level between these groups. I isolated RNA from treated cells and used poly-A selection and did sequencing on the illumina platform. I aligned the raw reads to the human genome using STAR. I used the featureCounts subpackage of subread to to get a count of reads that overlap with genes, the resulting gene count matrix is what I fed into edgeR to get differential gene expression results.
Now I would like to design some primers against a few select genes for qPCR to validate my differential gene expression results.
I am familiar with using Primer-BLAST, but I have previously only used it to design primers that span the exon-exon junction of a given transcript isoform by supplying the NCBI Reference Sequence of that isoform to the Primer-BLAST tool.
If I follow this approach I am afraid of designing primers that span the exon-exon junction of an isoform I don't even have in my sample.
I have also produced the equivalent of an "exon-count-matrix" with featureCounts by counting at the exon level rather than the gene level. I guess I could design primers to bind within a specific exon that I know is expressed, but then I worry about amplifying contaminating genomic DNA (which I realize might not even be an issue because I did a genomic DNA elimination step in my RNA isolation).
How do you usually go about designing primers for qPCR based RNA-seq validation?