Is there any consensus of opinion on the best RNA-Seq approach for quantitation of alternative splicing on well annotated genomes (e.g. human)? We don’t want to look for novel splice isoforms, we want to detect regulation of alternative splicing by our treatment among the annotated isoforms of a gene. The options we have are:
30M x 50 bp single end reads
30M x 50bp paired end reads random primed cDNA
30M x 125 bp paired end reads using strand specific cDNA library
The cost of the last two are similar, roughly double the first one. I guess in part this depends on the method chosen – something that just counts exon inclusion should work just as well with single ends. If you look for exon exclusion then you need reads that map across splice junctions, I don’t know if the chances of that are improved more by increasing the read length, or increasing the read depth (naively it would seem doubling either would have the same effect). Finally I don’t know what the benefit of strand specific libraries are for quantitation, one concern is that if they are more involved to produce then they might introduce variability.
hi,
my experience, do whatever, have replicates, save day. Having said that, shorter length would give more read numbers (more X sampling of the transcriptome) and this might be useful in case you want to call variants. If you are interested in alternate 3prime ends (where many a times repetitive sequences reside), longer read legth would be useful. But all this would be best for your hypothesis if you keep replicates to apply statistics.
Strand-specific I think you would go if you are interested in teasing out antisense transcription. In that case you would have to look into total RNA (rather than polyA select) as input. Sequencing total RNA (even after ribo minus) leaves enough amount of ribosomal RNA (ribosomal repeats in fact) to eat into the depth of transcripts of your interest. So, you would need to do very high depth sequencing if effect of ribo RNA is to be negated.