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We've done 36mers until now and we just got the data from our first 75mer run. We had only 20% of alignable reads and when trying to figure out why this was so bad we noticed the quality scores dropped in a big step after cycle 41. I know the reagents come in sets of 36 cycles but I remember from a meeting with the Illumina rep something about reagents having to be added after n cycles. Could this be it? Anybody seen something like this? When we we trimmed down to 40mers we got ~85-95% alignable reads to the genome, so the last 35 cycles generated unusable data. Thanks in advance for any suggestions you might give me before I get back to the lab that runs our samples.
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We routinely run out to 120 cycles for genomic & mRNA and we don't see drops until we get out past 100 cycles, usually around 110-115 we start to notice a drop, but it's not significant enough to effect our data like what you're describing.
Did you have a control on the flowcell, or a PhiX spike? If so did it also show the drop in mapping? Is the drop due to a lot of 'n' bases in later cycles?
Reagent kits are usually combined & loaded at the beginning of the run, unless they're running with an old chiller that can't hold all the reagents. It's possible they had to add a second kit that wasn't as robust as the first, but it's unlikely.
Do your intensity plots indicate a sharp drop at a certain cycle? If the drop was across the whole flowcell it could be an instrument problem. However, that is difficult to diagnose unless you have access to the images, Status.xml, and reports folder from that run. I'd contact whoever performed the run for you immediately so they can review the logs before they are deleted.
good luck!
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