Over my last couple dozen amplicon libraries, I've had an average filter pass rate of about 50-55%. There were a few that were in the low 40's and I had three others where I got a filter pass rate in the low 70% range (don't ask me how that happened; just a good day, I guess), but most are a little over 50%. It seems the rates have been a little higher since I started using the Fluidigm Access Array for the amplifications.
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That is pretty good for amplicons, I think. Are you using Lib-A or Lib-L? We´ve done so far only 16S amplicons using the one-way approach and using the protocol without modification, like using less primer for amplification. We also use amplicon pipeline for analysis. I think our best was around 47% (whole plate avg)
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I've used both. I originally started out with the standard chemistry, then switched to Lib-L adapters when I started using Titanium chemistry. With the Lib-L adapters I found that the sequence quality wasn't quite as good, particularly in homopolymer regions. Reducing the amount of primer helped. When we got the Access Array last year I switched to Lib-A adapters and I've noticed a small improvement in sequence quality, especially in homopolymers.
I also use the amplicon processing pipeline. If I use the standard pipeline, the filter pass rates are higher, but I find that I don't have any more useful reads. The extra data obtained by using the standard pipeline doesn't help, but can make analysis harder by introducing noise.
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Originally posted by ajthomas View PostI had a bottle of the RNA SPRI beads that I knew I wasn't going to use before they expired so I called to ask the company if they would work for DNA. I was told that they are the same product, except that the RNA beads have been produced to be sure they are RNAse free and are checked for RNAse contamination, so yes they should work fine for DNA. Assuming that's true, the reverse should also be true--that you should be able to purify ssDNA with AMPure beads. I suspect the ratios will need to be adjusted, but I don't know that for sure.
Thanks for the info! I’ll try to run some tests with the AMPure and denatured DNA, along with the HS DNA and RNA Pico chips and see what happens.
It seems like 45-55% passing filters is pretty standard with amplicons? Our best was 61.9%, with an amplicon only 150bp long. Wish I could get > 50% all the time!
Anthony
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Originally posted by Anthony.287 View PostI’ve had trouble in the past getting the enrichment low enough to even bother with sequencing, but between 0.01 and 0.05 cpb seems to be the sweet spot (after quantifying with the KAPA qPCR kit) for 8-20% enrichment.
Then, based on this, you conclude that your calculation of "copies" is being thrown off by some small (primer dimer) amplicons? If the primer dimers are only 10% the length of the full length amplicons then they fluoresce only 1/10th as brightly during qPCR.
I ask, because that seems like what is probably happening, but I am not clear on whether that is the accepted outlook.
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Phillip
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Originally posted by Anthony.287 View PostAfter following this thread, I’m wondering if it would be feasible to denature the DNA immediately prior to the AMPureXP purification? Will the AMPureXP beads bind ssDNA, or would the RNA SPRI beads be an option? I’ve had some issues with small fragments that don’t show up until sequencing, and AMPure-ing denatured DNA seems like it might alleviate the problem, if it would work.
So there are lots of factors in play.
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Phillip
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Originally posted by pmiguel View PostSo, the issue you are wanting to address is that at 0.01 cpb you would expect a maximum of 1% enrichment, but you are getting results at least 8x higher than that? That is, with one 1 copy of your amplicon per 100 beads, at best, you expect 1% of the beads to be templated.
Then, based on this, you conclude that your calculation of "copies" is being thrown off by some small (primer dimer) amplicons? If the primer dimers are only 10% the length of the full length amplicons then they fluoresce only 1/10th as brightly during qPCR.
I haven't actually tried 0.01cpb, but with a recent 16S library, it appears that somewhere between 0.01 and 0.05cpb would work best; this will change if I can remove more small fragments.
Anthony
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Originally posted by pmiguel View PostI am very interested in your results. The issue I am concerned about is the primer dimers reannealing to the large fragments during AMPure treatment. Seems like the high salt concentrations in PEG precipitations, like AMPure, would encourage strands to re-anneal.
So there are lots of factors in play.
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Phillip
Anthony
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Originally posted by TonyBrooks View PostWe routinely do 16s runs and they almost always look good. I agree with Philip that primer-dimer seems to be a major problem in all 454 amplicon work. All our amplicon libraries undergo gel-cut post pooling. The one library where we didn't do this gave us problems.
FYI I've attached results from our typical run. We use software version 2.5.3 with everything set as default. You have to remember that the Bioanalyser is a plot of DNA mass against size. What's important is not mass of DNA but molarity, so small peaks that are low in length are actually a big problem as they contain a high number of molecules. These are also favoured in emPCR over the longer amplicons you want to sequence. Also I don't think the Bioanalyser is sensitive enough to pick up any low level dimerisation.
It's might also be a good idea to also keep the results of the first NaOH wash during the bead enrichment step. This will contain the reverse strands of the emPCR which you can then run on a gel or Bioanalyser. You will probably need to concentrate on a Qiagen column first after neutralising with a small volume (40ul) of 3M sodium acetate (make sure the Qiagen indicator solution is yellow).
I am at a loss for why the Ampure and the PippenPrep were not sufficient for removing these fragments. Any suggestions would be greatly appreciated.Attached FilesLast edited by sequencingerrors; 02-28-2012, 08:16 AM.
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Quick update on the AMPureXP test with denatured products – it doesn’t seem to work. My guess is that the AMPure and DNA don’t bind well, if at all, below a certain temperature. At the end of the process, the pools that were denatured retained very little, if any, DNA, while the controls still had plenty.
If anybody wants actual details, let me know! I’ll be happy to share.
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Originally posted by Anthony.287 View PostQuick update on the AMPureXP test with denatured products – it doesn’t seem to work. My guess is that the AMPure and DNA don’t bind well, if at all, below a certain temperature. At the end of the process, the pools that were denatured retained very little, if any, DNA, while the controls still had plenty.
If anybody wants actual details, let me know! I’ll be happy to share.
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Phillip
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Not quite. I denatured the DNA at 95C for 2 min, then 5min on ice. I added the AMPure while the sample was on ice (5 min) then took it off the ice and let it incubate at room temp for 10 minutes. I then proceeded with the AMPure protocol like normal. I also included controls of the same DNA that was not denatured. When I ran the samples on the Bioanalyzer when all was said and done, the controls had nice, strong peaks where they were expected, and the denatured samples had nothing. I repeated the test with new sample pools spiked with a 100bp ladder as an additional control, with the same results.
On a related note, does anyone have experience using the Roche Sizing Solution in place of the original AMPureXP buffer?
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Originally posted by Anthony.287 View PostNot quite. I denatured the DNA at 95C for 2 min, then 5min on ice. I added the AMPure while the sample was on ice (5 min) then took it off the ice and let it incubate at room temp for 10 minutes. I then proceeded with the AMPure protocol like normal. I also included controls of the same DNA that was not denatured. When I ran the samples on the Bioanalyzer when all was said and done, the controls had nice, strong peaks where they were expected, and the denatured samples had nothing. I repeated the test with new sample pools spiked with a 100bp ladder as an additional control, with the same results.
On a related note, does anyone have experience using the Roche Sizing Solution in place of the original AMPureXP buffer?
Maybe it is your DNA being single stranded? That would mean half the molecular weight. My mental model for how a PEG precipitation happens is PEG occupying a percentage of the "solvation" sites sufficient to begin to force other molecules out of solution. Then the longer the molecule the more water molecules are needed to hold it in solution. But a single stranded molecule should take less water molecules because, well, only one strand to keep solvated. That would suggest it would take more PEG to precipitate single stranded molecules. Right?
The experiment I have not seen done is running the AmPure calibration curve on a single-stranded ladder. Maybe one could use the one included in the BioAnalyzer nano-chip kit?
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Phillip
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