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  • #16
    Zippy, did you use rapid libraries for your shotgun? If so, were you sure you did the denaturation step in the emPCR set-up? A low enrichment, plus such a high proportion of mixed reads would point to non-denatured rapid library in the emPCR. You would have dsDNA going into the emPCR with each strand being in opposite directions, hence all reads would come out as being mixed.

    There is a theoretical limit for how many beads you can get back from emPCR, so it could be that you're well beyond that limit. Reducing the input may not result in the same proportional drop in enriched beads. For our 16s studies, we've found that using the Bioanlyser sizing, plus a Qubit/Picogreen quantification gives us hightly reproducible results on our FLX. We pretty much get 10% enrichment +/- 2% every time with 0.5cpb.
    If in any doubt, use qPCR (such as the Kapa kit) but be aware that we've found different library types require different emPCR input.

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    • #17
      Many emPCR reactions can be started with dsDNA, the oligo bound to the bead is usually the B' or A'. Starting emPCR with ssDNA just avoids too much solution phase PCR. I'm not sure how you could get reads in all directions as described.

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      • #18
        Well, I could not stand the lack of logic in my experience described in the previous post and made a seq run using beads that by the protocol must have been considered failed because I got about 4-6 million enriched beads. Guess what? I got the best result so far with this ancient DNA, with 23% passed filters, while with all these diluted samples before the best was 17%. The distribution of reads was quite intersting though, with two maximums, one around 450-550 bases and another around 100 bases, with minimum around 250 bases. The total number of reads was about 50% of what I got with the worst case (1% usable reads).
        In fact, after reading the original paper and the emPCR protocol I realized that it is quite possible to get over 2 million enriched beads, contrary to what is stated in the protocol. The original paper says they got about 30% enrichment. The kit does not say how many capture beads are supplied, but assuming that the formula provided in the manual gives a real example, one gets 10 million beads with the kit. So, simple math gives you 3 million beads of normal enrichment output.
        I also feel that amount of beads one gets is quite dependent on how harsh the washing steps are performed. The enrichment beads are likely 0.8 micron streptavidin magnetic beads (I scoped them), much smaller than ~30 micron capture beads, meaning that strength of binding to a magnet will depend on how many brown enrichment beads are attached to the white capture beads. Instead discarding unbound beads right away as recommended, put these in another tube and stick into a magnetic separator - you will be surprised how many almost white beads will keep sticking to the magnet, meaning these also bear amplified library fragments! The harsher wash the more of these weak-binding beads will go down the drain.
        I feel the lack of logic in my experience has something to do with small fragments present in library that distorts library quantification. Perhaps, sizing did not work the way it is supposed to work. I found one reference, which uses double-step sizing, so I want to try it to see if I get more logical results.
        I do not think that I have broken emulsions. I get very consistent picture every time - white sediment and clear oil layer on top.

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        • #19
          OK, again another curious result. I used a totally different sample library this time, made from DNA extracted from an about 100-old tooth, as compared to previous excercises with ~1000-old bones/teeth. Library concentration was estimated to be at ~1.5e7 mol/ul, so I used 1 ul. From the enrichment step I got about 3 million beads (by GS Jr bead counter), which means my emPCR supposed to be failed again. Nope. I did a seq run with recommended 0.5 million of these beads - 37% reads passed all filters. Out of 173.5K reads (E.coli traning generated 240.5K reads), the largest loss was not after the mixed filter but after the dot filter. Overall distribution was 45.7K lost at the dot filter, 21.8K lost at the mixed filer, and 25.5K lost at the short quality filter. Read lenghts peaked at 500 nt, with a minimum around 300 nt and a substantial number of reads of 50-250 nt. The longest read was 1038 nt, not bad at all. This likely means that at about 30% enrichment the picotiter plate was still not loaded to its full capacity (in terms of beads carrying amplified fragments), many wells had beads with either too short fragments or produced no signal. Which means that the manual has a very narrow applicability and considering the variability of samples everyone is basically on his/her own to get the best from his/her libraries. Do not throw away your purportedly failed beads! Question authorities...

          I keep y'all posted on furter developments.

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          • #20
            epistatic, it was my understanding that the Y adapters contain both the A and the B sequences, annealed at one end. When ligated to an adenylated dsDNA library, this would cause both strands to have the A and B adapters attached, as such:

            A-LIBRARY-B
            B-LIBRARY-A

            If you didn't denature, then both strands would go into the micro reactor, anneal to the same capture bead and then amplify. Sequencing using the A primer, would then give one read: LIBRARY and another read YRARBIL on the same bead, which would be mixed signal.
            It's different for amplicon libraries as only one strand of the amplicon would contain the A and B sequences in the correct orientation. ssDNA libraries only have one strand anyway, so there's no issue with mixed reads.

            yaximik, we did a run recently where we had large numbers of small fragments in our library. We used the Bioanalyser/Qubit to estimate concentration, using the largest peak as our average size.
            Our enrichments were much higher than normal, (approaching 30%). We believe that this was because our quantification was wrong (smaller library lengths mean a higher molecule/µL per ng of DNA). We sequenced anyway and got much higher dot fails than normal.
            Roche said, and I quote "These shorter fragments will lead to brighter signals, ending up in elevated dot filtration".
            I think, you may need to be more stringent in your method of size selection. Ampure XP beads may not be removing all the small library fragments. Some of your read lengths do look a bit on the small size. Perhaps you should look at a gel-cut option for a run and see if you can improve your selection.
            Can I ask how you are estimating your library concentration and counting your beads? Do you get a stable raw read output from all your runs?

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            • #21
              My undrstanding is that Y adapter is not simply double stranded AB, and the sequencing primer site is on one end only. Tired from a lot of confusing or mutually exclusive responses from Roche specs about rapid library adaptors, I simply sequenced a few subcloned library fragments, the deduced adaptor is quite different from AB adaptors.

              I found a useful paper on double step sizing, do not have it handy to attach, but it worked very well even with already made libraries, so I am going to sequence one of these resized libraries if this makes a difference.

              I use SYBR qPCR with emPCR primers to quantitate libraries, using one of cloned library fragments as standard and recalculating based on the peak ditribution size of fragments by Bioanalyser2100. The problem is I cannot get below 1e5/ul in sensitivity yet, although vendors of mastermixes claim 1e2.

              Answering your question - I do not get consistent raw data, perhaps because I work with very small amounts of pretty bad DNA. Buried in the soil bones are not the best source of DNA. So each library requires several iterations to figure the right ratios. I am counting beads using GS Jr bead counter.

              Comment


              • #22
                Originally posted by yaximik View Post
                My undrstanding is that Y adapter is not simply double stranded AB, and the sequencing primer site is on one end only. Tired from a lot of confusing or mutually exclusive responses from Roche specs about rapid library adaptors, I simply sequenced a few subcloned library fragments, the deduced adaptor is quite different from AB adaptors.
                You can get the adapter sequences by looking at the IDT website. They are the approved suppliers of the extended MID adapters (i.e. >MID012) and they give you the sequence of the oligos when you get to the shopping cart. Ignoring the MID barcode, they do align at one end to potentially generate a Y shaped duplex. Interestingly, the region of alignment also contains a number of deoxyinosines.

                Comment


                • #23
                  Problem - Very low number of enriched beads

                  Hi,
                  Related to bead enrichment (for the Roche Junior), we're having the problem where we are not getting enough beads, essentially its looks like the vast majority of the beads is being lost. We're following the rapid library protocol and the DNA QC and quantitation look good, pretty much identical to the example Bioanalyser figure (Roche RL prep manual). The emPCR emulsions look good, no broken emulsion, but something happens (i don't know what?!!) between then and the collection of the enriched beads, we seem to lose almost everything?

                  We've done the 8kb PE protocol a couple of times and they've worked, but when working from the rapid library its just not working?

                  We following the example in the emPCR lib L protocol for determining the volume of DNA library, using 2 cpb as our target? To do this we dilute the aliquot prepared (based on the Roche calculator) which is set to 1x10^7 down to 2x10^6 molecules/ul.

                  Anybody have an idea how to resolve this?

                  Comment


                  • #24
                    Originally posted by Raj View Post
                    Hi,
                    Related to bead enrichment (for the Roche Junior), we're having the problem where we are not getting enough beads, essentially its looks like the vast majority of the beads is being lost. We're following the rapid library protocol and the DNA QC and quantitation look good, pretty much identical to the example Bioanalyser figure (Roche RL prep manual). The emPCR emulsions look good, no broken emulsion, but something happens (i don't know what?!!) between then and the collection of the enriched beads, we seem to lose almost everything?

                    We've done the 8kb PE protocol a couple of times and they've worked, but when working from the rapid library its just not working?

                    We following the example in the emPCR lib L protocol for determining the volume of DNA library, using 2 cpb as our target? To do this we dilute the aliquot prepared (based on the Roche calculator) which is set to 1x10^7 down to 2x10^6 molecules/ul.

                    Anybody have an idea how to resolve this?
                    If your PE libraries are working, that would suggest a problem with the library, rather than emPCR. We use the Kapa qPCR kit to quantitate/QC our rapid libraries. I don't think a Bioanalyser trace alone is enough to confirm a "good" library. All it does it confirms the Ampure size selection worked, it says nothing about how well your ligation went.

                    Comment


                    • #25
                      Hi TonyBrooks,
                      Thanks for getting back to me so quickly!
                      I understand your point regarding the bioanalyser. I should have said that in addition we perform library quantitation using the standards and method outlined in the protocol, the concentration of our library (molecules/ul) fits in the middle of the standard curve. We use the 454 calculator to work this out as well as the dilution needed to get it down to 1x10^7. Have you found this method to be "not the most accurate" or to say far enough out to cause the problems we're experiencing?

                      Comment


                      • #26
                        Originally posted by Raj View Post
                        Hi TonyBrooks,
                        Thanks for getting back to me so quickly!
                        I understand your point regarding the bioanalyser. I should have said that in addition we perform library quantitation using the standards and method outlined in the protocol, the concentration of our library (molecules/ul) fits in the middle of the standard curve. We use the 454 calculator to work this out as well as the dilution needed to get it down to 1x10^7. Have you found this method to be "not the most accurate" or to say far enough out to cause the problems we're experiencing?
                        I've never quanitified using the Roche method, so I can't really comment. My major beef with the that approach is that it only counts the adpaters. It doesn't tell you how the library will perform in PCR. It's generally agreed that qPCR is the most accurate measurement of library quantification. However, you will need to empirically determine the correct cpb number from your emPCR result though. We've found qPCR to be invaluable in assessing libraries and now do it as standard for all our rapid libraries.

                        Comment


                        • #27
                          Dredging this thread from the depths as have some updates that might be useful:

                          We've been having problems with really random runs (16S amplicons) on our Junior and have solved a couple of problems that might help others, will cross post this to seqanswers too:

                          1. Short read spikes that suck up lots of reads. Down to primer coming through from the PCR. We do an ethanol precipitation to pool together quadruplicate PCRs, quantify and produce the normalised pool of multiplexed amplicons. This is then followed by two rounds of Ampure XP bead purifications with a harsher size selection than recommended in the Roche protocol - 0.7:1 beads:sample (our amplicon is 676 bp including primer). This has removed the short read spike almost completely.

                          2. Mixed reads and random runs. These have been reduced considerably by using the eppendorf plates and caps recommended by Roche in their 'things required but not supplied' list on the my454 site. The Abgene plates and seals that serve us well for conventional PCR interfere with the emPCR. The belief is that releasing solvents used by plate manufacturers can cause a slow breaking of the emulsion that does not result in the clear banding pattern, but does result in increased mixed reads. Using these plates and caps solved this problem, and there was also a noticeable change in the consistency of the bead suspension during oil and breaking (less clumpy and easier to handle) and fewer washes required during the enrichment step to remove non-enriched beads. Hopefully they're going to start including them in the kits as they have such an impact on the run.

                          3. Broad size distribution in shotgun processing of amplicon reads. Shotgun processing allows trimming, which is useful for 16S amplicons, however we and others have seen a broad size distribution of reads, which doesn't make sense as the amplicons should all be the same length.

                          As we have a long amplicon that we're sequencing, slightly over the suggested maximum of 600 bp, on Roche's recommendation we changed to a version of the long amplicon emPCR method in Tech bulletin 001-2011. We changed the thermal cycling program and increased the buffer amount slightly (presumably to effectively increase magnesium concentration a little), but kept the sequencing primer concentration as it was for fear of increasing short reads. It's not possible to translate the method in there directly to the Junior anyway, as not enough of each of the reagents is included in the kits. Now, instead of seeing the broad size distribution with shotgun processing that we had before, the size range is similar to that of amplicon, but with a couple of thousand more reads. It looks like the changed elongation and annealing temperatures allow for more complete extensions of the amplicons.

                          I would guess that even if you don't have an amplicon over the 600 bp in length, but you see the broad size distribution range, then you would probably benefit from a painless adjustment to the emPCR protocol.

                          Hope that's useful to someone!

                          Originally posted by Palecomic View Post
                          OK - to update, we have the same problem again. A very small volume of clearing at the bottom of the emPCR, no classic banding of the emulsion and this time it is possible to see a small pellet of beads.

                          I will be setting up the same library yet again with an older kit to double check that it's not other aspects of our work flow.

                          In the meantime if anyone comes across a similar problem to that detailed by myself or Yaximik it would be really useful if you could mention it here as well as to Roche. The lot number of oil I have a suspicion of is 93807320.

                          Thanks!

                          Comment


                          • #28
                            Hello,

                            We are doing sequence capture and then sequencing on a GS Junior. We have had many problems with the sequence capture and are trying to work that out. As far as the bead issues, we get no correlation between SV titrations and full scale MV emPCR. We found in our case that we almost always need to use 0.1cpb which has been yielding in the 10% -18% enrichment. they also had told us that 2cpb was optimal, but it is WAAAAAAYYYYY too much in our case. Anyways, because we get no correlation between SV and MV, we are considering skipping the SV altogether and going directly to MV.

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