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  • #31
    I've just been drying 5 minutes under a hood, spinning down, and resuspending. Ampure XP has been so efficient regardless that I always have plenty of library (usually 10x more than doing the 2nd gel sizeselection, and just as effective at removing the adapter dimers).

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    • #32
      re-using AMPure beads

      Originally posted by yorkzhou View Post
      Theoretically, there in no reason that the beads can not be reused.

      Beckman Coulter can't be happy about this idea. At least they may argue that you need to decontaminate the beads before binding other samples.

      The homebrewed buffer seems to be promising; maybe someone can do some tests.
      There was a paper published this year by the Broad institute, and they re-use the SPRI beads.

      Genome targeting methods enable cost-effective capture of specific subsets of the genome for sequencing. We present here an automated, highly scalable method for carrying out the Solution Hybrid Selection capture approach that provides a dramatic increase in scale and throughput of sequence-ready li …


      BTW I advise anyone doing NGS to find and read all method papers from the Broad institute (Chad Nusbaum), they are the pioneer in NGS techniques and produce really nice & informative publications in (usually) public domain journals.

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      • #33
        Re. size of Ampure beads, I've also been wondering and reading a bit, and by looking through the Hawkins 1998 patent, I stumbled upon Biomag particles (formerly from PerSeptive, that had been acquired by PE, that became ABI a.f.a.i.k.).
        I understood from searching around that they have a mean diameter of ~1.5 µm and are of irregular shape, thereby having a higher surface area than spherical beads. (However, there are varieties of Biomag particles that can be larger.)
        Hope this isn't too far away from the real size of AMPure beads (I would assume Backman buys the particles somewhere).

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        • #34
          Originally posted by edawad View Post
          When using Ampure XP beads, the included protocol says to dry for <5 min after the EtOH wash to avoid beads drying out too much and cracking. However most online protocols I see, plus Illumina's Truseq protocol (which uses Ampure XP) all say to dry for 15+ min until the beads crack. Anyone have any experience what difference this makes?

          Also most protocols say to use fresh 70% ethanol to wash, but the Truseq calls for 80%. My understanding is that the higher ethanol concentration might be less efficient at washing away smaller molecules. Has anyone played around with this?

          thanks
          We've had experiences with both: completely dry cracking beads, and also still moist. We can't seem to see a difference in yield between the two using AMPure XP (we haven't used regular AMPure). We use 70% EtOH for Ion Torrent library prep PGM workflow, and 80% EtOH for Illumina library prep Hiseq workflow just because it's in their protocol. We've adapted the Ion Torrent methodology on the Illumina purification protocol: Following aspiration of the last EtOH wash on the magnet, we spin the tubes down and then place back on the magnet. Then suck up the residual EtOH. This allows the beads to dry faster. If you stick to the Illumina Tru-seq protocol you'll have to wait 15+ minutes for the beads to dry (or at least for the residual beads to evaporate. I'm not sure about the EtOH 70/80% differences, I'm rather curious myself. (maybe mean bp size for Sheared DNA input?)

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          • #35
            Originally posted by katsigner View Post
            Can the beads be reused?? Any idea?
            Sure, why not - if you can trust them in regard to contamination. I guess a very aggressive nuclease can be used for cleaning, like benzonase or cyanase, but can you trust them then in regard to DNA integrity?
            A cheaper alternative could be carboxyl SeraMag from Fisher dispersed in PEG/NaCl solution, but you will have to titer mixing ratios vs precipitation range. Several papers online can give guidelines. In my experience 25% PEG4000/2.5M NaCl works at DNA:beads ratio of as low as 1:0.5.

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            • #36
              anyone knows the DNA capacity of AmpureXP beads?

              hi, i am using AmpureXP to perform DNA purification to library chip-seq DNA. i change the size selection ratio from 0.9*-0.2* to 0.8*-0.4*to get a wider size range. one problem i have confront is that no matter how much initiation DNA i use(from 10ng to 160ng), i always get an equal amount of DNA(~30ng/ul*17ul) after PCR amplification(15 cycles). i cannot explain this result, can it be due to DNA capacity of AmpureXP beads?
              Last edited by junorose; 12-27-2012, 06:26 PM.

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              • #37
                Ampure beads don't settle well

                Used AmpureXL on Zymmo purified samples (no salts or detergents), got reasonable yields but the beads did not align well to the tube side but rather settled to the bottom.
                has anyone encountered this- what does it mean?

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                • #38
                  Originally posted by junorose View Post
                  hi, i am using AmpureXP to perform DNA purification to library chip-seq DNA. i change the size selection ratio from 0.9*-0.2* to 0.8*-0.4*to get a wider size range. one problem i have confront is that no matter how much initiation DNA i use(from 10ng to 160ng), i always get an equal amount of DNA(~30ng/ul*17ul) after PCR amplification(15 cycles). i cannot explain this result, can it be due to DNA capacity of AmpureXP beads?
                  I think it more likely that you've exhausted your primers during your PCR rather than bead saturation. 15 cycles is a quite a lot of amplification.
                  1µL of AmpureXP should contain enough beads to bind a few µg's of nucleic acid.

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                  • #39
                    Originally posted by ooriw View Post
                    Used AmpureXL on Zymmo purified samples (no salts or detergents), got reasonable yields but the beads did not align well to the tube side but rather settled to the bottom.
                    has anyone encountered this- what does it mean?
                    What kind of magnet are you using?

                    Comment


                    • #40
                      Originally posted by TonyBrooks View Post
                      I think it more likely that you've exhausted your primers during your PCR rather than bead saturation. 15 cycles is a quite a lot of amplification.
                      1µL of AmpureXP should contain enough beads to bind a few µg's of nucleic acid.
                      i uesed 25pmol primer,both universal and index primer, in theory, it should be plenty enough; last time i did this experiment, i first added 50ul water to 50ul pcr reaction, then perfromed Ampure XP clean up. i got as much as 55ng/ul *17ul DNA.

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                      • #41
                        Originally posted by junorose View Post
                        i uesed 25pmol primer,both universal and index primer, in theory, it should be plenty enough; last time i did this experiment, i first added 50ul water to 50ul pcr reaction, then perfromed Ampure XP clean up. i got as much as 55ng/ul *17ul DNA.
                        15 cycles is still a lot. We have over amplified with less cycles and similar amounts of primer. It would depend on how much of the PCR input is competent. Have you run the samples on the Bioanalyser? Do you see an over amplification bubble?

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                        • #42
                          Anyone have any experience/thoughts on the new Agencourt product SPRISelect? Is it just higher quality beads of Ampure XP or is something new in the buffer to help with size specificity? All I see is a $50 more expensive product that is already over priced, but I digress....

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                          • #43
                            I heard there should be less lot to lot variation that you sometimes see when doing size selection with Ampure XP. So in other words, sounds like there is more QC that goes on for SPRISelect. Not sure if there are differences in buffer etc...

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                            • #44
                              As mentioned earlier with Ampure XP, we were seeing inconsistent results. Our recent testing with SPRIselect has been going well and the data so far looks amazing! What seem to work best for us is 0.8X and 0.61 double SPRI. Peak at 300 +/- 100bp.

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                              • #45
                                The bead is overdried and there is no DNA in the supernate.What should I do?

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