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Old 07-12-2011, 02:08 AM   #21
niceday
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if the beads dry to cracking it can be difficult to elute the DNA in the time mentioned in the protocol.

Some people make up 70% by eye and because 7ml of EtOH and 3ml of water has a volume less than 10ml they make it up to less than 70%.
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Old 07-19-2011, 08:11 AM   #22
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Originally Posted by katsigner View Post
Can the beads be reused?? Any idea?
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.
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Old 10-07-2011, 02:26 PM   #23
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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
Has anyone tried this out yet or had any thoughts? I'm getting ready to start using Illumina's Trueseq kit as well, and this struck me as a little odd too.
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Old 10-07-2011, 04:52 PM   #24
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Has anyone tried this out yet or had any thoughts? I'm getting ready to start using Illumina's Trueseq kit as well, and this struck me as a little odd too.
I've done a bunch of bead purifications. I see pretty good results either way in terms of drying time. One thing to keep in mind is that if you use 70 or 80% ethanol, the ethanol will evaporate first and the last liquid you see will be water. So it can be a tiny bit wet and you can feel pretty confident that you have gotten rid of all of the ethanol. No idea about which ethanol percent to use.
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Old 10-07-2011, 05:19 PM   #25
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I've done a bunch of bead purifications. I see pretty good results either way in terms of drying time. One thing to keep in mind is that if you use 70 or 80% ethanol, the ethanol will evaporate first and the last liquid you see will be water. So it can be a tiny bit wet and you can feel pretty confident that you have gotten rid of all of the ethanol. No idea about which ethanol percent to use.
Awesome! Thanks for the prompt reply! Hopefully I will have similarly good results.

Woops - should have noted - do you use a short dry time (<5 min like the Ampure XP protocol calls for), or do you go for a longer drying time (i.e. 15 min) as specified in the Illumina protocol? It seems like for such a small volume, this is a fairly excessive dry time, but I'm not sure how it will affect the Ampure Beads.

Last edited by chronicle; 10-07-2011 at 05:21 PM.
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Old 10-14-2011, 03:05 PM   #26
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Awesome! Thanks for the prompt reply! Hopefully I will have similarly good results.

Woops - should have noted - do you use a short dry time (<5 min like the Ampure XP protocol calls for), or do you go for a longer drying time (i.e. 15 min) as specified in the Illumina protocol? It seems like for such a small volume, this is a fairly excessive dry time, but I'm not sure how it will affect the Ampure Beads.
I go for as short as possible that will allow me to feel confident it's dry. If I'm working with only a couple of tubes, I will suck out drops with a pipette to make it faster, haha.
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Old 12-07-2011, 09:38 AM   #27
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In my case the drying time is making only resuspension time a little bit longer and usually to be really sure that it's dry I dry beads 30 min under vacuum and nothing is happening to the product. But I have to note that mostly I'm working with the PCR products so maybe there are more stable.
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Old 12-07-2011, 09:59 AM   #28
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In my case the drying time is making only resuspension time a little bit longer and usually to be really sure that it's dry I dry beads 30 min under vacuum and nothing is happening to the product. But I have to note that mostly I'm working with the PCR products so maybe there are more stable.
If it works then great, no worries. A lot of people state that if they dry to the point of cracking then they lose yield, and I think the official protocol states this now as well. But if it works, no need to change.
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Old 12-20-2011, 09:33 AM   #29
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Hi,
This question is related to AMPure XP but comparing it with the RNAClean XP. In theory the have different features:

http://www.beckmangenomics.com/produ...aclean_xp.html
http://www.beckmangenomics.com/produ...ampure_xp.html

but some people told me that they are the same but with different buffer. Any idea or experience on this issue.
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Old 12-26-2011, 02:50 PM   #30
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Has anyone tried this out yet or had any thoughts? I'm getting ready to start using Illumina's Trueseq kit as well, and this struck me as a little odd too.
I use 80% ethanol, and also, I have let my beads dry to the point of cracking (15 minutes) and have seen no ill results from that. I always have gobs of library for sequencing. However, it's good to know that I can also let it dry for a shorter amount of time. Now all those Ampure XP cleanups will go a lot more quickly.
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Old 12-26-2011, 06:53 PM   #31
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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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Old 12-27-2011, 06:34 AM   #32
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Default re-using AMPure beads

Quote:
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.

http://www.ncbi.nlm.nih.gov/pubmed/21205303

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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Old 08-28-2012, 01:32 AM   #33
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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 m 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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Old 12-13-2012, 10:04 AM   #34
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Quote:
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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Old 12-14-2012, 06:32 AM   #35
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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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Old 12-27-2012, 05:44 PM   #36
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Default 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 at 06:26 PM.
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Old 01-08-2013, 05:38 AM   #37
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Default 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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Old 01-08-2013, 05:45 AM   #38
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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.
1L of AmpureXP should contain enough beads to bind a few g's of nucleic acid.
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Old 01-08-2013, 07:16 AM   #39
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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?
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Old 01-08-2013, 04:49 PM   #40
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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.
1L of AmpureXP should contain enough beads to bind a few g'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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