Ampure versus Ampure XP

[seqgirl123]

Does anyone use Ampure (aka SPRI) in their library prep to replace the other types of PCR purification methods? Ampure XP is a newer version of the existing Ampure, supposed to be faster cleanup.

I cannot find a good text description that will answer these questions. Does anyone know?

1) what is the Ampure buffer made of ?
2) what are the beads made up of? are they of a polymer substance?
3) how does the DNA bind to the beads and then to the magnet? (all I know is that probably the positive charge on the beads bind the negatively charged DNA, and this complex then binds to the magnet, but what is the actual chemistry here, such as carbon or phosphate bonds of the DNA involved or how the makeup of the beads react to the DNA?)
4) are the 70% ethanol washing steps to remove any contaminants that are bound to the beads?
5) how does dna elute off in Tris while the beads are still bound to the magnet, again want to know the chemistry behind it?
6) how does XP performs faster cleanup versus Ampure, meaning what is the difference in the binding chemistry? Are the beads bigger in XP versus regular Ampure, or does bead size matter at all?

[ECO]

Great questions. Many of these questions are answered in the original patents (the main one is attached...I really like http://patentstorm.us for a free source of patents if you need more).

PEG and some combination of high salts (based on the patent...most likely ~10-15% PEG6-8k, + 1-1.5M NaCl + some divalents) precipitates nucleic acids (and some other molecules)...directly onto the bead surface (which is carboxylated).

It's a very old technique to PEG precipitate via traditional centrifugation, the beads just allow you to avoid spinning by acting as nucleation sites for the precipitation.

As far as the exact chemical differences between standard and XP, I don't know.

[seqgirl123]

thanks, the info helps a lot.

If anyone knows of any info about XP it would be helpful, such as if bead size or surface area of binding is larger than that of regular Ampure, or how binding chemistry is faster now?

[greigite]

I don't know how the chemistry differs, but a rep told me the main difference between the kits is a longer shelf life for XP (1 yr versus 6 mo for regular).

Originally Posted by seqgirl123

thanks, the info helps a lot.

If anyone knows of any info about XP it would be helpful, such as if bead size or surface area of binding is larger than that of regular Ampure, or how binding chemistry is faster now?

[nextgen]

Ampure is just more expensive. Some of the time saved is just from reducing drying time. I think PCRClean is better in terms of band tightness.

Originally Posted by seqgirl123

thanks, the info helps a lot.

If anyone knows of any info about XP it would be helpful, such as if bead size or surface area of binding is larger than that of regular Ampure, or how binding chemistry is faster now?

[ECO]

Originally Posted by greigite

I don't know how the chemistry differs, but a rep told me the main difference between the kits is a longer shelf life for XP (1 yr versus 6 mo for regular).

Azide in XP.

[H5N1]

Originally Posted by seqgirl123

1) what is the Ampure buffer made of ?

AMPure buffer: 1.25 M NaCl, 7% PEG8000. See the old file from Roche. I don't know if the AMPure XP buffer is exactly same.

[peterwang]

Originally Posted by seqgirl123

2) what are the beads made up of? are they of a polymer substance? ....
6) how does XP performs faster cleanup versus Ampure, meaning what is the difference in the binding chemistry? Are the beads bigger in XP versus regular Ampure, or does bead size matter at all?

I found some answers in a Powerpoint/PDF on the Beckman-Coulter AMPure website (attached, see pages 5 & 6).

AMPure XP beads have two magnetite layers vs. only one layer in AMPure beads; so they may be bigger. In AMPure beads, the magnetite is sealed by a "non-styrene polymer"; it is unclear from the figure whether XP beads are sealed with something else or the same. For both, the surface is covered with carboxylic acid groups.

The increased magnetite content in XP beads makes them bind faster to magnets. However, the major speed-up is because the XP protocol eliminates the 10 min. air-drying step after 70% EtOH wash that is in the original protocol.

[Hyped]

Anybody know how big the Agencourt Ampure XP magnetic beads are?

[squeak]

How is the drying time supposed to be reduced so much? I still do a 5 minute as I think it's necessary

[der_eiskern]

Anyone know about the ampure XP binding capacity for the beads? other magnetic bead systems claim 2 micrograms per 100 micrograms of beads.

Thanks,
Der.

[janejane]

Originally Posted by ECO

Azide in XP.

wouldn't azide interfere with library preparation?

[peterwang]

@ squeak:
I agree with you, I still let XP beads dry completely; not sure it's necessary, but it seems sensible to evaporate off the last traces of EtOH.

@ der eiskern:
The capacity of beads is probably quite high, since it is really a PEG precipitation and the beads just provide a surface to accumulate on (see ECO's post above).
There is a paper giving a "home-made" SPRI protocol using Dynal(Invitrogen) MyOne-carboxylic acid beads which states that 10 ul beads was sufficient for 0.5 ug DNA.

I haven't tried their protocol, but it is intended for construction of high-throughput deep-sequencing libraries.
Lundin S, Stranneheim H, Pettersson E, Klevebring D, Lundeberg J. "Increased throughput by parallelization of library preparation for massive sequencing." PLoS One. 5:e10029 (2010).
http://www.plosone.org/article/info%...l.pone.0010029

@ janejane:
any azide in the buffer would be washed away before the DNA is eluted.

[sjcire]

Thanks everyone for the information you posted here. I use the ampute XP product and have a need to purify a large volume of a low concentration DNA solution. I cannot use one ml of Ampure XP for every purification...again. One concern however it the low end cut off. Apparently this quality of the beads can be modified by the concentration of the crowding reagent. It is important that I know that I recover at least 150 bp and above. To this end I ran the following experiment. Using the stock reagents found in the attached document I created three PEG solutions. Each solution was made so that when 90ul was added to 50ul of sample it would have a NaCl concentration of 1.25M, MgCl2 concentration of 10mM and either 7, 10, or 13% PEG 8000. I followed my own Ampure XP procedure(also found in the attached document) with the following modification. Three 90ul alloquotes of Ampure XP were collected by magnet and washed twice with either 90ul of 7, 10, 0r 13% PEG solution. 90ul of each solution was added to the washed beads. 25ul Hi-Lo DNA marker(bionexus) and 25ul of dH2O was added to the three washed bead suspensions and 90ul of Ampure XP (not washed). After beads had been washed with EtOH and dried, 25ul of dH2O was used to elute. 5ul of 40% sucrose was added to each sample and then 10ul of each sample was loaded into 4-20% TBE acrylamide gel, unpurified Hi-Lo was also loaded. Gel was run at 300v in 0.5 TBE until BPB ran off gel. Gel was stained with ethidium bromide and destained in dH2O. Gel was visualized with an alpha innotech.

Results:

WTF, the ampure treated samples did not run correctly. LOL, what a waist of time. It may have been the sucrose, or the excessive heat used or maybe quick temperature change. Any ideas? I have noted that sucrose loaded samples run differently from glycerol samples. Perhaps Hi-Lo was not meant to be run with sucrose.

Samples are as follows:
Ladder:Bionexus Hi-Lo - lowest band is 50bp. (Most people can get the rest from there. If you cant google bionexus hi-lo)
1. Ampure XP
2. 7% PEG
3. 10% PEG
4. 13% PEG
It is clear from the gel that all preformed similarly. However note that low molecular weight bands (whatever they are) are faint in the 7% sample. If that band corresponds to 50bp than it would appear that 7% was less able to precipitate low molecular weight oligomers. 10% and 13% are more or less indistinguishable from AmpureXP. It remarkable that during the experiment each sample collected the magnetic beads at different rates. This may have been due to the viscosity of the solution they were suspended. 7% cleared beads quickest followed by Ampure XP then 10% then 13%. This is in agreement with the gel.

Conclusion: Quit f**king up gels. My god I've been running DNA for almost a decade and I'm still screwing it up. F my life. Also, use 10% PEG with 1.25M NaCl and 10mM MgCl2 for similar results to Ampure XP. 7% works and could be used to size select DNA.


I may rerun this experiment. Any pointers? Has anyone ever seen a ladder run like this? I'm interested in eliminating DNA up to 150bp and it would seem that lowing PEG concentration to 7 or lower may accomplish this.

[sjcire]

Above experiment was repeated with the following changes. Sample was dried in a vacufuge for 5 min. Samples were eluted at room temperature. Sample key is the same. Additional sample was loaded, Hi-Lo with sucrose.

It appears that the weird looking gel was a result of the heating steps, either drying or eluting. This gel ran normally. It seems obvious that less PEG does effect the recovery of short nucleotides. 7% PEG recovered only ~25% 50bp nucleotide, ~50% 100bp. Recovery was nearly 100% at 200bp and above for all concentrations of PEG. It would appear, as I stated above, that Ampure XP acts as we would expect if the PEG concentration was between 7 and 10 %. Furthermore, it is obvious that the size exclusion characteristic is not an abrupt cutoff but a broad slope. Clearly, further research is needed...robust, dynamic, synergy. Oh wait this is not a paper.

[Susanne]

Does anybody know what the composition of the elution buffer is? Shouldn't that be almost water?

[ECO]

Water will work. Typically I use 10mM TrisHCl pH 8.5.

[Erika Feltrin]

Anybody know how big the Agencourt Ampure XP magnetic beads are?

[edawad]

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

[katsigner]

Can the beads be reused?? Any idea?