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greigite 01-26-2010 02:48 PM

ampure purification of illumina libraries
 
1 Attachment(s)
I'm hoping for some advice/protocol sharing on the use of Ampure beads for purification of Illumina libraries post-PCR. I ran a calibration of my beads with a titrated ladder, which suggested an 0.85:1 bead:template ratio to get rid of adapter dimers. However, following repeated tries at purification I am still not able to completely get rid of a peak at ~130-140 bp (see attached trace). In multiple library preps it usually remains at around 10-25% on a molar basis, still too high for sequencing, as I don't want to waste that much flow cell real estate on adapters. I have tried other bead:template ratios as well, including the standard 1.8:1 bead:template of the standard protocol, and gotten generally worse results. Any tips on optimizing the Ampure purification protocol would be greatly appreciated!

upenn_ngs 01-28-2010 07:23 AM

from my experience, it is necessary to to a gel sizeselect, especially if you are doing whole exome protocol. the peak at 130 is self-ligated adapter with pcr extension. invitrogen has a great precast gel that makes the sizeselect quick. run the gel after ligation and before pcr to separate.

greigite 01-28-2010 07:55 AM

Interesting to know that the 130 bp peak is self ligated adapter. I did actually do a gel size selection step following ligation, but it seems that was insufficient to actually eliminate self-ligated adapter. I guess that migration through the gel was imperfect such that there were still some adapters "trapped" with the higher MW DNA, and that they preferentially amplified in the PCR.


Quote:

Originally Posted by upenn_ngs (Post 13106)
from my experience, it is necessary to to a gel sizeselect, especially if you are doing whole exome protocol. the peak at 130 is self-ligated adapter with pcr extension. invitrogen has a great precast gel that makes the sizeselect quick. run the gel after ligation and before pcr to separate.


nextgen 02-14-2010 07:58 AM

The reason is that you have a lot DNA at 130. Ampure size selection is not as tight. When you have a lot DNA in the vicinity of your target size, it will just carry over. A double selection or tighter selection will help. The problem with gel is that when you load a lot, DNA gets tangled together. The target DNA has mix from non-target DNA. That is why Saga bio tries to limit the DNA input for their automated robot.

Quote:

Originally Posted by greigite (Post 12986)
I'm hoping for some advice/protocol sharing on the use of Ampure beads for purification of Illumina libraries post-PCR. I ran a calibration of my beads with a titrated ladder, which suggested an 0.85:1 bead:template ratio to get rid of adapter dimers. However, following repeated tries at purification I am still not able to completely get rid of a peak at ~130-140 bp (see attached trace). In multiple library preps it usually remains at around 10-25% on a molar basis, still too high for sequencing, as I don't want to waste that much flow cell real estate on adapters. I have tried other bead:template ratios as well, including the standard 1.8:1 bead:template of the standard protocol, and gotten generally worse results. Any tips on optimizing the Ampure purification protocol would be greatly appreciated!


greigite 02-15-2010 12:39 PM

Quote:

Originally Posted by nextgen (Post 14010)
The reason is that you have a lot DNA at 130. Ampure size selection is not as tight. When you have a lot DNA in the vicinity of your target size, it will just carry over. A double selection or tighter selection will help. The problem with gel is that when you load a lot, DNA gets tangled together. The target DNA has mix from non-target DNA. That is why Saga bio tries to limit the DNA input for their automated robot.

Hi nextgen,
Could you provide more info or a link about the robot you mention? Also, by double selection do you mean a post-PCR gel size selection in addition to the post-ligation gel size selection?

nextgen 02-16-2010 04:48 PM

Check this out:
http://www.genomeweb.com/sage-scienc...gen-sequencing.

Double selection means you do the same selection twice. You will lose more DNA with each selection. Ampure selection ability is limited, especially towards 500bp and above. So there is a trade off between yield, DNA size tightness and yield.

Quote:

Originally Posted by greigite (Post 14060)
Hi nextgen,
Could you provide more info or a link about the robot you mention? Also, by double selection do you mean a post-PCR gel size selection in addition to the post-ligation gel size selection?


gogreen 03-10-2010 06:24 AM

1 Attachment(s)
Hi Greigite,

I see a similar band at 130 bp in my sequencing library too! Did you figure out why this happens and any way to get rid of this? I tried going down on the adaptor concentration. But it was no good. The first lane is the library prepared with the solexa adaptors and the 2nd one is a multiplex adaptor that we made. I wonder why this doesnt happen with the solexa adaptors used at a higher concentration than the multiplex!! any suggestions would be great, Thanks in advance :)

greigite 03-10-2010 07:21 AM

Quote:

Originally Posted by gogreen (Post 15222)
Hi Greigite,

I see a similar band at 130 bp in my sequencing library too! Did you figure out why this happens and any way to get rid of this? I tried going down on the adaptor concentration. But it was no good. The first lane is the library prepared with the solexa adaptors and the 2nd one is a multiplex adaptor that we made. I wonder why this doesnt happen with the solexa adaptors used at a higher concentration than the multiplex!! any suggestions would be great, Thanks in advance :)

As upenn_ngs said above this band is most likely self-ligated adapter amplified with PCR primers. Your library looks concentrated enough that gel extraction should work well to eliminate the 130 bp peak. I've heard that there are some completely ampure-based protocols now for size selection but I have not been able to get rid of that peak without gel extraction. most likely I am doing something wrong :)

upenn_ngs 03-10-2010 09:50 AM

Quote:

Originally Posted by greigite (Post 15227)
As upenn_ngs said above this band is most likely self-ligated adapter amplified with PCR primers. Your library looks concentrated enough that gel extraction should work well to eliminate the 130 bp peak. I've heard that there are some completely ampure-based protocols now for size selection but I have not been able to get rid of that peak without gel extraction. most likely I am doing something wrong :)


we have had success eliminating excess adapters by performing two rounds of purification after the ligation. good luck!

zerone 06-17-2010 04:01 PM

Did you use two rounds of purification using Ampure beads or with gel extraction?

silin284 07-18-2010 02:16 PM

I have tried AMPure to clean up the ligation product and both 1.8 and 1.0 volume of AMPure beads work. I have never seen a 130 bp band after purification..... Is it possible that too much adapter have been used? or the homemade adapters are of poor quality therefore lots of self-ligation?

aperera 08-03-2010 08:35 AM

Has anyone tried the "Size Selector" beads from Aline Bio?

sjcire 12-09-2010 12:28 PM

One thing that may help is that Ampure XP uses PEG 8000 as a crowding reagent. If you use the quick ligation kit, the 2X quick ligation buffer has 17% PEG 6000. Therefore, Ampure XP PEG concentration ends up higher and thus the oligo selection cut off is lower. Try Ampure selection twice and dont wash with 70% EtOH but with a 6-7% PEG 8000 solution with 1.25M NaCl and some Mg2+ in there. I stepped into this same trap. DARN YOU PEG!!

nextgen 04-13-2012 10:51 AM

Sizeselector from Aline Bio
 
Quote:

Originally Posted by aperera (Post 22927)
Has anyone tried the "Size Selector" beads from Aline Bio?


You may talk to Tufts University core facility. They have some good experience. It is very effective in removing adaptor contamination.

TonyBrooks 05-11-2015 07:33 AM

Here a a few tips that I've found through experience:
1) titrate your adapter to your sample input. We use 1L of 15L for every 500ng of DNA. For making homebrew adapters we follow the protocol in Kozarewa et al. http://www.ncbi.nlm.nih.gov/pubmed/21431778.
2) If in doubt, double Ampure. If you want to save on beads, you can use a homebrew binding buffer (20% PEG8000, 2.5M NaCl). After resuspending beads in elution buffer, just add this at the required concentration, incubate, rebind beads, wash with ethanol twice, dry and elute.
3) 1:1 is a good ratio to use for most libraries. This should be good a removing anything below 200bp. We don't bother calibrating beads against a ladder. It's mostly a waste of time and reagents.
4) Try and do the double Ampure pre-PCR. PCR will preferentially amplify smaller material, so it will favour any adapter dimer over sample. Failure to do this will lead to large dimer peaks and lower yield from your library.
4) Make sure the beads are dry after the ethanol wash. Spin down the tube/plate once you remove the second wash. Place it back on the magnet and use a low volume pipette to remove any residual ethanol. Look for cracks in the bead pellet so you know it's dry. You can also speed up drying by incubating on a 37C block. If you do that, check every 30 seconds to make sure you don't over-dry the beads.

Lovro 09-08-2016 02:46 AM

Thank you for all the info and this nice summary.
I was wondering if you guys have any advice on eliminating large fragments. The step is included in TruSeq DNA PCR-Free library prep from Illumina. They dilute 100ul of SPB with 60ul water for 350bp insert and with 100ul of water for 550bp insert size and then use supernatant in the next step.
I'm trying to prepare 1000bp insert size library and am wondering if I should perform this step at all since according to gel, there aren't any larger fragments in the samples (DNA was sheared with Covaris)
Are large fragments problematic downstream in the protocol, including sequencing on MySeq?
If I will perform this step, maybe I should dillute SPB with water 1:3 and this will leave 1000-1500bp fragments in the supernatant? This is only speculation, so any advice from experience would be appreciated.

On the other hand, since I will later enrich the library with custom IDT probes, the 130bp adapter ligates probably would not be a problem since they will not be pulled ?

There is also no PCR step before enrichment, so biased replication of the shorter fragments shouldn't be a problem.

Would you recommend skiping both cleanup steps in order to preserve the sample?
Your thoughts ?

Thank you,
Lovro

nucacidhunter 09-08-2016 05:06 AM

To prep a library with fragments over 1kb, you need to shear DNA to 1kb with mini-tube and do one sided size selection using 0.45-0.48x AMPure beads. Larger fragments will not interfere with sequencing.

After ligation excessive adapters need to be cleaned up otherwise they will persist throughout the capture and might saturate blockers resulting in inefficient and off target pull down.

Lovro 09-08-2016 05:49 AM

Thank you!
Almost forgot about the blockers used in enrichment.

By one sided you mean I have to get rid only of the shorter fragments? With 0.45-0.48x AMPure beads you suggest I keep the beads or the supernatant? In the official protocol they use such bead concentration to bind large (I guess over 600bp) fragments and then use supernatant (remove large fragments).
Keeping the beads would then preserve everything above cca 600bp for further steps.
However, I am concerned how much DNA will I loose in the process ??
The reason I'm worried is because with enrichment step, most of the DNA will be discarded.

I would loose less DNA with, say, 0.7X spb. However this would preserve shorter fragments (still above 300 bp probably?) as well. Is there a reason I should be worried about this in regard to sequencing efficiency. In essence I'm interested in sequencing shorter inserts as much as I am in longer. I know the shorter fragments will cluster more, but I don't know to what extent. As long as I get 10% of 1000bp inserts it should be fine.
But I don't want to risk more sequencing errors or data loss because of uneven insert sizes.
So i guess my next question would be: "does uneven insert size hinder the sequencing process? "

Thank you!

nucacidhunter 09-08-2016 06:09 AM

You need to keep the beads and after washes elute large fragments bound to them.

The amont of DNA lost will depend on size distribution of sheared DNA. You will loose fragments below cut off and a bit above it.

To sequence 10% of 1kb fragments libray size roughly should be 800bp-1.5 kb.

I do not know what you mean by uneven because regardless of size cut off you still will have bell shaped size distribution.

Lovro 09-09-2016 10:18 AM

I was under impression that bead size selection is one sided and different concentrations of SPB bind DNA from certain length upwards. Setting the threshold at lower lengths would therefore result in wider insert size distribution.
On the other hand, if certain SPB concentration binds DNA with fragment lengths inside a given interval, the wideness of insert size distribution would remain constant.
I've read somewhere that 1:1 ratio of spb and sample binds everything of 200bp upwards. Therefore I thought the size selection is (predominantly) one sided (??is it).

Although I think that the wideness of the distribution should not be a problem sequencing-wise (there will not be more sequencing errors), I am aware of clustering being short fragment biased.

On one hand, I want to have large insert size. On the other I want to preserve sample. Too large fragment size will waste sample also because I plan to spike the library into routine runs with shorter fragments.

Anyhow, I will try with one sided size selection like you suggested, but with 0.55 AMPure beads. I'll report the results :)

tnx


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