AMPure adapter dimer carryover THROUGH CAPTURE

[ojm101]

Hi All,

I have been having some trouble with 120 bp adapter dimers showing up after ligation during Illumina library prep. We have a homebrew system here, and previously we incorporated SPRIselect (distinct from AMPure XP) but we are phasing that out because of the low yield issues and subsequent bias. As soon as we took out SPRIselect, we started seeing adapter dimers all over the place, which prompted us to use a lower (<1.8x) ratio of AMPure beads to "cut" the dimers out.

And that is where the confusion started. I used 100 bp ladder and 0.9x and 1.2x ratios of ampure beads and basically got no cutoff whatsoever. I could still see the 100 bp band in all samples. I am confident I used the correct volumes of beads and that there was no evaporation/discrepancy regarding my ladder samples. The ladder was in 50 uL and the beads were 45 and 60 uL respectively.

What else could be going on here? It's occurred to me that the lot of these beads might have some slight variation but not such that I would still be seeing 100 bp fragments with a 0.9x ratio. I have seen a lot of people using reduced binding ratios and getting results that I have thus far been unable to reproduce. Based on these results it's no wonder that adapter dimers are making it through in my libraries.

What's even crazier is that these dimers are somehow finding their way into our probe captured libraries! Has this happened to anyone before?

Let's please assume that all pipetting was done accurately and there is no adapter contamination anywhere in the lab. We have considered this and taken every measure possible to avoid it and I do not want this thread to focus on contamination problems unless you feel like you have something extremely profound to share.

Thanks in advance

[pmiguel]

We calibrate our batches of AMPure and generally see a 50-80% decrease in the amount of a 100 bp band with 0.9X volumes of AMPure. We always do a series of calibrations down to 0.5X volumes. You should probably do the same for your home brew.

Keep in mind it isn't the beads themselves that effect the size cut, but the PEG concentration of the solution and other factors that effect PEG precipitations (eg, temperature, time). So if you are using "home brew" beads it could be your PEG is a little more concentrated than you intended.

--
Phillip

[ojm101]

Thanks very much pmiguel.

When I mentioned the homebrew system, I was really referring to the incoming reaction compositions. These tests were all with AMPure beads.

Reducing the ratio makes sense but my target fragment size is ~200 bp before ligation and 300 bp after ligation. As such, I worry about losses with a ratio as low as 0.5x.

I have seen many gel images posted on this website and elsewhere that showed distinct losses of bands of ladder with reducing the bead ratio. The fact that there is any trouble reproducing what should be very straightforward is troubling.

Does anybody have any literature or advice on HOW the factors pmiguel listed (temperature, time, etc) can affect binding? Could any of them account for what I am seeing, which appears to be binding all the way down to 100 bp regardless of ratio?

[pmiguel]

Did you see no reduction in the amount of 100bp band compared to the 300bp band with a 0.9X AMPure? There should have been some, although it will not have been a complete removal of the band.

--
Phillip

[ojm101]

I've attached the image of my gel. The first well is just ladder. Afterwards, there are 4 samples cleaned up with ampure beads and 2 samples cleaned up with a competing brand.

The order of the ratios is 0.9x, 1.2x, 0.9x, 1.2x, 0.9x, 1.2x

[Hoban]

What PEG concentration are you using and what NaCl concentration are you using?

[ojm101]

The concentrations are whatever Beckman Coulter AMPureXP beads are. I'm not aware of the specific numbers. Same story with the competitor.

https://www.beckmancoulter.com/wsrpo...n%2Findex.htm/

[nucacidhunter]

Originally Posted by ojm101

Hi All,

I have been having some trouble with 120 bp adapter dimers showing up after ligation during Illumina library prep. We have a homebrew system here, and previously we incorporated SPRIselect (distinct from AMPure XP) but we are phasing that out because of the low yield issues and subsequent bias. As soon as we took out SPRIselect, we started seeing adapter dimers all over the place, which prompted us to use a lower (<1.8x) ratio of AMPure beads to "cut" the dimers out.

I wonder if you could post Electropherogram of a ligation reaction before and after bead clean up as well as the library after PCR (before any clean up) for the same sample. 0.9x bead ratio should clean 100 bp fragments.

[pmiguel]

Under the conditions your did those clean-ups it looks like you need to go to lower Ampure amounts. Try 0.6x and 0.8x, to get an idea.
Here are some factors that may impact precipitation:
-Time precipitated
-Salt, concentration. Divalent cations are said to be much more effective at preciptitation (eg Mg++) The salt concentration is cranked sky high in PEG ppts, so it may not be a factor.
-Temperature during precipitation.

Since you are probably controlling those factors pretty well, it is probably either a weird batch of ampure--possibly with higher PEG than it should have or maybe it went through a freeze-thaw that damaged the beads.
Alternatively, maybe the bead pellets aren't getting washed sufficiently? There is quite a bit of solvent volume in the pellet initially. That will contain lots of the lowest molecular weight band. If it doesn't get fully mix and diluted out, it might explain why you are retaining so much of the 100bp band.

But really I would suggest just doing a series of Ampures at 0.1x intervals from 0.5x to 1.0x on your 100 bp ladder to find where you get highest 300bp/100bp recovery ratio.

Also, if necessary, you could do two Ampures in a row. This might have more impact than you are expecting. Say 0.9x is giving 90% recovery of 300bp and 30% recovery of 100bp. After a 2nd ampure you expect about 81% (.9^2) recovery of 300bp and 9% (.3^2) of 100bp.

--
Phillip

[Hoban]

Originally Posted by ojm101

The concentrations are whatever Beckman Coulter AMPureXP beads are. I'm not aware of the specific numbers. Same story with the competitor.

https://www.beckmancoulter.com/wsrpo...n%2Findex.htm/

Have you tried using lower concentrations (0.5x-1x) yet? I'm interested to see the result of that.

[NextGenSeq]

Are your adapters homebrew also?

The Illumina adapters have modifications at the 3' end which increases ligation efficiency
(see the sticky)

[pmiguel]

Originally Posted by NextGenSeq

Are your adapters homebrew also?

The Illumina adapters have modifications at the 3' end which increases ligation efficiency
(see the sticky)

Eh? What "sticky"?
What do you mean "increases ligation efficiency"?

--
Phillip

[NextGenSeq]

Under the Illumina forum
http://seqanswers.com/forums/showthread.php?t=198

People use either phosphothioate or LNA between the last two 3' bases. If you don't do this you will get much more adapter artifacts.

We've found that the Bioo adapters give fewer adapter artifacts than even purchased Illumina adapters.

[pmiguel]

Originally Posted by NextGenSeq

Are your adapters homebrew also?

The Illumina adapters have modifications at the 3' end which increases ligation efficiency
(see the sticky)

Originally Posted by NextGenSeq

Under the Illumina forum
http://seqanswers.com/forums/showthread.php?t=198

People use either phosphothioate or LNA between the last two 3' bases. If you don't do this you will get much more adapter artifacts.

We've found that the Bioo adapters give fewer adapter artifacts than even purchased Illumina adapters.

Okay, but phosphorothiate linkages are said to protect the linkage against some types of exonuclease activity not "increase the ligation efficiency". I gather LNA linkages stabilize correct base stacking interactions -- but would a single base overhang with an LNA linkage really result in a more efficient ligation?

Interesting about the bioo adapters -- to what do you attribute their giving you fewer artifacts? (Do they use LNAs?)

--
Phillip

[nucacidhunter]

I have not seen any reference, but anecdotally I have been told that through unknown mechanism phosphorothioate linkage increases ligation efficiency with increasing number of linkages. That is the reason that 454 adapters had 5 phosphorothioate linkages not just one which is enough for protection against 3’ to 5’ exonuclease activity of enzymes.

[luc]

It seems unlikely that LNA bases would be used - to my knowledge, LNA in the template inhibits most polymerases.
I assume the phosphorothioate linkage helps predominantly preventing 3’ to 5’ exonuclease activity of the ligase - reducing adapter dimers.
We too, had good experiences with the Bioo adapters though.

[NextGenSeq]

Reduction of non-insert sequence reads by dimer eliminator LNA oligonucleotide for small RNA deep sequencing

Here we describe a method for constructing small RNA libraries for high throughput sequencing in which we have made a significant improvement to commonly available standard protocols. We added a locked nucleic acid (LNA) oligonucleotide—named dimer eliminator—that is complementary to the adapter-dimer ligation products during the reverse transcription reaction. It reduces adapter-dimers, which often contaminate standard libraries and increase the number of non-insert sequence reads.

http://www.biotechniques.com/Biotech...es-304403.html

Tufts LNA Adapter Protocol
http://www.tucf.org/htseq_protocol_f...ina_paired.pdf

[M4TTN]

Dear NextGenSeq, (apologies for dragging up an old post!)

I have a DNA enrichment project where our inserts range from only 40bp and up. We'd like to get them all.

We've just done a run and it is clear that we have sequenced a lot of adapter dimers (we expected that to be the case from the bioanalyser trace). My question is how applicable your strategy for dimer removal would be to standard Illumina DNA-DNA adapter dimer junctions? From the % base trace on MiSeq reporter it appears that most of the adapter junctions have gained an A (complementary to the overhanging T).

i.e. the pure adapter dimer junction should be: GCTCTTCCGATC/GATCGGAAGAGC
But we see (I think): GCTCTTCCGATCtaGATCGGAAGAGC

Or rather: in the sequence trace we see: AGATCGGAAGAGC...

And since the read primer ends: ...GCTCTTCCGATCT

This presumably means that the overhanging T is present (for the read primer to work) and an A has been added upstream of the GATCGGAA...

So...questions:
Would an oligo that spans this junction inhibit PCR?
What is the role of the LNA within this oligo?

[nucacidhunter]

Originally Posted by M4TTN

i.e. the pure adapter dimer junction should be: GCTCTTCCGATC/GATCGGAAGAGC
But we see (I think): GCTCTTCCGATCtaGATCGGAAGAGC

Or rather: in the sequence trace we see: AGATCGGAAGAGC...

And since the read primer ends: ...GCTCTTCCGATCT

This presumably means that the overhanging T is present (for the read primer to work) and an A has been added upstream of the GATCGGAA...

So...questions:
Would an oligo that spans this junction inhibit PCR?
What is the role of the LNA within this oligo?

I think adapter-dimer formation is more complex. I would suggest analysing a TruSeq Universal and Index adapter for hetero-dimer formation for instance using OligoAnalyzer. You will see that there is more than one way for hetero-dimer formation with relatively low Delta Gs (they will anneal during annealing step) and each of those can be extended and then those products can form new versions of hetero-dimers in the following cycles.

In your target range the best way for getting rid of dimers is preventing or reducing dimer formation by careful titration of adapters or using dimer-free adapter technology.

PS. I have not seen any proof that Illumina uses LNA technology in their adapter oligos but LNA will increase the stability of annealed adapter oligos.