Primer dimer problem

[Chipper]

Originally Posted by molly burns

I would like to understand more about double SPRI method to remove small fragments or adapters. I read that if I use 1.8 V of Ampure beads, only large DNA fragments will bind to the beads. On the other hand, if I go sequentially, that is use 0.8 V beads, it will only bind small fragments and large fragments will stay in sup. I can then recover the large fragments by adding another 1 V of beads. How is it possible then to remove small fragments by adding 1.8 V of beads? How does then small fragments stays in solu with 1.8 V of beads?

1.8 V will bind all fragments > 100 bp, start with a lower volume (eg 0.4) to bind large fragments and then add more beads to the sup to capture smaller fragments. Dial in the ratio with a 50 bp ladder or sonicated DNA.

[captainentropy]

take a look at the image posted by pbluescript earlier in this thread. 1.8 volumes isn't represented but I think the point is clear.
http://seqanswers.com/forums/showpos...8&postcount=16

[odile]

I need to remove fragments > 500bp and already removed everything <100bp (I basically have a smear that starts around 100-150bp). I would like to try to use 0.7x (from bluescript gel) and keep the supernatant.
What would you do next? Clean-up the supernatant with Qiaquick or MinElute or add 1.8x beads and start all over again?

[Heisman]

Originally Posted by odile

I need to remove fragments > 500bp and already removed everything <100bp (I basically have a smear that starts around 100-150bp). I would like to try to use 0.7x (from bluescript gel) and keep the supernatant.
What would you do next? Clean-up the supernatant with Qiaquick or MinElute or add 1.8x beads and start all over again?

You can add 1.8x beads and it will work. The idea is that you need to get the PEG concentration above whatever the threshold is for the size you care about. You could actually get away with less than 1.8x beads since you already have 0.7x bead solution, but it won't hurt to add more.

[DavidAntonio]

Originally Posted by pbluescript

I have attached an image showing the effect of altering the ratio of AMPure beads/sample. You can see that using a lower ratio will remove increasing sizes of DNA fragments. You can even use this for size selection by using two different concentrations. For example, if you wanted to get a band centered ~400 bp, you could use 0.7X beads and then take the supernatant. This would include all the DNA fragments that were smaller than ~500bp and didn't bind to the beads. Then mix the supernatant with 0.9X beads and everything larger than ~300bp would bind. Elute the bound DNA and you'll have a fairly broad, but good enough to sequence, library.
If you try the beads, there can be a bit of lot-to-lot variability, so I would test them with a 100 bp library as I did in this gel.

I haven't used the Truseq kit, but I have used the Truseq adapters with outsourced enzymes/buffers and the same problem can occur.

I am sorry about the Necro Post, but I have a problem in this moment about this. Could you tell me the weigth (in bp) of your 1 kb ladder? It would be very useful for my thesis. Thanks a lot.

[pbluescript]

Originally Posted by DavidAntonio

I am sorry about the Necro Post, but I have a problem in this moment about this. Could you tell me the weigth (in bp) of your 1 kb ladder? It would be very useful for my thesis. Thanks a lot.

The 100 bp ladder in the image is this one:
http://products.invitrogen.com/ivgn/product/15628019
The 1kb+ ladder in the image is this one:
http://products.invitrogen.com/ivgn/product/10787026

[rnaeye]

pbluescript,
what is the DNA ladder you used on this gel. thank you!

[captainentropy]

Originally Posted by rnaeye

pbluescript,
what is the DNA ladder you used on this gel. thank you!

He's using both. The 100 bp ladder was what he was size-selecting (to illustrate the use of the SPRI bead ratios), the 1kb+ ladder is the size marker for the gel.

Take a look again at the text above each lane.

[yuehuang]

Originally Posted by Simone78

I had the same problem(s). To get rid of the adaptor-adaptor dimers I now do size selection using 2% E-gel (great because you can also select more than one single fraction, so you have a backup in case something goes wrong during the pre-amplification step). And you don't need an additional purification step, just pipette out the water (containing your DNA) from the well. To remove the primer dimers I use AMPure XP (1.8:1 ratio) which removes (almost) 100% of the fragments <100 bp.

Thanks much for the tips. I was always bothered by the primer dimer when used NEB kits for library prep. The AMPure XP worked fabulously for size selection, however didn't work for getting rid of the primer dimer. I had to use Pippin Prep to clean-up the products, which caused more than half loss. After switched to Nextera kit, no more primer dimer problem, but primer contamination presented sometimes. The manufacturer's suggestion is only 25uL beads for 50uL products. I would try your 1.8:1 ratio next time.

[yuehuang]

Originally Posted by pbluescript

I have attached an image showing the effect of altering the ratio of AMPure beads/sample. You can see that using a lower ratio will remove increasing sizes of DNA fragments. You can even use this for size selection by using two different concentrations. For example, if you wanted to get a band centered ~400 bp, you could use 0.7X beads and then take the supernatant. This would include all the DNA fragments that were smaller than ~500bp and didn't bind to the beads. Then mix the supernatant with 0.9X beads and everything larger than ~300bp would bind. Elute the bound DNA and you'll have a fairly broad, but good enough to sequence, library.
If you try the beads, there can be a bit of lot-to-lot variability, so I would test them with a 100 bp library as I did in this gel.

I haven't used the Truseq kit, but I have used the Truseq adapters with outsourced enzymes/buffers and the same problem can occur.

Great job! I played with this too. But my concern focused on the size adjusted concentration, your gel is very convincible.

[yuehuang]

Originally Posted by monad

2nd gel purification is the best option here, but you lose lots of library DNA. In this case, you can set additional round of PCR will take care of it. Stick with not more than 5 round of PCR. If your bioanalyer peak is good, you will be fine even with 10-15 cycles of PCR.

Did you mean, the already purified DNA library to go another PCR at about 5 cycles, checked by bioanalyzer then go amplification again in 10-15 cycles to accumulate enough yield for sequencing?

[yuehuang]

Originally Posted by pmiguel

I have a partial answer. We tried a few modifications to the TruSeq protocol, one of which was to do the PCR step on non-size selected library, followed by size selection on the Pippin Prep. Our yield of final library was much lower than what we got following the normal protocol. But it was still enough to sequence.

It worked.

But there are a couple of additional issues for a non-amplified library:

(1) The Y-adapters apparently cause library fragments to migrate aberrantly on some, but not all, electrophoretic conditions. Details are in another thread. I think they may run at their correct size on e-gels, but appear larger than they actually are under most commonly used electrophoretic conditions. After enrichment PCR the adapters are no longer Y-ed, so the effect disappears and you see the amplicon's true sizes.

(2) Just to state the obvious: when you work with very limiting amounts of DNA, you have to watch losses to binding against plastic-ware, etc. A typical microfuge tube may be able to bind 1 ng of DNA (complete guess.) So, if you have 200 ng, who cares. But if you have 1 ng total, you might lose the majority of your library if you don't use low-bind plastic-ware.

--
Phillip

thank you so much for the low-bind tube tip. I tried the same procedure, got the perfect size selected and clean library prep products, but much lower yield. Lucky to me, it's still sequencable, but very scary.

[XIAOXIAO]

Hey if you used AmPure beads for purification, what is the ratio of library versus beads for purificaiton?

[Simone78]

Originally Posted by XIAOXIAO

Hey if you used AmPure beads for purification, what is the ratio of library versus beads for purificaiton?

DEpending on which fragments you want to retain and which ones you would like to get rid of. To remove short fragments (dimers/primers) use a 0.8-1:1 ratio beadsNA. If you want to pull down everything use a 1.8:1. If you want a specific size range, do a double (two-sided) purification where in the first your DNA is in the supernatant and in the second is on the beads, or vice versa.
Best,
Simone

[XIAOXIAO]

thanks! i will try 1:1

[liuqy]

Originally Posted by yuehuang

Thanks much for the tips. I was always bothered by the primer dimer when used NEB kits for library prep. The AMPure XP worked fabulously for size selection, however didn't work for getting rid of the primer dimer. I had to use Pippin Prep to clean-up the products, which caused more than half loss. After switched to Nextera kit, no more primer dimer problem, but primer contamination presented sometimes. The manufacturer's suggestion is only 25uL beads for 50uL products. I would try your 1.8:1 ratio next time.

You may try NVIGEN MagVigen. It's observed that AmPure XP lags MagVigen in primer dimer/adapter dimer cleanup. You can check these different contamination levels (>200%, >100%, >50%, 20%) (https://www.nvigen.com/dna-clean-up/)

Disclosure: I work at NVIGEN. We cordially invite you to try MagVigen. In case you find problems using it, we will be happily work with you to troubleshoot or fine tune it to meet your needs.

[liuqy]

Originally Posted by XIAOXIAO

Hey if you used AmPure beads for purification, what is the ratio of library versus beads for purificaiton?

There is a charter picture here, which demonstrates beads volume vs sample volume (note: not sample to beads). NVIGEN MagVigen uses the same protocols as AmPure XP.
https://www.nvigen.com/dna-size-sele...utorial/#ratio