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ddRAD protocol by Peterson et al. Hanne Sample Prep / Library Generation 1 11-22-2013 06:41 AM

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Old 03-02-2013, 06:09 AM   #1
blair.chr
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Default ddRAD-Seq

Hi all,

I am following the ddRAD-Seq protocol of Peterson et al. 2012. I start with ~40-100 ng/ul DNA as measured on a nanodrop. I take 5 ul of this and double digest with two different enzymes. I then clean this entire product with Ampure XP beads.

The problem is that I am losing A LOT of DNA at every step. If I run the uncleaned digests on a bioanalyzer I get about 3-9 ng/ul. After Ampure cleanup my concentrations decrease to less than 1 ng/ul. As the standard protocols for restriction digests and bead cleanup are fairly straightforward, I am not sure where the problem lies. Any thoughts would be appreciated.

Chris
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Old 03-04-2013, 07:38 AM   #2
ZWB
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Hi Chris,

We've had similar problems with determining concentrations. The problem may actually be the nanodrop rather than your protocol. We switched to a Qubit and get much more reliable results. We found that any impurities in your sample really throw off the nanodrop, but using a fluorescence based dye as with the Qubit fixes that issue. Good luck.
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Old 08-26-2013, 03:16 PM   #3
srdoyle
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Hi Chris,

Just wanted to check in and see how your ddRAD experience is going... we might be having similar issues and am keep to see how you dealt with them. Happy to chat via email too if you have some time.

Cheers,
Steve
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Old 11-22-2013, 06:37 AM   #4
horbachc
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What platform are you running your ddRAD on? I am using MiSeq, and have had issues with low cluster density compared to other library prep protocols (Nextera XT) we have used. What concentration are you loading?
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Old 05-21-2014, 12:27 PM   #5
maxbangs
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Hello blair.ch, I do not know if you have solved your problem or not, I know this response is very late. I have just started with the Peterson protocol and may have a little help. Please note I am a complete novice (started two weeks ago) and may be way off base.

First off if you loaded 5ul of 40-100ng/ul DNA into a standard 50ul reaction then your concentration should be 1/10th of what you started with, so 4-10ng/ul. This would explain why you got 3-9ng/ul concentrations for your digest.

Second with the AMPure cleanup only DNA fragments of >100bps will attach to the beads, thus if most of you digest is <100bps then this may explain why the DNA disappeared after the clean up. I had this problem with a couple of samples. These samples looked great on the Qubit (>1000ng/ul of DNA) but when I looked at the quality of the DNA on a gel the sample was highly degraded and lacked any intact genome DNA. Thus when I looked at the digest on the gel all of the DNA fragments were <100bps.

Hope this helps, also I am curious if you figured out your problem and had any advice for us just starting.
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Old 06-05-2014, 02:55 PM   #6
rosatoc
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Hi. I have just tried my first double digest (PstI/MspI) for library construction, but using the GBS method rather than the RAD-Seq. On wheat. The Bioanalyzer trace has a huge adapter dimer and a bit of other 'stuff' which is too close to baseline to interpret. With those disappointing results we ran a gel following a digest to see whether the sample(s) are restrictible, and though we see a smear there is evidence of a lot of high MW DNA which may not have been cut. Are there any suggestions on what to try next to get this method to work? We're using columns for clean-up rather than beads, and we may switch though I don't think that would make a difference here.

Many thanks for any tips, tricks, etc. you are able to share
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Old 06-05-2014, 03:32 PM   #7
nucacidhunter
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I suspect you have followed Poland et al protocol. I may be able to provided some suggestions. Could you post your Bioanalyser trace (is it library after PCR clean up?) and your digest gel photo with a bit of details (DNA input, duration of digestion and loaded amount). It is usual not to see much of smaller fragments. If column is used for clean up, a titration with different amount of adapters is essential, but one can save a lot of time just by doing a 1.2x bead clean up which will remove excess adapters and relatively short fragments which does not provided useful data.
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Old 06-05-2014, 05:44 PM   #8
maxbangs
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I am also using Pst I x MspI on some fish with a genome size of ~2.42Gbps. I attached a gel picture of a successful digest, the ladder is 1kb, 750bp, 500bp, 300bp, 100bp, 50bp. I used 10units for each RE and 1000ng of DNA in a 50ul reaction ran over night (17hours). Though this may be helpful to the discussion since it sounds like you have a problem with the digest.
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File Type: png Gel Doc 2014-05-23.png (124.8 KB, 65 views)

Last edited by maxbangs; 06-05-2014 at 05:46 PM.
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Old 06-05-2014, 08:43 PM   #9
SNPsaurus
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One issue with wheat is that PstI will very rarely cut since most of the genome is methylated, but MspI will cut the methylated DNA. I don't know how you calculated the adapter amounts to add, but the PstI adapter will only find overhangs at a very low rate compared to what is expected from the genome size.
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Old 06-06-2014, 07:17 AM   #10
rosatoc
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Thanks for all responses. As we are trying this using the Poland et al protocol, developed for wheat (and barley), we used the adapter concentrations according to the protocol without a titration. That may be something we will need to consider. We have a few ideas about how to do the next test library construction and we feel success is right around the corner.... Hope springs eternal.

Thanks, again, for all replies
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Old 06-19-2014, 02:01 PM   #11
rosatoc
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Hi - we are still trying to get the double digest to work in our lab. We know that the digest is working (gel imaging) but the ligation results in adapter dimer peak only post PCR. There is a heat inactivation step following the digest with PstI-HF/MspI, however neither of these enzymes are heat inactivated.... Is there any concern that the active enzymes will prevent ligation to occur?
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