As those in the lab will verify, I have an admitted obsession with the AMPure beads. I love to use them for any application possible. They are just so great, and the % recovery is excellent. And they are a lifesaver when it comes to eliminating adapter dimer from next gen sequencing libraries.
We have a ChIP-seq assay in development where a gel punch is required to eliminate the adapter dimers after pcr amplification, and also narrow in on the size of the library we move forward with into sequencing (our target size is around 200-500 bp). When you are processing just a couple samples, the gel punch and isolation is not such a big deal, though it does add an extra 3 hours at least, depending on number of samples. So I got to thinking, what if we used the "double-cut" concept of the AMPure beads to: First, eliminate the really big stuff (top cut), and Second, eliminate the really small stuff (bottom cut). This way I could transfer my entire ChIP-seq library prep to a plate format and greatly increase the number of samples feasibly processed at one time. I did a little experimenting, and here's a summary of what I found.
My basic AMPure protocol goes as follows:
1) Add beads at the desired ratio to the sample and pipet 10x to mix
2) Incubate 15min at room temp
3) Place on magnet 2-5min, until supernatant is clear
4) Remove supernatant (save if needed)
5) Wash beads 2x with 200-500ul 80% EtOH
6) Remove as much EtOH as possible by pipet, then let bead pellet air dry (up to 15min depending on pellet size)
7) Remove from the magnet, add desired elution solution/volume and pipet 10x to mix
8) Incubate 2min at room temp
9) Return to magnet for 2-5min
10) Remove supernatant - this contains the size-selected DNAFirst
, here is a gel showing the DNA size fragments found in the supernatant (step 4) versus what is eluted off the beads (step 10), for varying ratios of beads to sample. I used a low molecular weight ladder (100bp-1031bp) as my DNA "sample."
Interesting, I thought. You can clearly see that as the bead:sample ratio decreases, the beads selectively bind larger fragment sizes. A nice step-wise illustration where the DNA size lost in the supernatant is mirrored by the size retained on the beads. Also note the beads selectively eliminate the 100bp band no matter what other sizes are retained. This is key for the adapter-dimer removal in sequencing libraries.Second
, based on the gel above I tested different ratios for top and bottom cut, trying to zero in on a combination that could work to select out my 200-500 bp target region for the ChIP libraries.
data to come :)
We run into this all the time that people through around the word TE with know knowledge of the extensive variations that exist. Also most people don't realize common solutions in various kits are just variations of TE
1) Standard 1x TE = 10 mM Tris-HCl, 1.0 mM EDTA (pH 8.0, but this should not be assumed)
2) Buffer AE = 10 mM Tris-HCl, 0.5 mM EDTA (pH 9.0)
NOTE: Apparently Qiagen puts buffer AE in different kits and they may NOT be the same composition
Values From: DNeasy Blood & Tissue Kit (50) cat#69504
3) Buffer ATE = 10 mM Tris-HCl, 0.1 mM EDTA, 0.04% Sodium Azide (pH 8.3)
NOTE: Apparently Qiagen puts buffer ATE in different kits and they may NOT be the same composition
Values From: QiaSymphony DNA Midi Kit cat#931255
4) Buffer EB = 10 mM Tris-HCl (pH 8.5)
* Common elution buffer in many Qiagen plasmid prep and sample clean-up kits
4) DNA Hydration Solution = 10 mM Tris-HCl, 1.0 mM EDTA (pH 7-8)
* Provided in Gentra Puregene kits from Qiagen
5) DNA Suspension Buffer = 10 mM Tris-HCl, 0.1 mM EDTA (pH 8.0)
* Recommended by Affymetrix for SNP arrays (basically in standard TE it will not work!)
* Commonly called TElowE
A bit of ranting/suggestions
A) Never put DNA in water alone
B) The pH of the solution can effect things like 260/280 values, generally higher pH makes things look better
C) You want to watch the final concentration of EDTA in any enzymatic reaction. Our recommendations is to use buffers with 0.1 mM EDTA
In our lab all DNA samples are stored in DNA Suspension Buffer (ie. TElowE)
There are many great things about the GATK software, primarily their excellent level of support, but somethings are a bit irritating like "we change it all the time so check the --list function" which requires you to prep up a complete GATK command which can be difficult when you have no idea what you are doing.
So here are the lists I'm always wanting to have on my wall (The Genome Analysis Toolkit (GATK) v2.3-4-g57ea19f)
Variant Annotator List of Annotations
Standard annotations in the list below are marked with a '*'.
Available annotations for the VCF INFO field:
Available annotations for the VCF FORMAT field:
Available classes/groups of annotations:
If you're like most of us here in the lab, we can never keep track of what the difference is between the different types of Dynabeads. It seems like every new assay is asking for a different "letter" of bead. There's T1, C1, M270, M280, and well... I think that's the end of the list for now. At any rate, I decided to post a quick cheat sheet so it's easy to look it up and remember. So for anyone else out there wondering the same thing... hopefully this will make your life a little easier! For more details, check out the link for the LifeTech product sheet also included below.
I wanted to talk a little about the selection
characteristics of Agencourt’s AMPure beads,
a bead-reagent combination that purifies PCR reactions.
This stuff is incredible in terms of simplicity, efficiency,
and high-throughput compatibility. I have a sneaking suspicion that AMPure, not
unlike fire to Prometheus, was handed down from the gods to benefit humanity. You
just dunk it into your sample, slosh it around, stick it to a magnet, wash,
wash again, and elute in your favorite buffer. No muss, no fuss.
We were wondering, though, about its selection process. What
size fragments are selected by the AMPure beads, specifically at which ratio of
beads to sample? So, like diligent scientists, we rolled up the sleeves of our
labcoats and… read the protocol.
The protocol recommends washing your sample in a 1.8:1 ratio
of beads to sample, although it says that fragments less than 100bp will be
omitted at this ratio, it doesn’t say which sized fragments will be selected.
We found this remarkably helpful technical bulletin, which describes
calibrating each batch of AMPure beads with various ratios of DNA ladder.
So I did our very own calibration with AMPure beads using
Fermentas’s GeneRuler™ Low Range DNA Ladder (25-700 bp). I added 30ul ladder to
various concentrations of AMPure beads according to Agencourt’s instructions.
(Actually, if you’re looking for good AMPure instructions, I
recommend looking at Illumina’s TruSeq™ Sample Preparation Guide. Honestly,
their instructions are more comprehensive than Agencourt’s, and easier to
read.) After purifying each sample, I bookended the various AMPure:ladder
ratios with 10ul non-purified ladder on a 2% TBE gel for easy comparison.
Without any further ado, here are the results:
The results aren’t too surprising, I guess. Unless you’re
looking to select 100-150bp fragments, or if you’re using an extremely low
ratio of AMPure beads, the ratio differences aren’t that significant.
Basically, barring the first exception, you’ll be just fine following
Agencourt’s protocol and recommended ratio.
From this one image, it’s difficult to quantitatively
compare one ratio against another, so I plugged everything into ImageJ to give
me some numbers to play around with. I followed ImageJ’s guidelines for analyzing gel images. Then, I averaged the band intensities for both
non-purified ladder samples, multiplied them by three (knowing that I added
three times more ladder for purification), and normalized the band intensities
of the purified ladder by dividing them by their corresponding band intensities
for the non-purified ladder.
If you didn’t follow the grammatical train wreck that was
the previous sentence, don’t worry, you should just focus on the results:
Interestingly enough, according to ImageJ, the 1.6:1 ratio
has slightly more intense bands, and apparently slightly more purified DNA, than
the recommended 1.8:1 ratio. (If you want to see my exact analysis process, you can view the attached Excel file.) While those
values don’t mean percentages because the normalization isn’t exact, it does
suggest that different AMPure ratios to DNA can produce different results in
terms of fragment size and amount retained.
And, when you really think about it, isn’t that what experimental
PCR purification fragment analysis is all about?
After performing a test of heated DNA (95 C for 10 minutes) versus non-heated DNA for Kristi using the ARD_SSC.C8 cell line, we found that heating DEFINITELY denatures the DNA, literally to smithereens. We decided that I should try to fluctuate the heating times between 0 and 18 minutes and compare those to non-heated DNA. I have attached an image of the gel results which turned out beautiful I am proud to say!
I used the KMS34 cell line for this experiment. The non-heated DNA stayed at the top of the gel while the other 9 samples showed quite a variety of smear patterns. As you can see, when the DNA was heated for 18 minutes, it was denatured to the point that it lies between about 500 and 100 base pairs. On the other end of the spectrum, the DNA that was heated for 2 minutes shows a really nice, long smear from about the 12000 base pair marker down to about 650 or so.
After thinking about the effects heat has to DNA, there are both pros and cons to heat denaturing, and heating to differing times. If you want to get small fragments of DNA, for instance for use in a CGH protocol, heating for 18 minutes seems to do the trick. If you want to amplify long-length DNA via PCR, I would definitely say to avoid heating 18 minutes and try to stick to a time around 1 and 4 minutes. In talking with Jonathan, this image might explain why it tends to be difficult to amplify an DNA sequence of a large size, say 8 kb for example. Because we are heating the DNA at 94 C for 5 minutes and then proceeding to heat for another 15 or so, it is no wonder that we have a hard time getting many 8 kb amplicons. They are all shredded to pieces!
Just an interesting piece of data we figured out in the Keats lab today. Ending the week with interesting results is always a treat!
So the lab is now over one year old and finally getting going. I've had a number of people tell me that if you are going by six months and feel productive by one year you are doing good. Based on that I'm a bit behind as I only now feel like the lab is starting to really make progress, so that is a bit past the one year mark. But I really only started doing things in the lab in January when Kristi joined the group so maybe we are ahead of schedule? The good news is we have a decent bit of funding between my start-up, and two different grants. I actually had to hire another research associate recently as we just could not keep up. Then we have a big project ready to launch and when the contract is finally signed I'll be able to add two post-docs to the group.