[pmiguel]

Some of the DNA is damaged during shearing. My experience has been under certain conditions, only half of the ends can be repaired on one molecule. I don't know the reason yet, but I think the DNA was turned into single stranded. The damages,I think are pretty much mechanical, not chemical. Hydro shear is pretty low speed but will generate just a little better quality DNA. Some of the damages just can be repaired. I tried to repair the single-end adaptor ligated product, but it didn't go beyond that no matter how much enzyme I put in.

Could you give us a little more detail on what the conditions were where only "half of the ends can be repaired on one molecule"?

Any 5' or 3' overhand (locally single stranded) can be blunted by T4 polymerase's 5'->3' polymerase or 3'->5 exo-nuclease activity (respectively). The polymerase activity does rely on a 3'-OH, but T4-PNK should remove any 3' phosphate that would otherwise block the polymerase.

So, if your end repair regimen includes both T4 polymerase and T4 polynucleotide kinase (and most do) the remaining culprit would be non-phosphate, non-hydroxyl DNA fragment ends.

I have been able to find little in the literature about this possibility, the classic Richard and Boyer paper [1] mentions it and I summarized their results here:

http://seqanswers.com/forums/showthread.php?t=2759

As far as a comparison to the hydroshear, it seems like the results published by Oefner, et al. in their 1996 publication describing the prototype of the hydroshear [2] where 20-40% of fragments produced could be ligated with no end repair suggests it is qualitatively different from sonication.

1. Richards OC, Boyer PD (1965) Chemical Mechanism of Sonic Acid Alkaline and Enzymic Degradation of DNA. Journal of Molecular Biology 11: 327-240.
2. Oefner PJ, HunickeSmith SP, Chiang L, Dietrich F, Mulligan J, et al. (1996) Efficient random subcloning of DNA sheared in a recirculating point-sink flow system. Nucleic Acids Research 24: 3879-3886.

--
Phillip

[nextgen]

Some of the DNA is damaged during shearing. My experience has been under certain conditions, only half of the ends can be repaired on one molecule. I don't know the reason yet, but I think the DNA was turned into single stranded. The damages,I think are pretty much mechanical, not chemical. Hydro shear is pretty low speed but will generate just a little better quality DNA. Some of the damages just can be repaired. I tried to repair the single-end adaptor ligated product, but it didn't go beyond that no matter how much enzyme I put in.

[pmiguel]

Super-interesting thread. One obvious thing to reduce DNA damage in library prep is to use a visible light stain and a white light box for gel based size selection versus EtBr and UV. I have had good success with nile blue sulfate, an inexpensive visible light stain. I have not directly compared an identical library processed with EtBr/UV versus Nile Blue/white light so don't know the quantitative effect of UV damage though.

Yes, we don't use EtBr or UV light boxes in the lab at all. We just use SYBR Safe and one of those "Dark Reader" boxes. We bought the latter from Clare Chemical.

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Phillip

[greigite]

Super-interesting thread. One obvious thing to reduce DNA damage in library prep is to use a visible light stain and a white light box for gel based size selection versus EtBr and UV. I have had good success with nile blue sulfate, an inexpensive visible light stain. I have not directly compared an identical library processed with EtBr/UV versus Nile Blue/white light so don't know the quantitative effect of UV damage though.

[hanjaylee]

good challenge! i have been worrying about the same problem, too.

[DNADEB]

A side comment:
We found that the protocol for finishing the single stranded library did not neutralize the solution sufficiently and we were not recovering enough molecules here. Once we found that the pH was off and we had to add a lot more Na Acetate, our recovery sky-rocketed.

[pmiguel]

One can design assays for DNA damage using enzymes which target it and exploit the products the enzymes create. We've used antibodies to 8-oxoG to measure photo damage on arrays and it lights up like a X-mas tree without proper scanning buffers. So we dont have to live with this. It can be measured but requires alot of time and effort tuning a quantitative assay. Most assays are specific for an a priori lesion of interest so this is a long slog and quanting total DNA next to amplifiable DNA is probably too imprecise but should be able to pick up 90% effects like you mention (1 lesion eliminates a whole molecule).

Thymidine dimers from light through windows is one angle. I'd also point out low bind tubes and DNA adherence. While you are at, dont ship your DNA through certain zip codes or airports as the E-beam dosage varies. We're going through the steps which assume perfect DNA and seeing how poor these efficiencies are and then pushing them to highest we can achieve. If the simple hypothesis fails, we'll no doubt be digging into the damage more as the numbers in regards to loss are higher than anyone can explain.

great thread.. very pertinent to the field.

Naively, I would think that a method that hydrolyzes a DNA sample into mononucleotides followed by some flavor of mass spec would give the most global overview of DNA damage. Caveats there, of course. But damage introduced by the protocol itself could be discovered and corrected for, probably.

As to pyrimidine dimers, I seem to remember that there was an enzyme (from celery?) that reversed the dimerization process. Once we get to the point where we are repairing our DNA samples prior to assays, that would be a good enzyme to deploy. All the bacterial remedies for the pyrimidine dimer problem sounded far less appealing, if I remember this correctly.

--
Phillip

[Nitrogen-DNE-sulfer]

One can design assays for DNA damage using enzymes which target it and exploit the products the enzymes create. We've used antibodies to 8-oxoG to measure photo damage on arrays and it lights up like a X-mas tree without proper scanning buffers. So we dont have to live with this. It can be measured but requires alot of time and effort tuning a quantitative assay. Most assays are specific for an a priori lesion of interest so this is a long slog and quanting total DNA next to amplifiable DNA is probably too imprecise but should be able to pick up 90% effects like you mention (1 lesion eliminates a whole molecule).

Thymidine dimers from light through windows is one angle. I'd also point out low bind tubes and DNA adherence. While you are at, dont ship your DNA through certain zip codes or airports as the E-beam dosage varies. We're going through the steps which assume perfect DNA and seeing how poor these efficiencies are and then pushing them to highest we can achieve. If the simple hypothesis fails, we'll no doubt be digging into the damage more as the numbers in regards to loss are higher than anyone can explain.

great thread.. very pertinent to the field.

[seqAll]

...Also, even the 99% potential loss of this step only explains 2 of the >6 orders of magnitude of DNA loss in the Roche protocol...
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Phillip

Just wonder how precise the number 99% is. It was said >99%. But maybe 98.5%, 99.9%, 99.99, ...? So, that explains maybe 2, 3, 4...(unlikey to be 6 though) orders?

Loss or damage in other steps would be interesting to know.

[pmiguel]

You are right IF damage occurs randomly. IF NOT all the cool events happen in the 90% that you are loosing

There you go. Any time you lose 90% of a sample (for whatever reason), the remnant may be a biased representation of your initial sample. Because there is no reason to presume the loss is unbiased.

In the case of many of these library construction protocols we lose more like 99.9999% of our initial sample.

[What_Da_Seq]

You are right IF damage occurs randomly. IF NOT all the cool events happen in the 90% that you are loosing

[pmiguel]

In our case the DNA was not size selected so we cant blame the gel extraction or Ampure.

Not size selected at all? You must have size selected at some point, right? Otherwise you would get slammed with primer dimers, etc.

If you had size-selected prior to going into adaptor ligation, then all the DNA has the potential to form a legitimate amplicon. Whereas if not, a lot of that 750 pg would really be contributed by fragments outside a useful size range.

We spent some time looking into peroxide formation during the shearing step and monitoring heat. Curious if any one has used the NEB preCR kit or their Fpg and Hogg1 repair enzymes to repair other forms of DNA damage like 8-oxoG or glycosic bond breaks which may be induced by this method.
Alternatively, DNAse based forms of digesting DNA may be less caustic?

I have not. But I also have no clear idea how much DNA damage is present in a typical genomic DNA prep. If, for example, 90% of DNA spans longer than a few hundred bases contained lesions bad enough to stall out PCR replication, would we even notice?

PCR is an exponential process, after all. The 90% of strands that could not be extended would not contribute as template to later cycles. So the 10% that did extend far enough for the reverse primer to anneal would quickly overtake those that stalled.

Again, even if DNA damage isn't that bad in most DNA preps, it could be that a DNA prep you happened to have out near a window happened to pick up some pyrimidine dimers from the sunlight streaming in. Who knows. Heretofore all the assays I can think of would be insensitive to even fairly high levels of damage. If even 1% of the 1kb stretches of DNA in a prep are damage-free that still gives you 10 billion intact 1 kb stretches per ug of DNA. If the 10 billion work, then the 990 billion that do not will not be noticed.

We may just be entering an era where we do need all 1 trillion molecules. If so we need to either make sure this type of damage is not an issue or find ways to mitigate the damage.
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Phillip

[Nitrogen-DNE-sulfer]

Yes, Pairs are always better to measure start points and distinct molecules but with single ended reads we just assume if any single replicated start point is identical it was PCR induced. This is conservative and we only count unique placements in the genome..yet again an underestimate. The above example was 5 cycles of PCR with long extensions.
We have also performed no amplification libraries which eliminate the PCR replication problem. Taqman is another angle we use to measure at the linker step and I'll have to check our notes but I believe we are getting 300M-500M positive beads with 1ng of 200bp library. ABout 15% of our total beads (3B total) amplify which suggest few have 2 molecules in the bubble based on poisson. Not far off from your estimate of 5M molecules per 1pg of 200bp library or 5B per ng.

In our case the DNA was not size selected so we cant blame the gel extraction or Ampure.

We spent some time looking into peroxide formation during the shearing step and monitoring heat. Curious if any one has used the NEB preCR kit or their Fpg and Hogg1 repair enzymes to repair other forms of DNA damage like 8-oxoG or glycosic bond breaks which may be induced by this method.
Alternatively, DNAse based forms of digesting DNA may be less caustic?

Despite leaning high on beads in the emPCR this currently cant get a bead in every reactor without alot of clumping. Probably only 20-25% of the reactors populated. Raindance like techniques have been contemplated but would take days to make Billions of reactors.

[pmiguel]

Great Thread,

[...]
from just fragmenting DNA and adapting it and then counting distinct molecules on the back end we have gone as low as 750pg of already sheared DNA to generate 30-40M distinct 50mer human reads. I think this is a very key point. This is roughly 300 copies of the genome but most importantly, we didnt covaris this DNA. It came from Maternal blood streams so it was enzymaticaly digested in situ or in vivo.
[...]

(30-40M unique starts, right? Two reads that map to the same start position in a genome may (or may not) derive from a single input DNA molecule. That is why using single-sided reads makes this process difficult to assess.)

40M reads implies 40M 80-130 bp insert amplicons. What yield does that represent?

1 pg of 100 bp DNA is roughly 10 million molecules. So you started with 7.5 billion molecules (presuming they were all ~100 bp -- which, obviously, they would not have been). That would imply that roughly 1 in 200 of the original molecules were successfully converted to templated beads.

Was this DNA already size selected when you measured it as 750 pg?

I would expect any smear of DNA to be <10% in the fairly tight size distribution (150-200 bp for the full amplicon length == 80-130 bp of insert). Even as low as 1% would not be surprising.

So, yes that result would be consistent with nearly all the molecules being ligatable on both ends and amplifiable. Or less than 10% of them being so. Hard to say.

The reason I find this is intriguing is that all methods eventually go through a final Frag adaptor ligation so its important to know the efficiency of this step and its after all the simplest to measure.

By qPCR? By sequencing, it is not so simple. The human genome is replete with repetitive DNA, so single end reads are difficult to assess as to their derivation from a unique chunk of your original sample DNA. This is because the pre-PCR amplification step would make lots of copies of all amplifiable amplicons.

In terms of Covaris'd DNA, I will look through our data but we have performed 600M read on 1 ug buccal DNA Covaris'd from a patient and not saturated this library. We probably need to go deeper to understand if the different shearing methods are playing a damaging effect.

Yes, especially since minor changes in the shearing buffer may lead to different outcomes. I'm not a chemist, but if the C-O bond breakage that apparently predominates in sonication-mediated DNA fragmentation

http://seqanswers.com/forums/showthread.php?t=2759

can result in different fragment ends, then factors such as pH may influence which end-type does result. That is, a break between the C5' and O or C3' and O may result in the desired outcome: hydrolytic restoration of the end to a 5' or 3' OH. Or it could result in undesired outcomes such as ribose-sugar ring opening or maybe even loss of C5' entirely. (Again, I'm not a chemist, the above is rampant speculation.) Point being, mixtures of T4-polymerase/T4-PNK probably cannot repair the latter outcomes into something ligatable.

[...]
The final point I'd add to the discussion is that not all quantified DNA is amplifiable or makes it to a bead or to a cluster. We're working with emPCR on SOLiD and we assume a 1/2 to 2/3rds of our reactors have DNA and no beads. We lean on the pushing the bead poisson high and the template poisson low as 2 beads in a reactor dont kill us but 2 templates do.

Then it does not seem you would lose many amplicons in ePCR. That is, bead poisson high: therefore nearly all reactors have beads. Template poisson low -- most of the reactors have a bead but no template, but where there is a template it will almost certainly have a bead to bind it.

Similar effects may exist on the poisson curves for clusters...ie Flow cells must be flooded with 1 concentration where only a portion of this concentration can seed the flow cell surface but molecules exist throughout the whole volume and I'm still unclear if both surfaces amplify and only one being imaged creates another factor of 2 loss?

I don't have a Solexa, so I don't know. But I will note, tangentially, that it is interesting that after a couple of years of direct competition between Solexa and SOLiD, it now appears that the two platforms are veering into slightly different niches. Solexa, with paired-end 100 base reads seems poised to conquer the de novo sequencing niche. Whereas SOLiD appears to have abandoned longer reads to concentrate on increasing read numbers. Which, everything else being equal, would give them control of the resequencing niche (including digital gene expression). That said, everything else is not equal. Illumina had instruments out in the field at least a full year before AB did. And then there is the PacBio instrument looming...

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Phillip

[Nitrogen-DNE-sulfer]

Great Thread,

Stepping back a bit to dissect this we have been testing how far we can go with simpler Fragment libraries as a first measure of this. Most Circularized protocols have many inefficient steps and we began by quantitating how much DNA we can get from just fragmenting DNA and adapting it and then counting distinct molecules on the back end we have gone as low as 750pg of already sheared DNA to generate 30-40M distinct 50mer human reads. I think this is a very key point. This is roughly 300 copies of the genome but most importantly, we didnt covaris this DNA. It came from Maternal blood streams so it was enzymaticaly digested in situ or in vivo. It also has a very different GC content than Covaris DNA not surprisingly.
The reason I find this is intriguing is that all methods eventually go through a final Frag adaptor ligation so its important to know the efficiency of this step and its after all the simplest to measure. We will be backing up into the various circularization protocols shortly but already know the SOLiD circles are 10-20% efficient at the lengths mentioned above.

In terms of Covaris'd DNA, I will look through our data but we have performed 600M read on 1 ug buccal DNA Covaris'd from a patient and not saturated this library. We probably need to go deeper to understand if the different shearing methods are playing a damaging effect.

I found the complete genomic paper fairly well written in regards to exact pmols at each step. Lots of amplification along the way but its clear we need protocols which speak to these quants at every step with the other platforms as well.

The final point I'd add to the discussion is that not all quantified DNA is amplifiable or makes it to a bead or to a cluster. We're working with emPCR on SOLiD and we assume a 1/2 to 2/3rds of our reactors have DNA and no beads. We lean on the pushing the bead poisson high and the template poisson low as 2 beads in a reactor dont kill us but 2 templates do.

Similar effects may exist on the poisson curves for clusters...ie Flow cells must be flooded with 1 concentration where only a portion of this concentration can seed the flow cell surface but molecules exist throughout the whole volume and I'm still unclear if both surfaces amplify and only one being imaged creates another factor of 2 loss?