[Nighthawkrao77]

Is there evidence that you get very high RNA whole transcriptome coverage using this method compared to others? I know in the paper it claims that this comparison wasn't done because input/cost are also floating variables.

[EvilTwin]

Is there evidence that you get very high RNA whole transcriptome coverage using this method compared to others? I know in the paper it claims that this comparison wasn't done because input/cost are also floating variables.

Hi Nighthawkrao,


which metrics are of predominant interest for you?


In terms of library complexity on whole transcriptome for 1 ng (rRNA-depleted) and below, CATS gives very good results that are comparable to using > 50 ng with other kits.



We have verified the expression levels of several highly as well as lowly expressed genes in our CATS runs with conventional RT-PCR and get a very good concordance.


A direct comparison to conventional kits (using the same amounts) is truly not easy because they do not work with the CATS low input ranges.


The exact coverage of the transcriptome/certain genes depend on the composition of the RNA sample (what type of RNA is predominant) origin (cell line/tissue/plasma etc, cell cycle/expression status of genes of interest, etc) and aside from the input is a function of fragment complexity captured during library generation (which is the job of CATS) and of how many reads you actually sequence.


Did I understand your question correctly?

[Nighthawkrao77]

Hi EvilTwin,

Yes you did understand my question. Thanks for the response. Can CATS be used at equivalent quantities as the other kits? E.g. if you added 100ng into the reaction would it flood the system or create negative effects compared to other kits?

[jwfoley]

Why the 3' V anchor instead of the more common VN, on your oligo(dT) RT primer?

[HeidelbergScience]

Hi EvilTwin,

Yes you did understand my question. Thanks for the response. Can CATS be used at equivalent quantities as the other kits? E.g. if you added 100ng into the reaction would it flood the system or create negative effects compared to other kits?

We never tested >10 ng of DNA or RNA per reaction. However, if you use 10 ng (300 bp DNA or RNA) - the product is already strongly visible after pre-amp cycle 8 (if using the optimized protocol described in this forum). We never used less than 8 PCR cycles for the pre-amp, because pre-amp is aimed to generate dsDNA libraries with the Illumina P5 and P7 adaptors from single-stranded first cDNA strand.

Importantly, the complexity of libraries generated from >10 ng nucleic acids will be unnecessary high. Thus, 10 ng of input should yield a library with the complexity (a number of different fragments) that already exceeds the capacity of a 200M-read HiSeq run. For example: already 10 ng of 300 bp DNA is approximately 30 000 000 000 different molecules. If we assume that the efficacy of TSO at those concentration is close to 80%, the libraries would contain a similar number of different reads.

[HeidelbergScience]

Why the 3' V anchor instead of the more common VN, on your oligo(dT) RT primer?

The V primer has 4x higher efficient concentration as compared to VN, and it also decreases the probability of the formation of "empty" libraries.

[HeidelbergScience]

Importantly, the CATS protocol described in the paper (and in this forum) is optimized and tested only for 1 pg/µL - 10 ng/µL (final concentration in polyA / polydA tailing reaction) of RNA and DNA amounts. When using higher inputs (>10 ng/µL), additional optimization might be necessary (e.g. ATP/dATP concentration, primers concentration, incubation times).

[jwfoley]

Why do you use T4 PNK instead of a regular phosphatase (e.g. SAP)? Do you need 5’ phosphates for some reason?

Also, why do you say sonication results in 50% 5’ phosphates, 50% 3’? The literature discussed in this thread says there's a "great preference" for 5’: http://seqanswers.com/forums/showthread.php?t=2759

Have you tried using sonicated DNA without phosphatase treatment?

[HeidelbergScience]

Why do you use T4 PNK instead of a regular phosphatase (e.g. SAP)? Do you need 5’ phosphates for some reason?

Also, why do you say sonication results in 50% 5’ phosphates, 50% 3’? The literature discussed in this thread says there's a "great preference" for 5’: http://seqanswers.com/forums/showthread.php?t=2759

Have you tried using sonicated DNA without phosphatase treatment?

T4 PNK removes cyclo-phosphate and phosphate from 3-OH of RNA and DNA, but in the presence of ATP is also phosphorylates 5-OH. For CATS you need only to free the 3-OH group. The method doesn't care about a 5'-phos, and therefore there is no need to add ATP together with with T4 PNK.

https://www.neb.com/products/m0201-t4-polynucleotide-kinase

After sonication at least some % of DNA fragments should have 3'-phophates, although its proportion may vary from condition to condition. One should repair the 3'-ends to be on a safe side.

[HeidelbergScience]

Why do you use T4 PNK instead of a regular phosphatase (e.g. SAP)? Do you need 5’ phosphates for some reason?

Also, why do you say sonication results in 50% 5’ phosphates, 50% 3’? The literature discussed in this thread says there's a "great preference" for 5’: http://seqanswers.com/forums/showthread.php?t=2759

Have you tried using sonicated DNA without phosphatase treatment?

Other phosphatases may also work well, but we did not test them.

We tested sonicated DNA w/o T4 PNK, and the library yield was somehow lower, but not more than 50%.

[HeidelbergScience]

I can´t remember this detail off the top of my head, but I would say just slightly worse (a noticeable difference but not huge). All comparisons (Smarter2 vs the Smarter kit) can be found in the Suppl material of the Nat Methods paper.
Suppl table 4 has a list of all the comparisons with details in what they are different between each other.
Suppl table 3 has the detail of all the single cells analyzed with each protocol (sheet A) as well as the statistical analysis of the different variables (sheet B), where we looked not only at the TSO but also PCR enzyme, additives, etc etc.
Suppl Figure 1 and 2 give you an idea of the difference in the sensitivity and variability of different protocols.
In short, the rGrG+G oligo is better than rGrG+N, which is anyway better than the TSO from the kit.

PROTOCOL UPDATE
In our tests rGrG+G and rGrGrG ending TS oligos had equal efficacy (3 independent experiments on 22nt synthetic RNA and DNA).

Importantly however, the quality of the same TS oligo can vary from batch to batch, and affect the yield of the library to some extent.

[jwfoley]

In our tests rGrG+G and rGrGrG ending TS oligos had equal efficacy (3 independent experiments on 22nt synthetic RNA and DNA).

This is consistent with my tests on a different template-switching protocol: I found that the LNA actually gets slightly lower yields, in every test I've run, except a much higher background of self-priming artifacts. It probably depends on the sequence of the TSO (the Illumina P1 adapter has a run of three cytosines that might hybridize to the G-overhang), but I wonder whether the extra yield reported in the Smart-seq2 paper is actually due to non-specific strand invasion rather than proper template switching (http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3853366/).

At any rate, this should go without saying, but just a public service announcement since I've met many people who don't seem to be aware: you can reduce your batch-to-batch variation in oligo effectiveness by measuring the true concentrations of your stocks by spectrophotometry, rather than just assuming the manufacturer correctly measured the amount that's in the tube and your resuspension efficiency was exactly 100%. I often find that my "100 μM" stocks are as little as 90 μM, or sometimes even above 100. But then I dilute accordingly before I aliquot.

[Simone78]

This is consistent with my tests on a different template-switching protocol: I found that the LNA actually gets slightly lower yields, in every test I've run, except a much higher background of self-priming artifacts. It probably depends on the sequence of the TSO (the Illumina P1 adapter has a run of three cytosines that might hybridize to the G-overhang), but I wonder whether the extra yield reported in the Smart-seq2 paper is actually due to non-specific strand invasion rather than proper template switching (http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3853366/).

The strand-invasion is a possibility and the risk of that is higher when you use LNA-based TSO. However, if you would read our papers (properly this time) you might notice that this is only one of the changes we introduced in the original Clontech protocol. The effect of betaine and the extra MgCl2 (and, to a lesser extent, the dNTPs in the lysis buffer or the extra cycling at the end of the RT) are as important for increasing the yield.
You might also notice (reading the Supplementary), that a TSO with 2 or 3 LNA-G gives actually LOWER yield than the one with only 1 LNA. How do you explain that?
And I haven´t seen more self-priming artifacts with a LNA-TSO. Or lower yield, for that matter. Obviously it can happen...

[jwfoley]

However, if you would read our papers (properly this time) you might notice that this is only one of the changes we introduced in the original Clontech protocol. The effect of betaine and the extra MgCl2 (and, to a lesser extent, the dNTPs in the lysis buffer or the extra cycling at the end of the RT) are as important for increasing the yield.

I did see the parts about the betaine (trimethylglycine), magnesium, and denaturation/activation of the dNTPs, and I reproduced those results successfully. My comment was only about the LNA TSO.

You might also notice (reading the Supplementary), that a TSO with 2 or 3 LNA-G gives actually LOWER yield than the one with only 1 LNA. How do you explain that?

Multiple LNAs in the same oligo aren't typically used in several consecutive positions, especially at 3′ ends, because that makes the molecule too rigid and/or it anneals too tightly to be extended by a polymerase (see e.g. http://link.springer.com/chapter/10....387-32956-0_13). If you had asked Exiqon to design the oligo for you, I suspect they would not have given you those. This is also consistent with the self-hybridization hypothesis.

And I haven´t seen more self-priming artifacts with a LNA-TSO.

As I said, this should depend on the rest of the TSO's sequence. The Smart-seq2 TSO doesn't have more than two C's in a row for the G-overhang to stick to. The Illumina P1 adapter contains three C's in a row, though CATS cleverly truncates the primer to remove those. My test was on a version that included the three C's.

So to clarify, my hypothesis is that the LNA may or may not increase template-switching, but it definitely promotes strand invasion (as shown in the BMC Genomics paper), and it likely creates a problem with self-hybridization if your TSO has a certain sequence. The second factor may account for some or all of the apparently increased yield in Smart-seq2 (which is arguably still a good thing if your goal is to maximize sensitivity), and if it accounts for all of it, that would explain why we don't see the same effect with protocols like CATS that use shorter templates (shorter strands have fewer places to invade instead of annealing at the end). TSOs with more than one LNA don't work for the same reason that other primers with multiple LNAs at the 3′ end don't work.

[HappyMonocyte]

Can this be modified to work on the Ion Proton platform?