[RezaFalsafi]

deblocking FC

Hello I am new to this forum and sorry if the posts are old I just came across the all the above posts and the one about deblocking the FC and re-using it caught my eye. can it be please explained more as how to deblock? and is the FC used for the same run we tested with 6 cycles or just it is used for 6 cycle testings?
Thanks a lot

[pmiguel]

Thank you Phillip and SeqR&D,

The discussion was very insightful!! It has given me novel aspects to think about...
I would like to add on quick questions where your suggestions would be needed...

1. Is there a way to detect presence of EtBr in our sample library before we proceed to purification?

It would be possible fluorimetrically. But I don't have details. If you did some sort of clean-up after a step that used EtBr, you could at least hope the concentration of EtBr would drop below a threshold where it might be a problem.

2. I have observed variation in concentration of PhiX V2 & V3 libraries,the fragment size explains this.In this context do you suggest to quantitate phiX library with pico green as well to have a better comparison with sample library?

We now run a high sensitivity bioanalyzer chip on all new tubes of PhiX. That will give the length of the library and clue you into to abnormalities (like adapter dimer peaks...)

3. Do you use Truseq PPC for qPCR as well?As I have been using the in-house primers which was synthesized by the sequences provided by Illumina qPCR protocol.I have run melt curve and we see significant primer dimer peak which we have worked a lot on to remove but have been unsuccessful!!

A couple of things here:
(1) I don't think a bimodal melt curve is an indication of a primer dimer. Call me a heretic if you will, but why would we expect library melt curves to be monomodal? Any reason? Will not a not the complexity of a library, and the input sample from which it derives, cause one to see various melt curve modalities?

Anyway, I guess the TruSeq adapters themselves will be one of the peaks -- they don't have to be in an adapter or primer dimer to have their own melt temp.

A melt curve, I would hazard to guess, is for standard qPCR, where there will be a single gene being amplified. There your melt curve should be monomodal because it is a single sequence that is being amplified. But a library is a complex mixture of molecules all flanked by the same adapters.

(2) We just use cheap oligos we had synthesized for qPCR. I think of the PPC as being a limited resource, so it never occurred to me to use them.

--
Phillip

[Genquest]

Thank you Phillip and SeqR&D,

The discussion was very insightful!! It has given me novel aspects to think about...
I would like to add on quick questions where your suggestions would be needed...

1. Is there a way to detect presence of EtBr in our sample library before we proceed to purification?
2. I have observed variation in concentration of PhiX V2 & V3 libraries,the fragment size explains this.In this context do you suggest to quantitate phiX library with pico green as well to have a better comparison with sample library?
3. Do you use Truseq PPC for qPCR as well?As I have been using the in-house primers which was synthesized by the sequences provided by Illumina qPCR protocol.I have run melt curve and we see significant primer dimer peak which we have worked a lot on to remove but have been unsuccessful!!

At one point of time,i had used Illumina PPC for qPCR and the additional dimer peak disappeared and Efficiency improved but the PhIX conc. was even higher than previous ( i.e. when run by in-house primers).Don't remember which version of PhiX I used,will have to put a check on this as well!!
Suggestions and advises are welcomed!!
thank you once again

[pmiguel]

If you qPCR a non-ampified sample that has varying amounts of free adapter (with a keenly designed SPRI purification you can greatly minimize this), library elements with one ligated adapter and those with both, you will be quantifying the 2 adapter libraries, but the other stuff is in there as well.
I'm not exactly sure what ligation efficiency is, but from what I understand it's not great...60% to 70%? (correct me if I'm wrong).

You are right, I think. If anything 60%-70% is on the high side. Although if you presume that 60% means "60% of ends get adapters ligated to them", one would expect only 36% to get both ends adapter ligated.

Actually on Friday we looked at the results from a similar ligation chemistry -- the Roche Rapid Library prep kit for 454 library construction -- and saw 12% of the molecules were amplifiable via qPCR. Not entirely clear this is a result of whether adapters were ligated on or not. The genomic DNA given us was in questionable state -- prepared from old refrigerated leaf tissue or even twigs -- it was degraded to 5kb average length prior to any sonication. So the DNA may just have had a moderately high frequency of damage not circumventable by the qPCR polymerase.

Hmmm, actually the Roche Y adapters have FAM modification. So I guess we could actually assess the number of molecule ends with adapters ligated to them.

That could leave a lot of non-clusterable DNA fragments floating around that will "block" the flowcell surface from clusterable library. I'm not sure if this can account for a 99% reduction in cluster density from one to the other. Keep in mind, this is only my best guess.

Yeah, I like the way you think. The possibility of a blocking issue had not occurred to me prior to your previous post.

As it turns out the attempts we made with no-amp libraries were made from bacterial genomic DNA. So we were only shooting for 10% of a lane for each. (Even that would have been overkill...) So if there were material in the library blocking access of the flow cell oligos to library molecules we would see an effect on the other libraries sharing the lane commensurate with what we saw with with the no-amp libraries. But the other libraries seemed unaffected.

Also, we had previously ran amplified versions of these libraries and they seemed to produce plenty of clusters -- but they were terribly biased. All of this could be the result of the DNA of the organism (Deinococcus radiodurans) -- who know what modifications it hosts. But since Illumina tech support suggested that very poor clustering efficiency was to be expected for TruSeq libraries that had not undergone at least 1 cycle of amplification, I tend to draw other conclusions.

Obviously, I could have this all wrong. Just a couple of hypotheses...

--
Phillip

[SeqR&D]

If you qPCR a non-ampified sample that has varying amounts of free adapter (with a keenly designed SPRI purification you can greatly minimize this), library elements with one ligated adapter and those with both, you will be quantifying the 2 adapter libraries, but the other stuff is in there as well. I'm not exactly sure what ligation efficiency is, but from what I understand it's not great...60% to 70%? (correct me if I'm wrong). That could leave a lot of non-clusterable DNA fragments floating around that will "block" the flowcell surface from clusterable library. I'm not sure if this can account for a 99% reduction in cluster density from one to the other. Keep in mind, this is only my best guess.

[pmiguel]

Neither of those are true. I'm certain that if you run a denaturing gel on the adapters, that they will run as 2 bands.

What makes you certain?

And, PPC primers are complimentary to the adapter ends...nothing special.

Why do they run ~80 nt on a pico RNA chip if they are generic oligos?

Where you might run into issues are with A-tailing efficiency, ligation efficiency, possibly ligating 2 adapters together,

Okay, I'm with you so far...

having free adapters present when clustering a flowcell (they will take up space, but will not extend or show clusters).

And PCR will dilute out the free adapters, okay, I guess.

Doing some amplification enriches for library elements that have proper adapter ligation on both ends.

I see your point. But when we tried no amp libraries we used qPCR to determine the concentration of library elements that had proper adapter ligation on both ends. That is how we determined how much to add to the flow cell. qPCR (SYBR green) should not be confounded by any of the factors you list above.

Yet, when we clustered using the unamplified libraries, we saw much lower cluster yields from those libraries. (Like 1% of what was expected, if I recall correctly.) Our qPCR was not otherwise way off -- all the other libraries we quantified this way clustered at about the expected efficiency. So it seems like the simple explanation is that there is something odd about the TruSeq system that Illumina has not revealed.

--
Phillip

[SeqR&D]

Neither of those are true. I'm certain that if you run a denaturing gel on the adapters, that they will run as 2 bands. And, PPC primers are complimentary to the adapter ends...nothing special.

Where you might run into issues are with A-tailing efficiency, ligation efficiency, possibly ligating 2 adapters together, having free adapters present when clustering a flowcell (they will take up space, but will not extend or show clusters). Doing some amplification enriches for library elements that have proper adapter ligation on both ends.

[pmiguel]

There is no magic in the PPC. Adapters are not a complete "library element"...which is the beauty behind the Y-adapter. You do not have complimentary ends...they are "sticky", and only contain half of what you need to cluster (only half of your library elements have what it takes to hybridize to the flowcell). They should however qPCR with more accuracy than an order of magnitude...as only the first cycle of qPCR should be affected...but maybe thats enough....I haven't done the math.

From Illumina's FAQ:

Is there a PCR-free TruSeq Sample Prep protocol available?

The new index adapter design enables PCR-free protocols. (A single cycle of synthesis is required to separate the forked adapter.)

Upon querying Illumina tech support, I got the following response:

We have found that if you simply take the annealed fragment that has the forked adapters on each end and add it to the clustering reaction that even with NaOH treatment before clustering, there is poor initial hybridization and the resultant clustering is poor. By allowing at least one PCR cycle this separates the strands and the forked adapters, allows a single amplification of the entire strands, resulting in a clean, double-stranded species that gives cleaner clustering results.

This makes no sense unless one or both of the following is true:

(1) The adapter oligos are attached to each other in some way that resists their melting apart during NaOH treatment.

(2) The PPC primers themselves have some modification that causes them to bind especially well to the flow cell.

Or, at least, that is my take on the situation.

--
Phillip

[SeqR&D]

There is no magic in the PPC. Adapters are not a complete "library element"...which is the beauty behind the Y-adapter. You do not have complimentary ends...they are "sticky", and only contain half of what you need to cluster (only half of your library elements have what it takes to hybridize to the flowcell). They should however qPCR with more accuracy than an order of magnitude...as only the first cycle of qPCR should be affected...but maybe thats enough....I haven't done the math.

[pmiguel]

There are factors that might confound qPCR. They include:

(1) Ethidium Bromide in your sample. I did not do a careful test of this. But someone in the lab tried spiking EtBr into a sample at a low level. It shifted the Ct by a cycle or two. Ethidium Bromide will also confound fluorimetry, so purifying it away after some size selection methodologies is important.

(2) Your phiX may not be the concentration and/or length that you think it is. Most current (v3) phiX libraries from Illumina are supposed to be about 500 bp. So far every one that we tested has been around that size. I think v2 (including v2 multiplex) phiX is supposed to be 350 bp. The v2 multiplex phiX has always tested out to 350 bp or so for us. But we saw one case where the non-multiplex v2 was 500 bp, rather than 350 bp. Further, I have seen a 500 bp library that had a very weak, but visible, 350 bp peak. Also one library had a ~70 bp peak. I don't know what the adapter length is for these library types, so I don't know if this is an adapter dimer. If it was, then it was 10% of the molar total. But would produce little signal during SYBR Green qPCR.

Since the phiX standards are likely calibrated primarily to produce a given number of clusters, the actual size of the library molecules may not be a great concern for Illumina. However, SYBR green assays do not directly determine molarity -- instead they rely on you to supply the correct average size of libraries. So:

(3) You must have an accurate determination of both your library's average length and your concentration standard's length.

(4) One more: TruSeq libraries, if they are run no-amp, cluster at least an order of magnitude lower than qPCR would suggest they should. Illumina tells you, that you must do at least one cycle of enrichment PCR to "break the fork" of the adapters. Taken literally, this would suggest that the adapters are covalently linked strand-to-strand by a chemistry that breaks during extended heating but not by NaOH. Alternatively the adapters might lack some critical "magic" present in the PPC -- PCR primer cocktail -- of the TruSeq DNA/RNA prep kits. So, these factors could result in issues, if true.
--
Phillip

[Genquest]

Library qualification by qPCR

Hello everyone,
I am pretty new to this forum and stumbled on this post as even I am facing problem with qPCR quantification method
1.Has anyone encountered with, a higher qPCR concentration than the estimated concentration by pico green??
2. Also if assumed that qPCR is more accurate(w.r.t pico green) as it would give conc. of ssDNA ligated with adapters,then the cluster generation should be accurate with qPCR conc. but it doesn't happen so...the flow cell (V3,Hiseq 2000) remains under clustered.Can any one think of possible reasons??
3. This trend seems to change with every library.Is absolute quantification the right approach for Sybr green qPCR method?Also I am using the PhiX control provided by Illumina as my standard.
4.Do people take the "exact concentration of libraries" as determined by qPCR for cluster generation or they compare it with pico green to see the relative trend of concentration?

Would appreciate if any of the queries are resolved!!
Thanks in advance

[SeqR&D]

I wouldn't think so. The most difficult part here is aligning your qPCR results to your cluster #. After that, your cluster # should not change dramatically enough to have to redo the titration. If you see that your cluster # is going down/up all you have to do is change your concentration.
Example: If you are generally clustering at 8pM and you see that your cluster density has dropped across the board (even PhiX control lane), then you might want to start clustering at 10pM or 14pM (and change PhiX if necessary).....

It is possible to make a mistake when clustering the flowcell, so be careful. I occasionally get flowcells that will fail the first base report with low cluster density, but it was the process or the flowcell or the clustering that was the issue...not the concentration.

[lhandley]

Am I correct to guess that the titration run on the flowcell would have to be repeated with each software upgrade and/or formulation change since that would affect cluster #?

[SeqR&D]

This is how we did it:

1. We wasted a FC
2. Amplified (~14 cycles) gel purified/size selected library (in the size range that you use to sequence)
3. Purified the PCR products
4. Nanodrop quantified the PCR products
5. Made dilution series to hyb to a FC
6. Pick the "best" dilution that gives desired cluster #
7. From this info, create a qPCR STD which you will use to quantify every subsiquent sample that is going on to sequencing. Make tons of this stuff (alliquoted in strip tubes and left in the -20C), and do not run out without using a set to quantify your next batch of STD!!
Our qPCR STD dilution series is this:
100pM, 30pM, 10pM, 3pM, 1pM, 0.3pM, 0.1pM, 0.03pM

As long as you always cluster the amount that you found in step #6, you should be great.

Let me know if this doesn't make sense.

[simonandrews]

is this abt the in lane Phix experiment?

It's more the other way. Illumina now offer an option to spike small amounts of PhiX in with each lane for QC purposes, but these don't help if you need a control lane for calibration purposes. If you're still running a full PhiX lane you can spike in another library to that to get a low level of coverage which won't affect the normal function of the control lane.