[Smriti]

Thanks everyone for your input!
Right now I am looking for a preliminary estimation on % ds DNA recovery from my experiments. So, Qubit looks best to me after reading posts on this thread.

[pmiguel]

A word of caution about qPCR -- don't forget there are two types. Most commonly one uses the SYBR green, a dye the fluoresces in the presence of dsDNA. But there is also "Taqman" style qPCR, that is probe based. If you know the average size of your amplicons, the former should work fine modulo minor (hopefully) differences in your amplification yield due to sequence composition biases. TaqMan should be insensitive to the size of amplicon.

The specific issue that throws off SYBR green style qPCR is the presence of small amplicons (usually primer dimer or adapter dimers) in a library. If you have done a size selection on your library, you might be fooled into thinking you have no small amplicon contamination. However if you size-selected on dsDNA, this may not be the case. Small amplicons can anneal to the ends of larger ones, effectively "hiding" from size selection. You need to QC you library as ssDNA to detect this. Fortunately this issue is rare for libraries constructed with the TruSeq kits.

If you do get the right titer, and the small amplicons do not predominate, then your results will be okay. Some of your sequence reads will be wasted on primer-dimers. But not that big a deal. If you undershoot on your titer, then you are potentially in big trouble. First, this can result in overclustering. Second, under conditions of over clustering Illumina's infamous susceptibility to low-diversity sequence in the first 4 bases of amplicons can cause major issues. This is especially confusing because it can result in low cluster densities--even though the lane is packed with clusters.

--
Phillip

[FredericRaymond]

Hello,

I have a related question. In the illumina Nextera protocols, they suggest to use the HS DNA bionalyzer chips. Would it be ok to use the 7500 DNA chips for library validation? I will also quantify by picogreen and, most probably, validate libraries by PCR.

Thank you,

FR

[NextGenSeq]

The Bioanalyzer chips you use depend on the yield of your library. We find if our libraries are 10 ng/ul or less in concentration we have to use the HS chips. If they are more you can use regular DNA chips.

However, with DNA 7500 you may have a hard time resolving any adapter artifacts at 120bp. We usually use the DNA 1000 chips.

[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

[Jon_Keats]

Not sure if I'll just add to to the confusion but in my lab we QC each library and pool by Kapa QPCR, Qubit, and Bioanalyzer. The frustrating thing is that our 5nM pools created based on the library QPCR results calculated as follows in general:

Expected 5nM based on QPCR Directed Dilution
Kapa QPCR - 5nM +/- 10%
Qubit - 3.5nM
Bioanalyzer - 2nM

In the end our core likes to cluster based on bioanalyzer and typically hit their cluster density. When I asked them to cluster based on our QPCR, not surprisingly we ended up with around half the expected number of clusters (bad way to turn a V3 flowcell into a V1 flowcell). That being said you can see there is a trend between the assays and in my judgement I think they are all consistent but I don't think you can just cluster 10pM regardless of method. Instead if maybe 10pM by Bioanalyzer, 15pM by Qubit, and 20pM by QPCR to get the same number of clusters. One recommendation we are trying to implement was suggested by Heisman to use a known library that generated the target number of clusters and use that as a control with each QC run.

[Genquest]

Library quantitation by qPCR

Thank you Jon_Keats,

I am relieved by reading your post,as I am not the only one facing such issues.
Would appreciate if any of the members would like to give their insight to understand the possible reasons behind such an observation!!
e.g.After pico green estimation we dilute the libraries to 2nM conc. and then use this dilution for qPCR.Ideally we should get less than 2nM conc. by qPCR estimation but normally its higher i.e. around 2.3-2.5nM.Now if we use the exact conc. estimated by qPCR for clustering the flow cell,we would use less i.e. below 10pM assuming the cluster density should be appropriate but we end up under clustering!!

Could it be due to primer or adapter dimers that gives the false conc. by sybr green qPCR assay??

[pmiguel]

Adapter dimers in your library will produce signal proportional to their length, not just their molarity, using SYBR green qPCR. But that would have the effect of qPCR underestimating your sample's molarity, not overestimating.

Pico Green fluorimetry is insensitive to single stranded molecules. So if 10-20% of your library is ssDNA, it would explain your results. Also EtBr (and probably other fluors) would tend to through your results off. A sample taken directly off a PippinPrep and read with a fluorimeter gave a completely aberrant emission spectrum such that we had to remove the residual EtBr before we could get a concentration. Lower levels of EtBr might confound fluorimetry, or qPCR, to a variable extent.

--
Phillip

[SeqR&D]

I agree with Jon Keats on his qPCR thoughts.

1. qPCR is completely dependant on your standards, and your dilution of the standards.
2. qPCR is also dependant on your dilution of your samples.

If you mess up either of those...everything falls apart. It is all too easy to make ths smallest error in diluting samples for qPCR...I have done it many times. Make sure you are working in a clean area...ie no libraries from other experiments flying around and getting into your current samples. Also, having calibrated pipets and a clean water source are a plus.

I always use qPCR to quant for clustering...why?
-In order for something to cluster, it must be a complete piece of library...ie, it must have the adapters ligated on both ends. The only metric that will tell you how much of your library has adapters is qPCR. All of the other methods tell you total DNA...which is not exactly what you want to know. Also, depending on the size distribution of your library, one can get different results. If your qPCR standards are not the same length as your library...you will get "off" results.

One last point about qPCR standards: If you didn't make the standards yourself and test them on a flowcell, you will not be sure how a relative concentration, derived from the standards, will cluster. When we make standards we perform library prep, amplify the libraries, then qPCR with an existing set of standards (for which we know how it clusters). We always keep our standards rolling...as it were.

[Genquest]

qPCR Library Quantification

Hi,
I am back with qPCR related problems.It seems my qPCR related issues are not going to end at all.
Recently,I encountered some very contradictory results!!The concentration of standard control (in this case PhiX) turned out to be half of the original concentration.This screwed up the concentration of all libraries.Could not figure out what was the problem..as the only difference was that all the Illumina PhiX (10nm stocks) were pooled in and then diluted into 2nM and made into smaller aliquots.I got same concentration from qPCR in 3-4 subsequent experiments.
I am using the 1:10 dilutions for qPCR standards.Everything seems to be correct..I am getting good R^2 value i.e. >0.998 with good Efficiency i.e.. above 90%.I am unable to figure out what could have resulted the 50% decrease in conc.

Please help!!
Thanks in advance..

[HESmith]

DNA adheres to most plastic tubes, and this becomes an issue at low concentrations. Adding detergent (e.g., 0.1% Tween-20) or using low-binding tubes solves the problem.

[shawpa]

Hello everyone,
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.

Thanks in advance

I would suggest not to use the PhiX control. Initially we were told by illumina that we could use it. We got very bad quantification discrepancies resulting in a few crazy flowcell runs. Illumina later told us that the 10nM concentration of the PhiX control wasn't "exact" enough for qPCR purposes. Now we use the kapa illumina kit as suggested. We get better results and see less discrepancies between quantification methods. Either way, I always feel that loading a flowcell to the right dilution is an "adventure in clustering", so I don't get to worked up about it when it doesn't go exactly as planned

[SeqR&D]

Genquest

Hi,

My initial thought is...if you are exactly 50% off...maybe you diluted wrong. If you were to see a gradual decrease in concentration, then I would say that your dilutions are getting old. Tubes are sticky, but I have had dilutions around for a while and have not had a problem.

1. Have you had successful cluster generation before?
2. How do you know it is half the concentration you think it is?
3. IF you are seeing this as cluster density, just cluster at a higher concentration, and as long as your standards do not "drift" then you should be fine.

I have never used the Illumina PhiX control for qPCR quanting. What we did was to reverse engineer based on cluster density. We amplified 300bp library, purified, quantified using qubit or bioanalyzer, generated qPCR standards based on those quants, seeded a flowcell at increasing concentrations based on the qPCR results, picked concentration that gave us the cluster density we liked, then used that qPCR standard to quant the rest of our samples...that we seeded at the particular concentration we liked. I would assume that I do not cluster at the same concentration as anyone else, but I cluster where it works for me given the standard I use. Then, when you are running low on Standard, you PCR more 300bp library, and quantify using the first qPCR standard and make dilutions based on that...this way, you continue in a reproducible way.

[Genquest]

Thank you for your replies!!I appreciate...
First even I thought,my dilutions must have gone wrong so I repeated the whole experiment and got the same concentration for PhiX positive control as the first.I have repeated 3-4 times and I cannot get wrong each time.
PhiX positive control is diluted in the same manner as i would dilute my sample library.I was performing these experiments as suggested in Kapa Library Quantification kits for illumina platform(1:10 dilutions).I use PhiX as standard instead of Kapa standard(as my Kapa stnd. kit was out of stock).I was getting consistent results (i.e. around 2nM)with the same protocol until the aliquots were made!! I also observed a shift in Ct values(late by 1-2 Ct).
Then,i was suggested to repeat the expt. with Illumina protocol i.e. 1:2 dilution as well as Kapa protocol (1:10 dilution) in the same plate for comparative analysis.
And to my surprise,the 1:2 dilution gave the right concentration of 2nM while 1:10 dilution was again consistent at 1nM.I did not know how to interpret these results.
Can two different pattern of dilutions amplify differently?
The aliquots were made in 0.1% Tween 20 but i am not sure about the tubes whether it is Lobind or not(There are no such specifications made).
This made me think that there could be some DNA adherence,so the best thing I could think of was to vortex 2-3 tubes of 2nM PhiX aliquots vigorously for longer time and spin down..aliquot them in lobind tube and set up an assay again to see any change in concentration.And I could see the difference..the concentration rose upto 1.6-1.7nM.
1.Does this mean that DNA is showing some adherence?
2.If yes then why wasn't it detected in 1:2 dilution pattern?
3. Am I correct in vortexing and spinning down the DNA for prolonged time,will this make any difference?
I have been unable to find some satisfying answers so look upon you guys for some expert and experienced opinions!!
Thank you in advance...

[Genquest]

I would suggest not to use the PhiX control. Initially we were told by illumina that we could use it. We got very bad quantification discrepancies resulting in a few crazy flowcell runs. Illumina later told us that the 10nM concentration of the PhiX control wasn't "exact" enough for qPCR purposes. Now we use the kapa illumina kit as suggested. We get better results and see less discrepancies between quantification methods. Either way, I always feel that loading a flowcell to the right dilution is an "adventure in clustering", so I don't get to worked up about it when it doesn't go exactly as planned

I absolutely agree with your opinion because each lot of PhiX did show difference in concentration by 20-30% which matters a lot when you have to cluster on this basis.
We had also run a bianalyzer on phiX v3 which showed a wide range of insert length i.e from 440-630bp.Due to all these reasons we decided to pool down all the 10nM stocks into one, quantitate it by pico green and then dilute it to 2nM and make small aliquots to rule out the variations and also the minimize the freeze n thaw cycles.
But only after this I am encountering the qPCR issues,still not sure what is the matter!!