[Genquest]

I too agree with SeqR&D,that would be the best method for clustering.I have also used Kapa illumina Kit and this too gives consistent results but they are so expensive,have to manage with PhiX until I get next one.
And these PhiX standards introduce so much variation..Dont know how to get it right!!

[hawaii454-0]

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

What's the difference between the Illumina phiX control and the Kapa Illumina phiX?

Anna.

[shawpa]

What's the difference between the Illumina phiX control and the Kapa Illumina phiX?

Anna.

I don't know what the "standards" are in the kapa library quantification kit. If you say they are phiX then they probably are. However, they are already diluted and you just pipette them into the wells (full proof). I assume each standard is heavily QC'd by kapa (or whoever owns them now). I am not sure if the phiX control that you can order from illumina is QC'd to the same standard. I haven't had any issues with the kapa kit since using it and we use it a lot.

[Genquest]

Hi Shawpa,

Will the PhiX still show variation in qPCR,even if we have pooled all the lots of PhiX provided by Illumina and then the concentration adjusted to 2nM with the Picogreen?We have made small aliquots of this pooled 2nM stock of PhiX.

We are still facing variation with the PhiX concentration in qPCR@20-35% range.Not sure if this is acceptable or not.Have anyone seen so much of variation in PhiX with every assay that is set up?

Now Regarding the previous problem(concentration 50% off) that I was facing,it got resolved.I just changed to different tips (provided by another vendor).
Now this made me think,don't know how much to rely on the different type of tips that we are using!!
Any suggestions on tips?

[shawpa]

Hi Shawpa,

Will the PhiX still show variation in qPCR,even if we have pooled all the lots of PhiX provided by Illumina and then the concentration adjusted to 2nM with the Picogreen?We have made small aliquots of this pooled 2nM stock of PhiX.

We are still facing variation with the PhiX concentration in qPCR@20-35% range.Not sure if this is acceptable or not.Have anyone seen so much of variation in PhiX with every assay that is set up?

Now Regarding the previous problem(concentration 50% off) that I was facing,it got resolved.I just changed to different tips (provided by another vendor).
Now this made me think,don't know how much to rely on the different type of tips that we are using!!
Any suggestions on tips?

I can't give you any further advice on the phiX. If what you are doing works and you get good clustering from it, then I would keep doing what you are doing. All I can say is that we use the kit. I guess it is expensive (?), but if you are spending a few thousand on sequencing and the related reagents then the cost for a quantitation kit really is minimal. Good luck on your your qPCR. By the way, we use barrier low retention tips. We use either Sharp from denville or Avant Guard tips from midsci.

[Genquest]

Thanks shawpa

[NGCMRK]

Many reserchers started using Kapa Library Quantification Kit. qPCR is good way to quantify, and Kapa LQ comes with the validated standard, which is very helpful.

[DNA_Dan]

+1 on the Kapa kit. All these methods are relative to some standard. Once you know what factor produces the desired clusters, the rest of the game is just consistency. We've started usuing fixed volume pipettors for doing the qPCR dilutions, then use the same pipette when going back to the library stock for the run. Another main source of variation we have identified is library size distribution. The best way to assess this is to use the "region" function in the bioanalyzer software to get the average size under the curve for the whole region you identify. Don't just take the highest peak average for the distribution. In libraries with "leaning" distributions, this is critical to get an accurate read on the concentration. In non-size-selected libraries, you're best off titrating the sample and hoping future samples follow the same distribution, otherwise it's a crapshoot every time.

[Christopher Odom]

What's the difference between the Illumina phiX control and the Kapa Illumina phiX?

Anna.

I don't know what the "standards" are in the kapa library quantification kit. If you say they are phiX then they probably are. However, they are already diluted and you just pipette them into the wells (full proof). I assume each standard is heavily QC'd by kapa (or whoever owns them now). I am not sure if the phiX control that you can order from illumina is QC'd to the same standard. I haven't had any issues with the kapa kit since using it and we use it a lot.

Great question!

Actually the DNA Standards used in the KAPA Library Quantification Kits aren't PhiX, nor are they sequencing libraries. Instead, we use a defined, pure, linear, dsDNA amplicon for each set of DNA Standards. This allows us to rigorously validate their efficiency and reproducibility for use as qPCR amplification standards. Because PhiX is a library, it is very difficult to manufacture reproducibly through multiple production lots and over extended periods of time.

Before accepting a newly manufactured lot into our inventory, we use a stringent qPCR assay to compare each new lot of KAPA Library Quantification DNA standards to a reference set of standards during manufacturing and quality control. We compare Ct scores for each standard in a newly manufactured set with Ct scores in a reference set of standards, and we ensure that each standard lies within 0.1 Ct of the respective reference standard, and that the resulting standard curve lies on top of the reference standard curve.

I hope this is helpful and thanks for the positive feedback!

Chris Odom
KAPA Biosystems
Technical Support

[Genquest]

Thank you all for your informative repliesAnd please keep posting with new problems,so that we can learn from them.
And Thank you,Christopher Odom,very insightful!!

[1230Rock]

Hello,

I have been quantifiying my libraries with the Kapa system and find that I have to dilute my libraries 1:16000 at least in order to get in a reliable range of the standard curve. Also, my calculated molarity is 3-5 times higher than what I get with a Nanodrop. Any insight into what might be going on would be greatly appreciated!

Thanks in advance.

[GW_OK]

I serially dilute my libraries to 1:10,000, 1:100,000, and 1:1,000,000 to fall within the standard curve (and run all three). Not a big deal. And I never trust anything a Nanodrop says.

[DNA_Dan]

I'm with GW_OK. A nanadrop really has no business in NGS as far as I am concerned. It measures everything, whereas the Kapa kit is measuring only "functional" molecules that will actually contribute to amplification. That is why you see a difference. If you could imagine, there is unligated product, single end ligated product, etc. which will contribute to A260, but not actually be functional to amplify in the Kapa kit (or on the slide).

I am typically at 1:50,000 dilution for my libraries. I used to run different dilutions, but have seen up to 20% difference in concentration due to the added dilution steps. The measurement is more accurate on Ct the further out you go, but reproducing that exact same dilution is questionable and has a higher deviation. I have found it best to focus on the reproducibility of technique, using fixed pipettors, watching whether I "blow-out" on the pipettes, etc. After all, the hope is that what you do at the bench and calculate for the Kapa kit is reproducible when going back to your stock library tube. Doing it EXACTLY the same way every time is the key to repeated consistency.

[protist]

I echo GW_OK & DNA_DAN with regard to the nanodrop - should not be relied on for any library quantification at any stage. My advice if you want to quantify outside of your KAPA protocol use a QUBIT it is more sensitive and consistent than the nanodrop.

[DaanV]

I used to run different dilutions, but have seen up to 20% difference in concentration due to the added dilution steps. The measurement is more accurate on Ct the further out you go, but reproducing that exact same dilution is questionable and has a higher deviation.

Just figured I would hop in in here to make my first ever post on this forum after a long time of lurking and learning (thanks for that all of you!).

I have found that indeed, you calculate different starting quantities (SQ) for different dilutions of the same sample. However, it should be kept in mind that SQ is calculated based on the PCR-efficiency of the standard.. and not all libraries run through the reaction at the same efficiency.

Now one might argue that the differences in efficiency between the standard and the sample are relatively small (most of the time <5 percent-points difference) and can thus be neglected. I would like to stress though that the difference between 95% efficiency and 98% efficiency is not negligible after 20 cycles of qPCR (typical Ct score for a heavily diluted sample) due to the exponential nature of PCR: ((0.95+1)/(0.98+1))^20 = 0.74. I.e. if your standard has a 98% efficiency, and the sample has 95% efficiency, your calculated SQ value can be up to 25% off!

For this reason, I always run duplicates of three dilutions (1.000x, 16.000x and 256.000x) for every sample and calculate the efficiency of each sample, then use this efficiency to make a more accurate estimation of the SQ. Using this method, the results for each subsequent dilution are much more consistent than when simply using the SQ values as calculated by the qPCR software (standard deviation of the 6 values is almost always <10% of SQ using my method).

Of course this does mean that I'm limited to 13-14 samples per plate of qPCR, but I find that this is only a minor investment compared to the improved accuracy.

I hope this helps some people out.