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  • which qPCR to use for quantitating lib?

    Hi,

    We want to do qPCR quantitating lib. So for the different kinds that I am aware of:

    SYBR gree qPCR (From micrograms to picograms: quantitative PCR reduces the material demands of high-throughput sequencing)

    digital PCR (Digital PCR provides sensitive and absolute calibration for high throughput sequencing)

    Taqman qPCR (A large genome center's improvements to the Illumina sequencing system)

    Taqman MGB qPCR (Titration-free massively parallel pyrosequencing using trace amounts of starting material)

    We are considering KAPA qPCR kit. Anyone uses it and can you share your experience - how much input lib is needed (pmol or copy-per-bead ratio)? Or I have to use one good lib that run well previously as guidance? Do you need to have different 'good lib' for different samples (plant, virus, plasmid, etc)?

    Any thoughts appreciated!

  • #2
    Hi,

    I'm wanting to pursue this as well to replace titrations for 454. Has anyone used any of these with the Rapid Libraries or Titanium amplicons? Also, what approach do you take to do this in place of titrations? Do you need to run some libraries with both methods first to sort of correlate the results? I'm not quite sure how the qPCR results indicate how much DNA to put into the emPCR reactions.

    Thanks!

    Comment


    • #3
      Other threads in this forum had mentioned that the Rapid libraries contain some hairpin, by which PCR amplification is somewhat inhibited. The 4th paper (Taqman MGB qPCR paper) showed AB-library and Y-library, both of them contain hairpin. The same hairpin exists also in illumina Y library.

      I would appreciate someone uses qPCR for Rapid libraries share some experience. Or some reps from KAPA gives some instructions on how to "Eliminate time-consuming and expensive titrations", as I see the ad everytime I log on seqAnswers?

      Comment


      • #4
        Hi everyone,

        I will try to address each point in the thread in chronological order.

        First, this from seqAll (and relevant to both subsequent posts):

        "We are considering KAPA qPCR kit. Anyone uses it and can you share your experience - how much input lib is needed (pmol or copy-per-bead ratio)? Or I have to use one good lib that run well previously as guidance? Do you need to have different 'good lib' for different samples (plant, virus, plasmid, etc)?"

        In general, we have found that there is some variation in optimal input library quantity with respect to: laboratory, sample type, library construction method, etc. For this reason, we suggest that qPCR is most usefully seen (and used) as a "relative" (within each laboratory) measurement tool, rather than as a method for determining an absolute, "universally true" amount of input library material for optimal emPCR or cluster amplification. Furthermore, each NGS technology/platform/sample prep evolves constantly and rapidly, and users seem to prefer a suprisingly wide variety of enrichment percentages or cluster densities.

        The short/conservative answer is therefore more or less as you suggest: use one or more "good (typical) libraries" as guidance to "calibrate" your qPCR results initially. You may require different "good libraries" for different samples or library construction methods (but you may not; experience will tell). You should only have to do this initially; thereafter, relative quantification by qPCR should provide a very good measure for any further optimisation that you might need to do.

        A longer explanation follows:

        Unfortunately, there are a number of factors that apparently confound the simple and reliable translation of accurate library quantification into optimal emPCR or cluster amplification. Most NGS users are aware of the (variable) difference between the total amount of DNA and the number of viable (or "effective") library molecules (i.e. those capable of initiating emPCR or cluster amplification). This is probably the major reason why methods such as electrophoresis and spectrophotometry often fail to produce reliable results, because they cannot distinguish between library molecules carrying the proper combination of adaptor sequences, and other DNA fragments that do not. For this reason, qPCR almost always yields lower values for library concentrations than methods such as Bioanalyzer assays or spectrophotometry (and one therefore needs to use correspondingly "lower" concentrations, as determined by qPCR, than one is accustomed to using when quantification is by another method).

        Apart from adaptor ligation, other factors such as GC-content, secondary structures, fragment length, and DNA damage (e.g. from UV exposure during gel purification) can affect the efficiency with which library fragments are converted into functional "polonies" during emPCR or cluster amplification. Unfortunately, qPCR and emPCR/cluster amplification are probably not equally vulnerable to these factors. In fact, because they occur on a solid support, both emPCR and cluster amplification (or "bridge PCR"; bPCR) are much less efficient than standard solution-phase PCR (i.e. qPCR), so it seems reasonable that they may be more vulnerable to failure as a result of various types of "difficult template".

        Finally, there is undoubtedly some inherent variability in the reagents and processes of emPCR/enrichment and cluster amplification. Thus, even when using the exact same library prep, people sometimes see significant variation in their results, which is independent of library quantification.

        Another reason for using a reference library initially to "calibrate" qPCR results to determine optimal input is that enrichment and cluster density do not increase linearly with increasing template input. The TaqMan MGB paper contains a good discussion of this, but I am not sure how well the theory translates to reality across a wide variety of laboratories, samples, etc.

        In general, we have found that when qPCR is used for quantifying 454 libraries (standard FLX or Ti), then the optimal cpb is somewhere around 5 - 10 times less than when quantification is by Bioanalyzer or spectrophotometry. In other words, the range of optimal cpb would be 0.1-1 cpb, rather than 1-10 cpb. We do not yet have enough feedback from customers using the 454 Roche Rapid Library Preparation kits to confirm that they behave in the same way. My understanding is that adaptor ligation with the new Rapid Library Prep kits should be more efficient, and therefore one might expect a smaller discrepancy.

        Most of our customers have found that ~10 pM input (determined by qPCR) results in ~220k clusters/tile on the Illumina GA platform, and this seems to hold for most gDNA libraries as well as RNA-seq libraries. On the other hand, some labs see up to a 2-fold difference in the optimal amount of input material for their gDNA libraries compared to their RNA-seq libraries. We therefore recommend that customers determine the optimal input within the context of their sample prep and cluster amplification protocols. Many Illumina customers now seem to be aiming for much higher cluster densities (up to 320k clusters/tile?), so this is a moving target.

        In summary, whether or not you are able to entirely do away with titrations (emPCR or cluster amplification) will depend on the diversity of samples and library prep procedures in your lab, as well as the degree of control that you are able to excercise over the workflows. We have high-throughput customers who use automated liquid-handling stations for most of their sample prep, library prep, and library quant, and they simply cannot practically perform amplification titrations with all of their samples. Because of the consistency/reliability of their workflows, however, qPCR has allowed them to reduce the variability in cluster density that they struggled with previously, and they have done away with titrations entirely. On the other hand, core facilities that accept samples and/or libraries from a variety of customers seem to find it much more difficult to control variability. Nevertheless, qPCR provides them with a very valuable tool for reducing the number of titrations that they need to do. While qPCR is not a perfect, cure-all solution for library quantification, it is a lot better than the alternatives, in many ways, not least of which because it reduces the variability in translating concentration into emPCR enrichment or cluster density.


        Second, in response to this from RCJK:

        "Has anyone used any of these with the Rapid Libraries or Titanium amplicons?"

        Unfortunately, as you are probably aware, there are some confusing issues with the adaptor sequences used in the various 454 library prep kits.

        We supply two kits -- one for the "FLX" adaptor sequences, and the other for the "Titanium" adaptor sequences. To our knowledge, our FLX quant kit is compatible with all FLX libraries. Confusingly, Roche 454 has used the FLX adaptor sequences in their Titanium amplicon libraries, so if you need to quantify Titanium amplicon libraries then you would need to use our FLX quant kit. However, the situation with 454 Titanium amplicon libraries seems to be further complicated, depending on whether you want to do unidirectional or bidirectional sequencing. Apparently the standard amplicon protocol uses so-called "Lib-A" Fusion Primers and Lib-A emPCR kits to allow bidirectional sequencing, but some people prefer to use "Lib-L" Fusion Primers and Lib-L emPCR kits if they are sure that they only want to do unidirectional sequencing of their amplicons. The Titanium "Lib-L" compatible Fusion Primers that Roche 454 recommends are compatible with our "Titanium" library quantification kit, and not with our "FLX" library quantification kit.

        Our Titanium quant kit is compatible with the Titanium Rapid Prep libraries, but we suggest that customers use a longer combined annealing/extension time in the cycling protocol to accommodate the longer fragment sizes that are usually used with these libraries.

        Third, seqAll raises this point:

        "Other threads in this forum had mentioned that the Rapid libraries contain some hairpin, by which PCR amplification is somewhat inhibited. The 4th paper (Taqman MGB qPCR paper) showed AB-library and Y-library, both of them contain hairpin. The same hairpin exists also in illumina Y library"

        This issue of "hairpin" structures that is discussed in the "TaqMan MGB paper" relates to the question of so-called "PCR suppression" effects that one sees with PCR templates that carry complementary sequences at either end. This occurs in library construction whenever you have a library fragment carrying the same adaptor sequence at both ends -- so-called "A-A" or "B-B" (also "P1-P1" or "P2-P2") fragments. In theory, such library fragments are amplified with very low efficiency (if at all), because the intra-molecular binding of complementary sequences at either end of the target efficiently out-competes binding by primers, due to both kinetics and thermodynamic stability. Thus, PCR should result in the enrichment of the desired "A-B" (or "P1-P2") library fragments. Similarly, qPCR should not quantify ("count") A-A or B-B fragments, since they will not be efficiently amplified during qPCR. This is a benefit, because A-A and B-B fragments are essentially inert in emPCR (or cluster amplification).

        Of course, one would expect much fewer, if any, such fragments in the Y-adaptor type schemes used for Illumina library construction, and in the new 454 Rapid Prep libraries.

        Comment


        • #5
          Thanks a million for the detail explanation! Very informative, cover many aspects from lab persons' perspective!

          And, very useful to know "the range of optimal cpb would be 0.1-1 cpb, rather than 1-10 cpb" and "~10 pM input (determined by qPCR) results in ~220k clusters/tile on the Illumina GA platform"!

          Comment


          • #6
            I have heard that Illumina's cBot generates more clusters per tile from the same input concentration of DNA than the cluster station? Can anyone else comment on this?

            Comment

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