Hello everyone,
I am trying to run a qPCR for detecting a unique group of archaea using specific primers which I have tested both in silico and in vitro. The samples I am using seem to have low copy numbers of these microbes (~100-1000 copies). When I run qPCR for 40 cycles, I seem to have unspecific product amplification (checked on a gel as it only showed as one peak after melting curve analysis). My samples give a Ct value of 24-25. My last standard (I run 6, 10 fold diluted standards) has a Ct value of 28 (theoretically 100 copies) whereas my negative control usually has unspecific amplification (~Ct value 33-35) (usually a product not visible on gel and about 10-15°C lower on Tm than my target). Thus I ran a few tests by changing some PCR parameters (decreasing annealing time and amplification time by half) and limiting cycle number to 35. When I check the outcome on a gel I dont see the unspecific amplification anymore. So I decided to run qPCR with these new amplification conditions and I observed similar Ct values for standard and samples. However my last standard and my samples just reach a plateau (not complete) on the plot: Delta Rn vs cycle number. Is this evaluation correct and viable? I was thinking that since all my samples and standards are amplified before 35 cylces and have a Ct values between 11 and 29 this should not be a problem. Any help, advice, criticism would be gladly welcomed.
Thanks in advance.
Regards,
Ajinkya
@Muthanna Hadi Hussain: Any reasons why that you recommend that? Just curious!
Hello...
If you have a negative control that "usually has unspecific amplification" (Ct value 33-35 of 40 cycles) this qPCR is NOT especific. You have to think in what contain your negative control (You don´t specify that... is water? soil?). Think that if you have some organism "not target" in your negative control with low title, you obtain hight Ct values, but when you test this qPCR with other sample and casually this "not target" organism appear in hight title, you will obtain low Ct values-->Problem.
With this explanation i have to say that if you are limiting cycle number to 35, you are not solving the problem, you're just hiding it.
In my opinion, if you are designing a new qPCR protocol... this could be a problem. I recomend you find out what are crossing with you primers and then try desing others primers and/or other probes that not inlude this not target organism.
regards
@Felix: Firstly, thanks for your useful insight.
The Ct values of my negative control i.e. just PCR water ranges from 33-35. Just to clarify, I run 7 standard dilutions, but I use only 5 to 6 for quantification as the 7th standard has a Ct value of ~31-32 and theoretically has only 10 copies. I find it a bit unreliable to rely on it. The negative control is even above this Ct value with values of 33-35 with a very hifh S.D. The SYBR fluorescence barely crosses the background ROX fluorescence in the negative control and should theoretically have less than 10 copies. So I think this should not affect my quantification and this probably happens due to aerosols formed during pipetting and vortexing of the DNA.
I said its unspecific because the product I get in the negative control is usually having a Tm of 65-70°C and the target Tm is 86°C. Furthermore, this product is not even detectable on gel. Finally, I have already tested primer specificity in vitro (using various clones from neighboring genus's and families of the target organism and testing on gradient PCRs) and in silico. I think I can still run 40 cycles and have the same outcome (since I optimized the qPCR to not give unspecific products), but I was just wondering if it was correct to run 35 cycles or not.
Just attaching a figure of the five standards, my samples and the negative control in red towards the end.
PS: My samples are marine sediments
OK!! That clarify more things!!
If the negative control is water, that is a contamination, as consequences of what you have mentioned (aerosol...). In this case Ii think that you can still run 40 cycles and have the same outcome (being careful) ;)
So, don´t worry and try again.
Good luck.
Yes, you are correct to limit the program to 35 cycles. More than that will start to create artifacts. have you tried increasing the amount of template for your low copy number samples? You can correct for the increased starting material in just those wells in your analysis.
@Artur: Thanks for your thoughts. Currently I have an efficiency of ~90% so I am working on improving it to at least 90 :) Will also test if I get no non specific products after 40 cycles, just in case.
@Katie: I did try increasing the template but it usually inhibits the amplification because a) either there is too much DNA or b) the inhibitor concentration increases. I have to start with high amounts of DNA from the get go as the target population is not so enriched in the marine sediments and I am targetting functional genes which in turn are one copy/cell.
I cannot avoid inhibitors because they co precipitate from the sediment itself and we dont know much about them yet to improve our DNA extraction protocol
Just wanted to give an update.
With lowering annealing time to 20s from 30s and amplification time from 40s to 20s and assessing efficiency for new conditions with new primer concentrations (-90%, R2=0.998) I was able to get a single quantifiable product at 35 cycles with no by products. Hopefully such kind of adjustments can help others too :)
PS:Testing for PCR amplification of functional genes with ApliTaq polymerase wasnt yielding products although with other polymerases it did (Just FYI)
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