I have been using the serial dilution method of analyzing the efficiency of Primers designed, via Standard curve. For the assay I take mixture of variety of cDNA's but every time the efficiency value turns out to be different let's say at first it was 85% , next time the percentage turned up in lakhs and even larger than that.I have attached the Standard curve of a primers for reference. Why is this happening could anyone explain?
If I'm reading this correctly, your Cq values decrease by fractional values even following substantial dilution (Cq 23.15, then diluted 1/1000 = Cq 23.8).
This is, in essence, telling you your standard curve hasn't worked, at all.
The variation in Cq values over this range is also good support for this. There is barely any correlation between dilution and Cq (the R2 for dilution series should usually be 0.99+).
If a standard curve is working correctly, you should see an increase in Cq of ~3.32 cycles for every ten-fold dilution (gradient of -3.32).
If I'm honest, what you're most likely measuring here is either primer dimers, mispriming or genomic DNA contamination. Or contaminated stocks.
So: does your PCR actually work? Does it produce a single amplicon, as determined by melt curve and subsequent gel electrophoresis? Does it NOT produce this amplicon if you omit cDNA from the reaction (-ve ctrl)?
Further to this, if you have two samples of cDNA, one of which should contain your target and one of which should not, do you get appropriate Cq values for these? (low Cq values for +ve, high Cq or no amplification for the other?).
Once you've confident your PCR is actually working, we can move onto troubleshooting the standard curve.
To get a standard curve which is also termed as absolute quantification, serially dilute your samples over a wide range. MIQE guidelines suggest that the standard curve should be prominent over atleast three powers of concentration (ex. 10^-1, 10^-2, 10^-3 or 1, 10 100 etc) Then you can calculate the standard curve and efficiency after plotting the graph of log of concentration vs Ct values.
If you are using mixture of cDNA and the respective gene is differentially expressed at different stages, make sure you use a cell stage with maximum expression for analysis.
Additionally, efficiency curve will hint you the concentrations of target gene which the primers can detect.
Henry Kolge Thankyou sir for your explanation I would surely look for my mixture of cDNA and yes I am using serial dilution approach 1:5,1:25,1:125 & 1:625.Should I increase this range ?
John Hildyard Thankyou so much sir for such an elaborate reply.You pointed out correctly that there might be some fault in my cDNA since there is no fold increase in Cq values.. even I wondered that. For standard curve I use several mixture of cDNA samples of the same cell line lets say undifferentiated, spontaneously differentiated etc.,stages .So can you tell me whether this approach is correct or do you suggest some alterations. Awaiting for your kind reply!
Honestly, I don't bother using cDNA for my standard curves.
If you think about the actual PCR reaction, after the first two or three cycles the majority of target is going to be a PCR product (rather than cDNA), and after another three or four cycles basically all of it is.
So I just use a PCR product.
I run my PCR (can be qPCR or just regular) and then run it on a gel to confirm it's a single band of the right size. I cut this band out, clean it up and then spec it, and (using the amplicon length and measured concentration) determine the number of molecules per ul.
I then use this in a serial dilution series from say...1e9 molecules per well down to maybe 1e-1, and establish the linear range (and ideally the LLOD and LLOQ, too). And from the gradient I can establish the efficiency.
In practice, what I usually do is: for a given GOI, I download the ensembl transcript sequence and look for large introns. I then design my primers to the exons either side (the idea being that now you will only amplify cDNA, not gDNA). I use primer3 for this.
I usually order two or three primer pairs for each gene, on the assumption that one will probably be rubbish, for...eh, reasons. I try out all the pairs with a cDNA sample I know should be robustly expressing my target, and then pick the pair that works best and throw the others away. I never bother to tweak annealing temps or anything: if it doesn't work under my bog-standard one-size-fits-all PCR protocol, then I don't use it.
Primers are cheap, your time is not.
THEN I move onto efficiency shenanigans.
This approach allows you to establish the basic safety checks ("does my PCR work, does it amplify the right thing, is it specific, etc") before committing too much time and effort to the enterprise.
Concentrations are good, but you can make it five, either take the original concentration or go for 1:3125
Most folks will clone the region of interest into a plasmid, set up a restriction digest to linearize the plasmid, and use that to make their standards. It's a bit of work up front, but then it's relatively easy (and cheap!) to make as much DNA as you want for the standard curve.
Good luck!
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