this is likely a very easy question, i have just been confused by everything i have been reading. i am looking at gene expression over time of two genes in a fungus, and i have two reference genes, actin A and B. i have never run qPCR, and it is time to do it with my calibrator sample for the first time, how do i set the plate up?
i know i need a serial dilution to generate a standard curve for each of the genes and their respective primer sets, which would take up an enormous number of wells if i do a serial dilution in triplicate for each of the 4 genes, is this what i should do?
do i need to test different primer concentrations at this point, or can i just try one for the first run? likewise, i think i have an appropriate annealing temp to try. what else goes on the plate, a triplicate negative control without cDNA?
thank you for the help. i am running this with BioRad iQ SYBR Green Supermix.
Try primers concentration first on a few known samples - preferably standards, to see which concentration is the best - if you work with SYBR Green (as you say) choose this primers concentration which will give you good amplification curve but with minimal primer dimers - this you'll see after melting curve. Make all necessary normalization reactions on a few known samples - together with optimum magnesium concentration (if you have it already in premix you don't have to) or annealing temperature.
Having this done you'll have to run your standards in dilutions and replicates - unfortunately this will take many wells and a lot of supermix, but that must be done.
For my last standard curve I used six replicates, in many publications I read there are from three to five replicates used per each dilution. Remember to use such dilutions that will cover all ranges of your possible cDNA concentrations you'll later test with this standard curve.
But now I've got a question - if it's gene expression you actually perform relative quantification? With callibrator or housekeeping gene of known concentration you don't actually need to construct a standard curve but you determine concentration of your target gene on the basis of concentration of a housekeeping gene. Unless you use absolute quantification which relies on a standard curve. You can always use standard curve to determine detection limit of your assay.
If you need a standard curve three replicates of each dilution is minimum for one target. You also need no template samples preferably in the same number of replicates.
This is the basis. I generally run with my samples negative samples as well - these are samples with different DNA than my target (to check specificity). When I start to develop an assay I run also different reactions but this is a different issue.
I hope I could help you a bit.
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Thanks so much for your response, it helps a lot. I am indeed doing relative quantification, but I was under the impression that to validate the primers I first need to generate a standard curve to be sure that efficiency is good. Do I not need to do this?
Once I do this, do I not need to run serial dilutions each time? Thanks
Yes, you can do this - this will indeed tell you what is the efficiency of your reaction and also detection limit if you use very low concentrations of your target DNA as a template.
Theoretically you don't have to do it in many replicates, but take into consideration you will not calculate standard deviation, variation coefficient and in result standard error between the replicates, so you won't know what possible error you have. And there are always some pippetting errors on the way.
This obviously depends on your budget, but remember this is your research and this is in your best interest to do it as good as possible. So if you aren't so restricted with money then do it in three replicates.
To see if the primers are good a standard curve is not needed so much. The efficiency of course depends on your primers but to lower extent than for example template concentration and purity and also your target DNA - number of bp, CG content, CG nucleotides arrangement (these are the factors that affect polymerase efficiency).
You will have to know the efficiency of your PCR for your further calculations anyway, so it's better to a standard curve using replicates.
Wish you good luck!
Last edited by Aga; 02-15-2009 at 02:57 PM.
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Aga, you have been very helpful thank you. I am comfortable with my primer concentrations and annealing temp; I ran my first qPCR of my calibrator sample with 7 serial dilutions of cDNA for one of my four primer sets in triplicate and got efficiency of 100.4% with good R^2 and good melt-curve analysis. I will in the future include a no-template control in triplicate.
I will do the same thing with my calibrator sample for my three other primer sets to ensure good amplification efficiency and specificity, but then what?
As the fungus I am working with grows, I will extract RNA at several time points. I will then synthesize first-strand cDNA from this RNA, then I am still not sure how to set up the plate for the test samples.
Do I do a serial dilution of the unknown test sample, or just one triplicate reaction using undiluted cDNA, plus a no-template control? Basically what I don't understand is, I have been reading about data analysis for relative quantification, and the different methods describe using a Ct, but I am unclear as to which Ct to use. For the calibrator when I did a 7-fold serial dilution in triplicate, which Ct value would I use as the calibrator Ct (for a given gene), since of course each dilution has a different Ct. Would I use the undiluted Ct?
For each test sample plate, does it only require a single triplicate undiluted reaction for each primer set and no-template control for each primer set?
Thanks so much, I really am doing a lot of reading on qPCR but your clarification is invaluable.
Probably you've read some publications already - the one which I recommend for you is Pfaffl's 'A new mathematical model for relative quantification in real-time RT-PCR'
You'll find it here: [Only registered users see links. ]
If you don't have full access to this article let me know.
What is generall done is the following:
- you do a serial dilution of unknown sample - you need it to establish PCR efficiency, plus a no-template control (NTC) and CONTROL SAMPLE as well - a standard sample with known concentration; to be alble to calculate intra- and inter-assay variation you'll have to do it in triplicate.
- you'll have to calculate the relative quantification of your target genes using reference gene as a comparizon; the equation is given for egample by Pfaffl.
You need the following data:
efficiencies of PCR reactions for target samples (unknowns)
PCR efficiency in amplification of reference gene
CP values for unknown samples and reference genes
CP deviation for control sample
Usually it is advisable to check reaction efficiency for each tested unknown sample. Unfortunately even a single thing can influence reaction efficiency and your results will vary. If you discover your efficiencies are almost identical you can save some time and money. For routine analyses make dilutions of target and reference gene in duplicates to save reagents.