I ask this question because I have found that the PCR efficiencies for my housekeeping and target genes are nearly identical when the former reaction uses 1:1000 diluted cDNA and the latter reaction uses 1:100 diluted cDNA. I intend to use the comparative Ct method to quantify the relative expression between samples and controls, so this efficiency similarity is crucial.
I can't think of a way that a difference in dilution between target and HK gene would matter mathematically, but I could be overlooking something.
Well, no. I must disagree.
There is no need to have the same dilution for the target gene and for the reference gene. As long as the dilution factor is the same for both, treated and controls, the factor will cancel out from the ddCt.
Take the formulas, add a dilution factor, and calculate what the ddCt is.
From a logical argument: the referenece gene will surely not have the same concentration as the target gene. If you's use a different reference gene, it likely would have a different concentration again. If bot are valid reference genes then it should not matter which one you chose (ok, the best it to use both, but that's for argument here). If you can take on or the other, and their concentrations are anyway different, then you can use any of them in any arbitray dilution (again, given the dilution is the same between the conditions).
The problems start when the efficiencies of target and reference gene are not identical. Then larger dCts will increase the bias, but then everything gets weird in qPCR...
It's best if you use the same dilution. I'm not aware of anybody use different concentration for reference and target genes. Most people prefer to use the same dilution because delta delta ct (relative gene expression) depends on the concentration of the target and reference gene. It would be wise to use the same dilution since you will be using them for the calculation. Also, try to use multiple reference genes if you can. More and more publications indicate that previously known "house keeping" are not stable in all conditions as previously thought. That's why people no longer call reference gene - "house keeping gene." If you want to publish qRT-PCR result, you should follow the MIQE guidelines. Good luck.
I agree with the previous answer. As far as I know, to use the ddCt method the two pairs of primers must have a similar efficiency (no more than 5% difference) at the concentration range that your samples are going to fit in. It is a relative quantification method, and the house keeping gene is just there to be used for normalisation, so that you can compare different samples, which might have different concentrations (and thus by normalising them with the housekeeping gene, you can compare then them). Ideally, you would run the same sample with the two sets of primers on the same plate, so that all the conditions are the same. If your primers have different efficiencies at the concentrations where your samples are going to be, I would try and design other pairs of primers for the gene of interest and maybe even the housekeeping gene, and then use the two pairs that have the most similar efficiencies. Otherwise, you would have to go for an absolute quantification method, which I've never used but both methods are very well explained in help booklets from Life technologies, biorad and qiagen (all free online as far as I know). There is a paper that states the MIQE guidelines that you should check, just search for it on pubmed/google scholar, it was most useful to me. Good luck.
For q-PCR analysis you have to have same level of dilution of cDNA for both reference gene and target gene.
Thank you, everyone, for your answers. I will work towards improving the PCR efficiency in the reactions such that they similar in equivalent dilutions.
I will also begin to assess more reference genes for use as endogenous controls, as required by the MIQE guidelines. I have been following those guidelines as well as I could, aside from the use of a single reference gene.
of course you have to use the same dilution or how could you compare your results???
Well, no. I must disagree.
There is no need to have the same dilution for the target gene and for the reference gene. As long as the dilution factor is the same for both, treated and controls, the factor will cancel out from the ddCt.
Take the formulas, add a dilution factor, and calculate what the ddCt is.
From a logical argument: the referenece gene will surely not have the same concentration as the target gene. If you's use a different reference gene, it likely would have a different concentration again. If bot are valid reference genes then it should not matter which one you chose (ok, the best it to use both, but that's for argument here). If you can take on or the other, and their concentrations are anyway different, then you can use any of them in any arbitray dilution (again, given the dilution is the same between the conditions).
The problems start when the efficiencies of target and reference gene are not identical. Then larger dCts will increase the bias, but then everything gets weird in qPCR...
I have one ore question. When the cDNA synthesis is done, (let say 20ul of cDNA at the end) first I add 80ul ddH2O to the cDNA and then I make the dilutions for primer efficiency. Is that correct? Otherwise the amount of cDNA is not enough.
You would usually add 2 µl template to a PCR. Thus, for a dilution series you would need an intial volumen of 2 x 2 µl = 4 µl (2 µl are used as 1:1, the other 2 µl are serially diluted). This allows you to measure about 5 independent dilution series from 20 µl cDNA. If your assay uses only 1µl template the numbers of possible replicates will double appropriately.
It's always a bit probelmatic to use cDNA as standard, because the gene of interest may not be expressed at reasonable levels. I would prefer creating a standard by a preparative PCR. It bears the risk of carry-over contaminations, but this is manageable when you are careful. You will then have really sufficient amounts of standard to replicate as often as needed, and you are sure to get your standard curve starting with reasonably low Ct values so that you have a good chance to reliably determine the dynamic range of your assay.
Jochen Wilhelm I am sorry but I did not understand the procedure. Should I add lets say 80ul ddH2O to my cDNA and then proceed to serial dilutions.
Btw. is it necessary to check efficiency for each primer pairs no matter if its a gene of interest or house keeping. Thanks
If you add the water, your lowest Ct will be about 2-3 cycles higher, what may be a problem if your lowest ct is already quite large. It is not a problem, when your lowest ct is low (about 18, to give a rough number).
And yes, you must validate all assays you are using - this means all the primer pairs. That's not something limited to real-time PCR... you should always make sure that a quantification is unbiased, and if it is, you should know the bias and consider it in the interpretation.
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