don't get stuck on measuring cDNA conc.

First, you can't really measure cDNA the way most people do. If you use a nanodrop (or any other spectrophotometer based assay) after cDNA synthesis, you are not measuring cDNA but a mixture of RNAs, cDNA, and nucleotides, and without any information on degradation or quality. If you'd want, you could purify cDNA and measure, but that's totally irrelevant and not needed for what you do, so don't bother. Second, since you run a QPCR amplification, ANY amount of cDNA will work, as long as it's not as low as a "homeopathic dilution" and not so high that you get inhibition of amplification due to.. other non-cDNA contaminants that are carried over from impure RNA or components from your cDNA synthesis etc. As long as it works, and your ct values are let's say

run the experiments taking diff. conc of cDNA ie. 10ng, 25ng, 50ng and 100ng check the Ct values and decide the conc. For us Rat liver cDNA we took 25 ng, it varies by the tissue. better you run the experiment by taking diff.conc. good luck

PCR is a very powerful Technique. I use my cDNA in pg. and find beautiful amplifications. But the key to success is a quality RNA, good reverse transcription, and specific primer designing (considering GC content, Tm, and complimentarity). I would also suggest you to read this article on proper guideline for qPCR (MIQE): http://www.ncbi.nlm.nih.gov/pubmed/19246619

Try some options in a plate: different cDNA concentrations (the above examples are good) combined with different primer concentrations. The cDNA concentration range might vary according with your cDNA souce/transcription kit/cDNA purity/cDNA integrity/RNA extraction; even, wether you use SybrGreen or TaqMan might vary. You can find some range references from the PCR reagents manufacturer instructions book (if not, check the PubMed: articles that match with your sample that performed real time pcr with your reagents).

Later, you should choose your best efficacy result: use the slope of the regression line (for each primer) to decide it.

Good luck!!

Kindly find attached this reference that may help

don't get stuck on measuring cDNA conc.

First, you can't really measure cDNA the way most people do. If you use a nanodrop (or any other spectrophotometer based assay) after cDNA synthesis, you are not measuring cDNA but a mixture of RNAs, cDNA, and nucleotides, and without any information on degradation or quality. If you'd want, you could purify cDNA and measure, but that's totally irrelevant and not needed for what you do, so don't bother. Second, since you run a QPCR amplification, ANY amount of cDNA will work, as long as it's not as low as a "homeopathic dilution" and not so high that you get inhibition of amplification due to.. other non-cDNA contaminants that are carried over from impure RNA or components from your cDNA synthesis etc. As long as it works, and your ct values are let's say

I agree with Chris, if you really want to know the concentration of your cDNA you probably need to use more fancy instruments such as bioanalyzer or a qubit fluorometric quantitation. However, the concentration of the cDNA will vary among samples, because even when we use same RNA concentration, there are several other factors that will determine the final concentration of the cDNA.

The proper way to setup real-time experiments is first evaluate the primers and find the limits of their performance. Usually people use PCR product but the result is different when you use a pool of cDNA, so I recommend using real cDNA rather than a purified product. What you can do is test serial dilution of cDNA with each primer, and this result (Ct) will tell you at which concentration this primer is not efficient to determinate the amount. If cDNA is too concentrate, primers generate enough product very fast and will reach a early Ct, and if is too diluted, the replication is affected. Then if you plot the Ct against log of the concentration, you will find the limit of your primers. After that, you will know if you cDNA from your sample is in the confidence range. If is not, and the Ct is very small or high, you may dilute or concentrate your cDNA and then you can multiply the result according to the dilution factor.

If you need more information I wrote in my Ph.D. thesis a chapter about real-time PCR, discussing the application in plants (I work with Arabidopsis). If you need it please don’t hesitate to ask me.

Good luck!!

Thank you everyone.

Hi Chris,

As you suggested I did the same. I used 100 ng of RNA samples for synthesising cDNA and after that I used 1:5 dilution of synthesised cDNA (1uL) for running my qPCR experiment and got good amplification. Somewhere I read about the quantity of cDNA used in real time experiment and that point was bothering me as I did not measured the concentration of cDNA. So, I think the way I performed my experiment is right?

@Cris

Great comment !... Completely agree. Your comment must be part of any qPCR manual. Best

Paco

Completely agree with Chris!

total agree with Chris comments.

I have completed dozen genes qPCR from the same cDNA. The first step is to run qPCR with series cDNA dilutions for each qPCR assay to determine the best dilutions in order to have the Ct value between 15-25. . In my experience, from 1.0 microgram total DNA, the cDNA can be diluted from 5-20 depend on the level of gene expression.

Yizhou

You perform a standard curve run with your refernce primerrs/genes, in which you use different concentrations/dilutions of your RNA converted to cDNA. The curve will then tell you in which range you get the best amplification for your reference genes, meaning which concentration or amount of cDNA is best suited to get the best amplification. Reference genes are important here not your test genes. Running a standard curve for your test genes is non-sensical as you are normalising test genes to reference genes not reference genes to test genes. 

Thank you for your answers, I am also getting some useful points from the discussion. However, I am still facing a problem on constructing RT-qPCR curves based on the cDNA. I am working with an RNA virus which I have grown in cells and extracted the RNA and made cDNA. I have carried out a two step RT-qPCR with selected one primer from each of the ten segments of the virus. The cDNA was diluted 10-fold serially and each dilution used for each primer. Now, can some assist me which parameter do I use in constructing the curves? I have contructed the curves using the concentration from each of the dilutions on the X-axis and Ct values on the Y-axis. But now from this discussion, I am not sure which one to use now.

Hi Augustino, In the x-axis you put the concentration but in log scale... for example if you have 10fold dilutions, the higher concentration will be 1, then 0.1, 0.01 etc... and what you have to put in the plot is 0, -1, -2, -3 etc... I the y-axis you just put the Ct value. the linear model from all the points (don´t average the technical replicates, use them as individual points in the plot) gives you the model to calculate the concentration from any Ct value. the concentration (x) will be in log scale, so you need to naturalized using 10^x

Cheers

Juan

Hi Juan, Thank yo very much for your answer. It is ok for X-axis and Y-axis, you have brought another challenge to me about the replicates. I did the RT-qPCR in triplicates, I have 10 primer sets, one from each segment of the virus genome. I want to show the curves for all the ten sets of primers using the different dilutions on one graph in order to compare them. If I do not average the triplicates, then it becomes very difficult for me to draw the curves. I will have three Ct values for each dilution from one primer set. Please assist on how best I can organize this to draw the curves. Thanks.

Hi Augustino. In Excel use the Scatter plot, and do I did in this example, where I created data from 5 primers set, and 5 dilutions. The trick is that I repeat each concentration 3 times to add the 3 different Ct. Then the plotting is automatic. Be careful to average Ct values, because is not correct just average Cts, you need to do a logarithmic correction and it is not easy (the real average of Cts is = log[(base = efficiency) mean(Eff ^ Ct)]. 

In your chart you can put the equation if you like, so you will have a model to calculate the concentration of any Ct according to its primer set.

Cheers

Juan

Hi again Juan, Thank you very very much. With the example in the excel, it is very clear now. I also did using the  scatter plot, the error was on the x-axis where I used concentration and the average of the triplicates. I also tried using the GraphPad prism to draw the curve but it was not coming well as in the scatter plot (may be I don't know it well). However, it gave me the slope, the equation, the R square which were similar to the scatter plot. Thank you once again for this useful assistance. 

Cheers,

Augustino

You're welcome, and here is the same chart with 10 primers, where I put the equation bellow, next to the labels.. Use it if you want. Cheers

 Thank you. I will use it.

Regards,

Augustino

Hi all Here I have one query, does the dilution of CDNA before qPCR affects the Ct value ? Lets assume , I have to check the expression of three genes of a sample. Then the concentration of cDNA for in three different wells, meant for three different gene of the same sample shpould be equal..?

Yes dilution of cDNA before qPCR affects the Ct value. If you intend to compare the expression of the three genes at the end from the same cDNA, it will be good idea to use one diluted sample for all genes (equal concentration).

Thans a lot. Let we have two different samples and we have to see the expression of a single gene, then the dilution of the CDNA should be equal.? 

Thanks all, very productive discussion for those who are the beginner for real time stuff. Got my problem solved. :)

Dear Scientist

It depends on the amount of transcript for the gene you're interested in or housekeeping gene that you're working on. That's why you need to run dilution curves test to make sure that the amplification of your gene/housekeeping gene is within the 30 cycle range.

Best regard

Hello everyone,

I have trouble with amplification of a lncRNA in qPCR. When I overexpress my lncRNA of interest, my Cq values are about 11 but I have no amplification in my control and silenced wells. I am using gapdh and beta actin as housekeeping genes and their Cq values are about 15-16. I used nearly 1000 nanograms of RNA for cDNA synthesis which is the upper limit of the cDNA synthesis kit that I am using. What should I do to increase the cDNA concentration to have amplification in control and silenced samples?

do not get confused . just go to the first step of cDNA preparation and the concentration of RNA that you have used for making cDNA is critical. according to your cDNA preparation cycle calculate the concentration and use for real time pcr. or you can dilute your sample with buffer or water as 1:10 , 1:50, 1:100 and you can find the concentration .

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.

Ali Alavian-Ghavanini , that is an interesting question and I would also like to hear the opinion of some RT-PCR expert, which btw I am not. Anyway, in my opinion, it is not wrong what you are doing, however if you do so all of your cDNA will be diluted 1:5. Therefore your dynamic range will not include a first point with undiluted cDNA. If then you are going to do a serial dilution 1:2, you should be fine and maybe even better off, because instead of having 1:1, 1:2, 1:4 etc u will have 1:5, 1:10, 1:20 etc so you should cover a wider dynamic range with less points. However, if you have a highly expressed gene, and you are planning to dilute 1:10, then your dilution would be already 1:50000 (probably useless) after 4 folds of dilutions, so your curve would be based on only 4 points which is probably not good enough. In this case (highly expressed gene) you would be probably good to go with a 1:5 dilution of your initial 1:5 diluted stock.

So, at the end of the day, the main real problem that I see in your approach, is that you are losing the information relative to the undiluted sample. Therefore you will probably have a hard time to recognize a loss of linearity of your curve at lower concentrations, and to recognize the presence of inhibition in your matrix which could account for an efficiency lower of what your curve shows. Personally I prefer to include an undiluted sample, be aware of possible inhibitor effects showed by the curve, and then, if necessary, to exclude the lower dilutions to have the real value of efficiency for a specific dilution folder.

Paolo Tenti Thanks for answering. So what you mean is that:

The best approach is to start with undiluted sample and go further down the dilutions (1:5, 1:25, 1:125, :1:1625) . And then from the curve I select the best dilutions, is that correct in your opinion?

Extra q. : it will be correct to do this efficiency for each set of primers? regardless if they are gene of interest or house keeping. right?

Appreciate your help!

yes, starting from 1:1 I dilute 1:2, if I know it is very low expressed, or 1:10 for HKG, or sometimes 1:5 works well too. If I have a "horizontalization" of the curves in last 2 or 3 most concentrated samples, I try to remove 1:1 and maybe 1:5 and if necessary even 1:125, but I try to maintain at least 5 folders points (or more if I diluted 1:2). Yes, you need to repeat dilution curves every time your assay changes, regardless if it is the primer used or the mastermix or even the RTPCR machine used. I know, it is very cumbersome