Hello, I am using Taqman probes in RT-PCR in order to identify gene expression in genomic DNA but this expression is very low and I am trying to do it in a higher concentration of the tamplate DNA sample. Which is the highest concentration I could use if I want to be in the linearity range?
Hi Jud,
It is not entirely clear what you are attempting to do. 'Gene expression' implies you are looking at mRNA (converted to cDNA), i.e. genes that are actually being expressed. Genomic DNA will not give you this information.
If you are looking at genomic DNA alone, then I can only assume you are looking for gene copy number (how many copies of the gene there are in the genome). This will almost always be low: typically 'one' for haploid genomes, and 'two' for diploid. Some genes are present at higher copy number (ribosomal RNA genes have copy numbers of 100-200) but most genes will only be present as single/double copy. This is...fairly basic genetics.
Even under these stipulations, you should have no trouble getting quantitative answers with nanograms of material (a nanogram of gDNA typially represents 100-1000 genomes, depending on genome size: well within the sensitivity of a good qPCR assay).
If you are instead looking at cDNA (i.e. you have isolated RNA and reverse transcribed it to cDNA), then the next question I would have is "how much are you using currently?"
Even relatively low-expression genes are readily quantified with 5-10ng of cDNA (I typically use 8ng for everything), and if you're picking up stochastic levels of expression (say, 0-10 transcripts per well) with 8ng of cDNA, it is reasonably safe to conclude your gene is not really being expressed at all.
The main danger with low expression in qPCR is using too much cDNA: cDNA synthesis buffer contains reagents that inhibit PCR, so adding a load of undiluted cDNA to a reaction not only wastes lots of cDNA, it also makes that reaction inaccurate (you may not get a Cq value at all, even). I would always dilute my cDNA after synthesis (by 1/10 or 1/20 at least), and then only use 1-4ul of that.
I would suggest you perform a serial dilution to establish the linearity of your data: start with a high [cDNA], such as 20ng or so, then dilute in series, perhaps 2 fold, or 5-fold, or even 10-fold: if your data is quantitative and within the linear range, your measured Cq values should increase in line with your dilution. Make sure you also do this for a gene you know to be expressed robustly (you will need a reference gene anyway -ideally two or three for MIQE-compliance).
If you get no expression at ANY dilution (not even with 20ng) while your reference gives you acceptable data, I would conclude your gene is not expressed. If you get nothing for your GOI or your ref gene, I would remake your cDNA.
Best way is to make dilution of cDNA in that way that you test multiple dilutions: i.e., 100ng, 50ng, 1ng, 100pg, 50pg, 10pg, 5pg, 1pg, 0.5pg per reaction mixture.
That will give you a range where you get right copies amplified. If still comes weak curve, than you may replace other reagents, start with fresh aliquotes of reagents,
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