I am doing qPCR but my HPRT1 Ct is not coming at the same point. I see a 2 cycle spread in Ct. I have checked my RNA quality A260/280, 230/280 values, did ethidium bromide gel. I used superscript for cDNA synthesis. no DNA contamination in -RT samples. I am pipetting 50ng cDNA in each qPCR well. The triplicates are exact replica so no pipetting error in qPCR. I tried using sybrgreen and eva green both for qPCR but results are same.
I used RNA samples from another colleague and its showing the same spread in HPRT1.
First, 50ng of cDNA is LOADS. A tenth of that would be fine.
Second, the whole point of a reference gene is to normalise for variation between samples. If your samples were all completely identical and somehow no sources of variation existed, you wouldn't need a reference gene.
But: there are mystery sources of variation we cannot account for (RNA integrity, RNA isolation efficiency, cDNA synthesis efficiency etc) and so the only way to handle that is to use a panel of genes we know to be reasonably stable in expression for the conditions studied. The assumption is that these genes are NOT changing in expression between samples, so any differences in expression we see MUST therefore be attributable to differences in underlying cDNA content, and we can therefore just normalize for that.
So...this is fine. Assuming you know HPRT1 is stable in your experimental conditions, then just normalize to the measured values.
And use less cDNA. And ideally use multiple reference genes.
I agree with John Hildyard . It is completely normal to have minor Ct difference of 1 to 2. But if this difference in Ct is more than that, you need to re assess your experimental conditions or use a different housekeeping gene.
Did you conduct a reference gene stability test i.e. GeNORM, RefFinder, BestKeeper etc? Ct values can be influenced by cDNA input and real time PCR dilution. If any of those change, then your Cts will shift. For example, you had extrated RNA from 100,000 HeLa Cells, then used 500 ng of RNA for cDNA syntehsis, then you made 1:100 dilution for cDNA as working PCR stock. Then if you extracted later from 100,000 cells, but this time used 1000 ng of RNA and 1:100 dilution, you'd get 1 Ct earlier results for the 1000 ng results compared to 500ng input. 2 ct is a lot of difference, that's 2^2 = 4-Fold change difference. I'm not sure I agree with the advice that 1-2-fold change is something to ignore, my take is it depends...did you do a reference gene stability test to see whether your samples appear homogenous or heterogenous samples i.e. with geNorm these are 0.5 & 1.0-M values. If you are using a cell line, one would expect your replicates to behave like a homogenous sample because cells are not biological replicates (won't behave like mouse tissue/human samples). If you are working with a tissue sample, 1 gene may not be enough. If you want accurate quantification, you may need two or more (as it happens in cancer samples). So the answer to your question lies in your reference gene stability test that you conduct with all your relevant conditions.
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