Hey guys! I have been working on Quantitative PCR for quite some time and am still having problems. I'll explain my situation.
I've got a set of candidate genes I'm interested in and am using B-Actin as a reference gene. I'm looking at relative gene expression differences in three tissues, one of which is serving as the reference sample for relative quantification. I've isolated total RNA and synthesized cDNA from pools of 10 individuals in each of the templates. ( 10, 10, 10) and then done this 3 times. (10,10,10) (10,10,10) (10,10,10) I've run my gene expression tests in each of these 3 Biological replicates of pooled individuals, but I've gotten rather inconsistent results.
It started out that I thought my primers were good, but they turned out to be non-specific. Then, I redesigned the primers to get specific amplification, but had primer dimer. So I ran a Primer Dilution matrix to get ideal levels to use in reactions. Now I have rerun the 3 templates with all the genes and the results are inconsistent. For those that show the same upward or downward expression, there's quite a bit of variance (ratios from .2 something to nearly .9 for ex.). Others don't even show the same trends b/t the three samples (some upregulated some down). Anybody know what could be going on here? Have any suggestions. This has had me held up for nearly a year now.
Note: I used the Pfaffl Efficiency Correction Method to compensate for Primer efficiencies.
Submit your paper to J Biol Methods today!
1 reply to this topic
Posted 05 April 2010 - 01:00 PM
I think that the expression of your genes is not significantly different among your sampes. The differences lower than 2 Ct are most probably bellow sensitivity of your assay. You can do some statistics to confirm this, but I think it can be seen at the first sight. The result should be: there is no difference in expression level of these genes among the three tested tissues.