Calibrator sample - (Feb/11/2013 )
I am trying to analyse gene expression over a time period. I used the delta delta Ct relative quantification method for analysis. My endogenous control is Actin and my calibrator sample is the cells obtained from the first time point (Undifferentiated cells).
After completing the first round of experiments, it is evident that my gene of interest is not expressed in my calibrator sample (time 0: undiff cells). Therefore I am unable to calculate the fold changes in gene expression relative to this time-point. The endogenous gene was amplified in my samples, but the gene of interest was not. Moreover, my no template control and negative control did not show any amplification.
Ultimately I would like to show that my gene of interest increases over time. Is there another method that any of you can recommend I use to analyse my data?
You say that the endogenous gene is expressed but the GOI isn't - I take it that these are done using separate primers? If so, do you have a positive control to show that your PCR for the GOI is working?
Yes, two separate primers were used. I do have a positive control to show that my GOI is working. Moreover, I do see an increase in expression of this gene at the other time points. However what I want to show from my results is, that compared to the undifferentiated cells (calibrator sample), the levels of gene expression increase over time.
Do you know if this is possible?
I am assuming that your GOI is in stable transfected cells and you have determined the expression at t=0 by other diagnostic tests? What you are asking may not be possible. I usually design primers specific for my overexpression vector containing my insert, just so I may determine that expression of my plasmid+GOI is occurring (or run a WB if tagged). What are you comparing your transfected t=0 sample against to determine if it has expression?
Expression of your GOI within a plasmid should be relatively consistent. It should not decrease or increase over time. That is why (basically) all vectors contain strong promoters before the MCS. You should also keep in mind that when you are running a RT-PCR you are looking at a snapshot of the cell. You are looking at all the RNA that is present at that point in time. If you were to isolate 10 RNA samples from the same cell line at the same point, expression of your plasmid+GOI would vary. It is important to determine if your plasmid+GOI is being expressed via RT-PCR, but I would personally stick to using primers that cannot distinguish between your endogenous and exogenous gene. Only because it will give you a better understanding of expression change at the RNA level.
Did that answer your question?
Edit: I can't write a cohesive sentence.
Thanks for your replies.
I am trying to use the relative quantification method (not absolute). Hence, I believe it is not necessary for me to create a plasmid.
Therefore my question is, how to calculate fold change in gene expression if your gene is not expressed in your calibrator sample? I think I may just have to choose another calibrator sample and calculate the fold changes, and merely note that my undifferentiated cells did not express my gene of interest.
Also does anyone know how to calculate delta standard deviations for fold changes?