1 reply to this topic

[BSM]

I ran my first RT-qPCR today with some cDNA created from brain tissue. I am looking for the expression profile of chemokine ligands to the CXCR2 receptor. I am using a bio-rad icycler with the Taqman Iq supermix.

The max fluorescence for the four genes is only around 500-600 RFU and the max fluorescence of my HPRT housekeeping gene is even shorter.  The calculated Ct value for these genes is around 31-32 cycles but since the HPRT gene ended so soon the derived ct value is too low. I diluted my cDNA 1:5 and ran about 150 nM for my probe concentrations in a 20microliter reaction volume.

The primer sequences and recommended molar concentrations of primers I used were based on a previous paper that performed a similar assay. The paper that provided this info was very hard to interpret. They listed concentrations of probes and primers for each gene and also the "optimal ratio" as shown in this graph:




Are the concentrations they list here the concentrations found in their final reaction volume? They seem insanely high compared to the usual recommended concentration of 100-500 nM. If they are the final concentrations, how come the optimal ratios do not match up correctly? For my primers, I just used the optimal ratio for forward and reverse. Can anyone interpret this graph and the issues I am having above?

Thanks!

[BSM]

I have also attached two plots representing my qPCR data. If you notice the plot with the background subtracted RFU, the RFU values at 0 cycles are only around 100-150. I looked at data from other lab members here and their background fluorescence at 0 cycles is near 1000 RFU. The HPRT probe I used was at 40x concentrate so I know for sure that I had the right concentration of it in my samples. Could the probes be too old? They were used in 2006. Just a thought.

Edited by BSM, 20 August 2009 - 10:51 AM.