I'm using the SpectraMax M3 from Molecular Devices to read 96 well plates which contain wells with E. coli transformed with vectors containing a fluorophore under an inducible promoter. On this same plate, I include wells with the same bacteria transformed with vectors that lack this fluorophore but are identical otherwise (my negative control) and wells with clean media. I take OD readings and fluorescence readings using the same plate reader and I normalize the RFU readings to the OD.
I'm trying to measure the difference in normalized fluorescence between the induced and uninduced samples on my plate. Before calculating this fold change, I want to subtract my negative control from all of my induced and uninduced samples but my problem is that these wells consistently have higher normalized RFU readings than my actual samples! I can't figure out why this is happening and would really appreciate some help with this problem.
For further clarification, my media is LB for all samples and the two fluorophores I use are GFPmut3 and mRFP1. With GFP I use an excitation of 500 and an emission of 513 and with RFP I use an excitation of 584 and an emission of 607. My plates have black wells and clear bottoms and I use a bottom read for reading fluorescence.
Can you detect a peak in the fluorescence spectra of the G/RFP proteins in the induced cells above the background scattered light spectrum? If not, the level of expression may be too low.
Also, there is a risk in normalizing the RFU to the OD if the different strains do not scatter light to the same extent due to a difference in shape, or if the OD measurement is not an accurate report of the number of live cells (dead cells also scatter light).
Another issue with the OD is that the RFP absorbs light at 600 nm, which could contribute significantly to the OD600 reading if it is well-expressed.
Finally, GFP/RFP proteins are stable enough to continue to fluoresce even after the cells that made them die, which may be a problem for some types of experiments.
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