I’m trying to use CellProfiler to determine DNA content and fluorescence signal (for my cell cycle protein) in each cell to see if the levels of the protein fluctuate during the cell cycle. Before trying with my cells I wanted to be able to get a multi-image set from human cells that would give me a decent histogram (with defined 2N and 4N peaks). I used 4 DAPI images from the exampleSBSimage set images and tried to correct for the image to image illumination and staining variation outlined in your BMC bioinformatics paper by taking the mode value of log(integrated DNA intensity), X, for each image, converting it to log2 (by multiplying X by 3.3219), raising 2 to the power (X*3.3219)-1] to get a denominator Z, that should, when the entire corresponding image is divided by it, and a second histogram generated give me a log(integrated DNA intensity) of 0.301 (would give 1 if log base 2). However that’s not what I get, there is still considerable variation between the modes. What am I doing wrong and what do I have to correct?
I get the mode from a cellprofiler generated histogram (pipeline test, then use ratio for the histogram, and 50 bins) and then carry out the calculations using excel, then adjust the images using a second pipeline and then rerun the original pipeline. I haven’t used cell profiler analyst, since my data set is very small (200 images and only 2 parameters), and there is no one in the department with expertise to set up the database required to run cell profiler analyst. A second question is: is there a way to automatically extract the mode value of a histogram for each image in cell profiler?
Thanks a lot for your help
testPIPE.mat (950 Bytes)