I want to quantify how many "green" cells also express a "red" marker. This seems to be easy to quantify by overlay (for example with the Mander's coefficient), but I am having a hard time because green and red markers, although expressed in the same cells do not colocalize in the same cell compartment, so they do not fully colocalize. The result is that with Mander's coefficient, I feel that I am underestimating the real proportion of green cells that also express the red marker (because the maximum M value I get in a cell/group of cells that express both markers is approximately M1=0.5 when I consider that as full match). Can anyone suggest another way of measuring marker overlay?
PS: There is no way, in which I can just count the cells as positive or negative because they form long membrane processes and it is difficult to distinguish which nucleus a cytoplasm/membrane corresponds to; they form a mesh.
depending on your setup, you might consider including another stain for e.g. ZO-1 to label junctions (ZO-1 in endothelial cells) and then using this signal to distinguish cells (in ImageJ/Fiji)
I guess that even with junction labelling it will be imposible to distinguish cells, but I will try once and I will see.... I was thinking in a some kind of algorithm that matches (by morphology for example) or interpolate coexpression of two markers even when there is no full celullar colocalization. I just want to measure coexpresion, not formal colocalization. In my immunohistological tissue samples I have noticed that there is huge variability (in "shape" not "contain") between stainings of the same marker depending on how you develop the primary antibody. The pattern it is not the same when using TSA, ABC, TSA+ABC, second antibody only... So one of my markers is brigth, continuous and smooth, and the other one is discontinuous and dotted-like, so obvioulsly Mander's coeficient gives me a low value... I really can not find a clue how to cuantify coexpresion (not colocalization) on my IHC slides...
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