I am trying to label mitochondria in human dermal fibroblasts to assess mitochondrial transfer via tunnelling nanotubes to osteosarcoma cells. I am using a direct coculture approach whereby seed fibroblasts first and then 24 hours after seed osteosarcoma cells and perform time-lapse imaging on a confocal microscope for another 24 hours. I am using a direct coculture approach whereby seed fibroblasts first and then 24 hours after seeding osteosarcoma cells and performing time-lapse imaging on a confocal microscope for another 24 hours.
No matter how many times I wash my fibroblasts (that had been labelled with MitoRed), I can see the osteosarcoma cells pick up the label uniformly after a 10 minute period of coculture. I have tried washing my MitoRed overnight, labelling and washing MitoRed while cells are adherent to the substrate and I have tried labelling and washing MitoRed labelling cells in suspension (up to 7 washers in HBSS). I have even tried using MitoRed in various concentrations from 1:10,000 - manufacturers recommendation, down to 1:200,00. Even at the lowest concentration, there seems to be residual dye that the osteosarcoma cells pick up and coculture. As a result - it is almost impossible for me to determine whether fibroblasts have transferred some of their mitochondria into osteosarcoma cells.
Are there any ideas or approaches I could try that will allow me to assess the extent of mitochondrial transfer from one cell type to another? Many thanks in advance
What is the lowest concentration of mitotracker you can use to detect it in the dermal fibroblasts? I'd use that followed by extensive washing and see how that goes. Also just out of interest you should set the experiment up as if you were expecting the osteosarcoma cells to pick up the mitotracker straight away and start imaging probably on a spinning disk to see where the mitotracker actually comes from, if its from the mitochondria or within the fibroblasts or if it seems to come from somewhere else.
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