Hello Everyone.
Currently I am working to characterize macrophages in the myocardium after ischemia-reperfusion injury in rats. Due to the low total cell number isolated from rat hearts I can only do ~200K cells per tube for flow cytometry. Recently I have finally been able to process the samples without losing the cells during the permeabilization. However, I am now facing another issue. When I perform flow cytometry I notice that the cells tend to stick together unlike when I do fixation alone. I attached an image exhibiting this; the FSCvsSSC data was captured during the same run and using the same voltages. This seems to be interfering with the antibodies I use (CD45-PE and CD68-AF647) as I can't get their emission peaks to show up when I do flow. I know these antibodies work since when I perform this experiment on isolated PBMCs everything goes well. I wanted to ask for any input on how to resolve this issue. I've been recommended to increase antibody concentrations, change SSC and FSC voltages, or use a different cell type with high expression for both markers for compensation controls. Can there be anything else that I am missing? Thank you in advance as I am quite new to flow cytometry.
The shift to the left in forward scatter is not because the cells are sticking together but because the cells have reduced in size as a result of the protocol. You may need to check cell viability using 7aad
Lora Benoit Thank you for the input. That makes more sense now that I think about it. Do you happen to know whether I can prevent this morphological change? I'll definitely check viability first. Thank you for your time.
Conceptually, i would recommend that you ensure that your cells are always in isotonic buffer. Also, it may help if you use a reversible perm detergent such as saponin-based rather than triton-x, which is non reversible. ; )
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