Dear Dr. Vahinipriya Manoharan

Better not. Let the macrophages adhere before performing the assay.

Macrophages generally need to be adherent to a surface to effectively perform phagocytosis. The initial attachment provides the necessary mechanical and signaling framework for the macrophage to carry out the complex steps of phagocytosis. This attachment allows the macrophage to form a stable connection with the target (pathogen) and triggers the necessary cytoskeletal rearrangements and receptor activation for engulfment.

The cytoskeleton, primarily actin filaments, is essential for forming the pseudopodia (cellular extensions) that surround and engulf the target. Attachment to a surface allows the macrophage to exert force and shape these extensions effectively. The membrane protrusions extend and fuse around the target, creating a membrane-bound vesicle called a phagosome, which is then internalized into the cell.

Without attachment, the process of phagocytosis is less efficient, or even impossible, due to the lack of proper engagement of receptors, cytoskeletal reorganization, and formation of the phagosome.

In a flow cytometry-based phagocytosis assay, you quantify the number of macrophages that have successfully engulfed the target cells. To do this, you detach the macrophages from the substratum they might be attached to (e.g., by trypsinization), and then the detached cells are analyzed. The flow cytometer then measures the fluorescence or other properties of the detached cells to determine which macrophages have ingested the target particles.

Regards,

Malcolm Nobre

You wrote that peritoneal cavity cells will be used for your phagocyte study. I understand that you have WT and gene-specific knock out mice, from which you will isolate peritoneal macrophages and compare phagocytic activity using flow cytometer.

If so, I have a better idea, based on our previous findings (1). You just inject living motile bacteria (Such as Salmonella typhimurium, Escherichia coli etc) to the peritoneal cavity and then collect macrophages by washing the cavity and count number of bacteria per cell. I would like to suggest the following protocol.

1. Inject 1 milliliter of bacterial suspension containing 5 x 108 into the peritoneal cavities of WT and gene-knockout mice.

2. Collect peritoneal cells after 15-30 min by washing the peritoneal cavities with ice-cold HEPES MEM containing cytochalasin B (5 ug/ml).

3. Wash the cells twice by centrifugation at 110 x g for 5 min with fresh medium mentioned above.

4. Use the cells for flow cytometry analysis

5. Alternatively, smear the cells on glass slides using Cytospinner.

6. Dry and fix with methanol and stain with Giemsa stain.

7. Cout number of bacteria per cell at least 200 macrophages.

8. Determine percentage of macrophages ingesting bacteria and the mean number of ingested bacteria per cell.

Reference:

Tomita, T. and S. Kanegasaki: Infect. Immun. 38, 865-870, 1982

(doi: 10.1128/IAI.38.3.865-870.1982)

Good luck!

If you insist on testing in vitro with a suspension culture, this is also possible using live, motile bacteria. See the reference for detail. You can find out why this is possible by reading it.

Since the cells remain on the culture plate for 5 hours during the phagocytosis assay, they become strongly adherent. Scraping them in 2 mM EDTA for flow cytometry is causing significant damage—I see most of the cells appearing as debris in the FSC/SSC plot. I’m using non–tissue culture–treated plates.

What else can I do to harvest the cells without damaging them? Do you think performing flow staining directly on the plate and then scraping the cells after fixation would help reduce cell damage?

If you use Teflon Petri dish to culture mouse macrophages, you can remove them easily. Write more details of experiments you want to do, so that I can provide better advice. Why use adherent culture?