The over goal of this technique is to monitor the different stages of phagocytosis using a confocal microscope. This method can help answer key questions in the field of cross pathogens interactions. Mainly to understand how pathogens enter the host cells during infection.

The main advantage of this technique is that it is simple and enable us to gain information about the virus cell of our events that occur during phagocytosis in a time sensitive manner. To begin, prepare the fluorescently conjugated zymosan as described in the accompanying text protocol. The previous day, seed about five times 10 to the fourth DC 2.4 cells in a chamber slide, so that the wells are 60 to 70%confluent at the time of the experiment.

Once the cells are at a proper density, place the chamber slide at 10 degrees Celsius for 10 minutes. Cooling of the cells is necessary to block endocytosis as well as when to toggle cytosis. And subsequently to have a synchronized phagocytosis.

Next gently remove the medium from each well using a pipette or aspirator. Then, mix the fluorescently conjugated zymosan with cell culture medium at a multiplicity of an infection of 10. Next, add 500 microliters of medium into each chamber well to ensure that all of the cell are covered.

Then spin down the chamber slide to both increase the contacts between the particulates in the cells and synchronize the bonding process. Following centrifugation, transfer the chamber slide to a 37 degree Celsius humidified incubator with 5 percent CO2. After five minutes at 37 degrees Celsius, remove the chamber slide from the incubator and aspirate the medium.

Then, wash the cells three times with ice cold PBS. After the final wash, check for the presence of unbound particles with a light microscope and perform additional washes if necessary. It's essential to remove unbound particles from the wells to minimize background noise during the analysis.

Next, fix the cells by adding 500 microliters of four percent paraformaldehyde in PBS to the wells, and incubating them for 30 minutes at room temperature. After fixing the cells, wash them by adding 500 microliters of PBS into each chamber and then gently aspirating the solution. Repeat the washing step two more times.

Following the third round of washing, stop the reaction by adding 500 microliters of 50 millimolar ammonium chloride in PBS to the cells and incubating them for 10 minutes. Then, wash the cells in 500 microliters of room temperature PBS. Next, permeabilize the cells by adding 500 microliters of binding buffer to each well and incubating the cells for 30 minutes at room temperature.

The next step is to stain for F-actin by adding fluorescently conjugated phalloidin to the cells and incubating them for one hour at room temperature. Then wash the cells three times with 500 microliters of PBS. Finally, capture images using a confocal microscope and analyze the images using ImageJ.

In order to measure the zymosan uptake, prepare chamber slides with cells as shown in the binding assay. Following the cooling and labeling steps, transfer the chamber slide to a 37 degree Celsius humidified incubator with five percent CO2. And incubate the cells for different time points.

Fix the cells after 30, 60, and 90 minutes post infection. Be sure to optimize these time points based on the respective cell type and fluorescent particles that are used. Then, analyze images using ImageJ.

Successful uptake is characterized by a complete internalization of a particulate inside the cytoplasm of a phagocyte. To begin, prepare a microscope stage environmental chamber by warming it to 37 degrees Celsius and equilibrating the chamber with 5 percent CO2. Wait 30 to 45 minutes for the temperature and the CO2 to stabilize before imaging.

Seeing the CO2 and the temperature levels are critical for phagocytosis, as important to turn on the microscope heater prior to the experiment to equilibrate the environment of that chamber. Next, prepare chamber slides with DC 2.4 cells which stabilate express mCherry F-actin as shown in the binding assay. Following the cooing and labeling steps, transfer the chamber slide from the centrifuge directly onto a pre-warmed microscope stage.

Finally, perform live imaging using a confocal laser scanning microscope as described in the accompanying text protocol, to not only capture the movement and dynamics of the F-actin network, but also to visualize the different phagocytic events within the living cells. The cells shown here in green is a wild type DC 2.4 cell that is being infected with Canadian BFP which is shown in red. Timelapse imaging is used to visualize the early stages of phagocytosis and is shown here in 30 second intervals.

The locations denoted by the asterisk are where actin remodeling is occurring. The entire process occurs in a matter of a few minutes. Here the role of sphingolipids were evaluated for their role in the early stages of phagocytosis.

Using this technique, sphingolipids were found to be essential for the binding of candida albicans to the cells and reduced overall phagocytosis. This reduction of phagocytotic binding was also shown for DC 2.4 cell binding to zymosans. After watching this video you should have a good understanding of how to perform live imaging to study the early stage of phagocytosis.

Once mastered, this technique can be done in one to two hours if it is performed properly. Following this procedure, the phagocytosis of other pathogens, including bacteria and the parasites, can be performed in order to understand the intricate relationship between the immune cells and of this pathogens. After its development, this technique paved the way for researchers in field of cell biology, to explore receptor mediated endocytosis including phagocytosis and a macropinocytosis in immune cells.