共焦点顕微鏡を用いた可視化マクロファージ細胞外トラップ

The overall goal of this method is to demonstrate how macrophage extracellular traps can be visualized and studied using confocal microscopy. This method can help answer key questions in the inflammation field, such as responses to infection and other immune stimuli. The main advantage of this technique is that it is a modification of a well established method that has been used to study neutrophil extracellular traps.

Demonstrating this procedure will be Roleen Sharma. After obtaining lung macrophages from humans and mice by bronchoalveolar lavage, or BAL, according to the text protocol, use trypan blue and a hemocytometer to perform a viable cell count on the BAL solution. Pipette one to four times 10 to the fifth macrophages in 500 microliters of culture medium onto cover slips in the wells of 24-well plates, then incubate the cells at 37 degrees Celsius overnight for the cells to adhere.

Remove the culture medium and use PBS to wash the cells once. Then add 2%paraformaldehyde-periodate-lysine, or PLP fixative, and incubate the samples for 10 minutes. After using PBS to briefly wash the cells, add 0.2%TWEEN 20 in PBS and incubate the cells for 20 minutes.

Next, to block the cells, add 10%chicken serum in 5%BSA, diluted in PBS and incubate the samples at room temperature for 30 minutes. Then replace the block solution with a one to 100 concentration of primary and isotope control antibodies and incubate the cells at room temperature for one hour. Following the incubation, use PBS to wash the cells before adding the corresponding secondary antibodies.

Then incubate the samples for 40 minutes at room temperature. After washing the cells in PBS, mount the samples with DAPI based mounting medium, then visualize the samples using a confocal microscope. To pretreat the FFPE samples with antigen retrieval solution, oven dry the slides at 60 degrees Celsius for 60 minutes.

Then transfer the slides into xylene solution for 30 minutes before transferring the samples to 70%ethanol at room temperature for five minutes. After using tap water to rinse the slides, place them in heat-proof plastic wrap and subject them to higher antigen retrieval by placing them in a pressure cooker in Tris-EDTA pH 9.0 for 10 minutes. Cool the samples for 20 minutes and transfer them to tap water.

Then place them on a rocker for five minutes. Repeat the tap water wash and then wash the slides once in PBS on the rocker. To block the samples, add 10%chicken serum in 5%BSA PBS and incubate them at room temperature for 30 minutes.

Then, to define macrophage extracellular traps, or METs in macrophages, add a one to 100 dilution of primary antibodies in 1%BSA PBS and incubate the slides at four degrees Celsius for 16 hours. Use PBS to wash the samples two times on a rocker for five minutes each. Add the corresponding fluorescent secondary antibodies in 1%BSA PBS and incubate the slides at room temperature for 40 minutes.

Following PBS washes, use DAPI containing mounting medium to stain the chromatin and mount the samples. Using a confocal laser scanning head attached to an inverted microscope, capture fluorescent images using 20X 0.1 NA air and 40X 1.0 NA oil objectives. Capture single plane 512 by 512 pixel images by clicking on the Line Sequential leveling button.

Obtain at least 10 fields of view per section for analysis and data for each result. To stain the sections in microcentrifuge tubes, add the relevant primary antibodies in 200 microliter volumes and incubate the samples at four degrees Celsius for 16 hours. Wash the sections in PBS three times for 10 minutes before adding the appropriate secondary antibodies and incubating the samples at room temperature for one hour.

Use DAPI to stain the lung sections in solution and incubate the samples for 20 minutes before adding cover slips to the sections on microscope slides in PBS. Use an inverted confocal microscope with a 40X 1.3 NA objective, equipped with 405 nanometers, 440 nanometers, 473 nanometers, 543 nanometers, and 635 nanometer lasers, to capture images and to record 3D Z stacks, 0.54 microns thick, with optimal Z sectioning. Finally, analyze the images according to the text protocol.

An example of METs in a BAL sample is shown here. The first detectable feature is the movement of the nucleus to the edge of the cell, as seen in this earlier stage MET that has a roughly spherical shape. This is followed by extracellular chromatin with other co-expressed mediators, such as H3Cit and granular proteases, as seen in this more mature MET.

Earlier stage MET is upper left and later stage MET is below this towards the center. Lung tissue METs are shown in this figure. As the lungs are inflated with fluid to define the lung architecture, the METs will be pushed against the alveolar walls.

The morphology of the METs will also vary depending on in which plane the tissue was cut. The standard sections used for lung tissue samples are four to five microns thick. Thicker tissue sections enable multiple images to be taken and the METs can then be viewed in 3D.

An example of a 3D image from mouse lung tissue cut in 25 to 35 micron sections is presented here. Once mastered, this technique can be done in two days of staining plus imaging if it's performed properly. While attempting this procedure, it is important to remember to be gentle with the washes.

Following this procedure, other methods like zymography can be performed to answer additional questions like protease expression. After its development, this technique may pave way for the study of inflammatory macrophage responses. After watching this video, you should have a good understanding of how to visualize METs using confocal microscopy.

Don't forget that working with PLP can be hazardous and precautions such as wearing gloves and eye protection should always be taken while performing this procedure.