Neutrophil Extracellular Traps: How to Generate and Visualize Them

Neutrophil extracellular traps or nets are a recently identified mechanism for neutrophil mediated clearance of pathogens by releasing a combination of cytoplasmic and granule proteins, as well as chromatin neutrophils form extracellular fibers that trap pathogens. In this video, granulocytes are isolated from human whole blood by histopath density gradient centrifugation. Neutrophils are isolated from the granulocyte preparation by per coral gradient.

Centrifugation seeded onto cover glass and stimulated to form nets using chemical or biological stimuli, the cells are then processed for immuno labeling or em for visualization of nets. Hi, I'm Christian Gosman from the microscopy core facility at the Max Plunk Institute for Infection Biology. Today we will show you a procedure to purify neutrophil granulocytes from human blood and stimulate them to generate neutrophil extracellular traps or nets.

We will then visualize nets by immunofluorescence and scanning electron microscopy. We use this procedure in our lab to study the mechanisms that lead to net formation and to analyze the antimicrobial properties of nets. So let's get started.

Begin this protocol in a lamina flow hood by preparing six milliliters of histo 1119 into each of four to 15 milliliter Falcon tubes. Carefully layer five to seven milliliters of whole blood over the histo in each tube centrifuge tubes for 20 minutes at about 800 times G, ensuring that the break is off once the centrifugation has completed. Carefully remove the tubes from the centrifuge.

The tube will contain four clearly separated layers from top to bottom. These are a dark red layer containing erythrocytes. A light red layer containing the neutrophil granulocytes with some erythrocytes, a whiteish yellow layer containing leukocytes and serum, and a clear yellow layer of serum.

Again, working in the lamina flow hood. Use the perpet attached to a vacuum system to carefully aspirate the top yellow layer. Then using a pipette transfer the reddish phase containing granulocytes from each tube into a new falcon tube.

Next, fill the tubes with PBS to wash the granulocytes. Then pellet them by centrifugation for 10 minutes at about 300 times G while the cells are spinning. Prepare a 100%per coll solution by mixing 18 milliliters of per co with two milliliters of 10 times PBS.

Then use the 100%per coll solution and one times PBS to prepare four milliliters each of 65 70, 75, 80, and 85%per call. Use these solutions to prepare per core gradients by carefully layering two milliliters of each solution into each of two tubes. Begin by petting two milliliters of the 85%solution into each tube.

Then on top of the 85%per coll, carefully layer two milliliters of the 80%per co. Continue layering two milliliters of each solution in decreasing order such that the 65%solution is layered on top. Remove the sedimented leukocytes from the centrifuge under the hood.

Discard the supinate and resuspend the cells in one milliliter of PBS. Then combine the resuspended cells in a fresh tube. Carefully layer two milliliters of the resuspended granulocytes onto each of the perol gradients.

Then centrifuge for 20 minutes at about 800 times G, ensuring that the break is off after centrifugation. The interfaces should be clearly visible due to the highest cell density. Discard the top layer and most of the 65%layer, which contain peripheral blood mononuclear cells or p BMCs.

Collect the 65%70%interface together with a 70%75%and 80%85%Interface from each gradient and transfer them to new fresh falcon tubes. Wash the cells by filling the tube with PBS and centrifuge for 10 minutes at about 300 times G to pellet the cells after centrifugation. Remove the supinate and resuspend the sedimented, polymorph and nuclear leukocytes or PMNs in two milliliters of PBS.

At this point, 95%or more of the cells in the preparation will be mns. Using a hemo cytometer determines cell counts, then see 200, 000 cells in 500 microliters of RPMI onto sterile round cover slips placed in culture plates with 15 millimeter size wells. Incubate for one hour in a 5%CO2 incubator at 37 degrees Celsius to allow the cells to attach to the cover slips after one hour.

Add 100 microliters of RPMI containing the desired stimulant. For a time course experiment, add the stimulant sequentially, starting with the longest stimulation time for each conditional time point. Be sure to prepare a positive control by adding 600 nano molars of PMA in RPMI.

Also, for each condition or time point, prepare a negative control by adding RPMI alone incubate for up to four hours in a CO2 incubator at 37 degrees Celsius during stimulation, the formation of nets can be monitored using bright field microscopy. Shortly after stimulation, nuclei changed their form dramatically. They lose their LOEs round up, then expand at the desired time point at 600 microliters of room temperature, 8%paraform aldehyde solution to each well.

To bring the final concentration to 4%incubate for two to four hours for immuno labeling. Place a paraform sheet on top of a test tube rack with three times eight bore holes of approximately 15 millimeter diameter. To form wells, fill wells with PBS.

Use a curved needle to lift up each of the cover slips from the paraldehyde on the culture plate. Then using forceps, grasp the cover slip and invert it onto the surface of a drop of PBS on the paraform. At this point, neutrophil extracellular traps are very fragile and great care should be taken during manipulation to avoid cell loss.

After five minutes, repeat the wash by transferring the cover slips to new drops of PBS. Repeat for a total of three washes. Next, transfer the cover slips to a drop of room temperature 0.5%Triton X 100 for one minute to permeate the cells.

Then wash three times in PBS for one minute each following the wash. Proceed with the staining according to the accompanying written protocol. Performing each of the incubations and wash steps by inverting the cover glass on the paraform.

Once the cells have been stained, place a 20 microliter drop of MO onto a glass slide and mount the cover slips face down with the cells between the cover slip and the slide. After one hour, the MOY should have solidified and the cells are ready for microscopic analysis. With immersion lenses, prepare the cells for scanning electron microscopy under a chemical fume hood.

Post fixx the cells on the glass cover slips in a 24 well plate with 2.5%glutaraldehyde in PBS for 30 minutes. During the incubation, place a paraform sheet on top of a test tube. Stand with three times eight bore holes of approximately 15 millimeter diameter.

To form wells. Fill the wells with distilled water. Use a curved needle to lift up each of the cover slips from the glutaraldehyde solution.

Then using forceps, grasp the cover slip and transfer it to a drop of distilled water on the paraform wash by transferring the cover slips to drops of distilled water on Paraform. Repeat for a total of three washes, five minutes each transfer cover slips back into the wells of the cell culture plate containing 0.5%osmium tetroxide in distilled water and incubate for 30 minutes at room temperature. Remove the cover slips from the plate and wash three times in distilled water as before.

Then return them to the 24 well plate with 0.5 milliliters of 1%tannic acid in each well and incubate for 30 minutes. Next, wash the cover slips three times five minutes each. Then repeat the fixation with osmium tetroxide and the wash.

Next, cover the test tube. Stand with a new piece of paraform. Dehydrate the cells through a series of incubations in the following ethanol solutions for five minutes each 30, 50, 70, 80, 9100, 100.

And finally, one more time in 100%ethanol. Submerge the rack in an ethanol filled specimen boat. Quickly place the cover slips into the metal rack, making sure that the specimens do not dry out.

Then transfer the specimen boat into a cooled critical point dryer or CPD. Fill the chamber of the CPD with a liquid CO2. Leave the inlet valve fully open to maintain liquid level and as soon as a phase of ethanol is visible under the CO2 phase on the bottom of the chamber, open the drain valve at the bottom.

To remove most of the ethanol. Allow the chamber to flush for about 30 minutes. After flushing, close the inlet valve and lower the liquid level to just below the top of the boat.

Run hot water through the jacket of the CPD. When the temperature reaches 36 to 38 degrees Celsius and the pressure rises to 1, 200 pounds per square inch, the liquid gas boundary line will disappear and the specimens will be above critical point. Once the critical point has been reached, vent the gas off slowly over a five minute period.

To avoid condensation, the temperature should remain around 40 degrees Celsius. During this time, open the door and remove the boat. Confirm that the dehydrating solvent has evaporated.

Stick the cover slips right side up to the adhesive on the aluminum specimen holder and transfer them into the high resolution coating system. Evacuate the coating cylinder to two times 10 to the minus five millibars. Then set the instrument to 1.8 kilovolts and 70 milliamps, and monitor the thickness of the platon carbon layer with a quartz crystal thickness until the layer reaches five nanometers.

Vent the coating cylinder of the high resolution coating system. Then using forceps, remove the samples from the coter. Bring the chamber of the SEM to normal pressure.

Open the door. Then insert the samples into the specimen holder and evacuate the chamber. For SEM analysis.

When fixed at different time, points after stimulation, morphological changes during NETosis can be observed by immunofluorescence microscopy. In this figure, the DNA is shown in blue histon. DNA complex is shown in red and neutrophil elastase.

A granule marker is shown in green during the first phase of NETosis, only the nu nuclear periphery in condensed nuclei. Later, when the chromatin is more decon condensed, most of the nucleus is stained in the decon condensed nets. Staining is homogenous.

This panel of images details the overlap of nuclear and granular staining that occurs with NETosis. This scanning electron micrograph shows non stimulated neutrophils after stimulation with Ella bacterium, the neutrophils flatten out and produce neutrophil extracellular traps.Traps. Here a gel bacterium has been trapped in nets.

Bacteria such as gel are immobilized and killed by the enzymes present in nets. We've just shown you how to isolate neutrophil granulocytes from human blood and to stimulate them in order to induce formation of neutrophil extracellular traps. These can then be further analyzed by immunofluorescence or scanning electron microscopy.

When doing this procedure, it is important to remember that nets are very fragile even after fixation. In order to minimize loss of nets, we take the cover slips with the stimulated neutrophils from the well cavities and do most of the procedure on the surface of buffer drops on. Even then, it's important to move the specimens gently and slowly.

So that's it. Thanks for watching and good luck with your experiments.