Traditionally, macrophages have been difficult to study in real-time chemotaxis assays because the cells are slow moving. This protocol provides a means to image macrophages migrating in a chemotactic gradient for up to six hours or longer. Compared to implant assays such as transfer assays, this technique has the advantages that macrophage morphology can be observed, and parameters such as cell velocity and chemotactic efficiency can be measured.
Macrophages are involved in many inflammatory diseases, and this technique can be useful to study drugs designed to inhibit chemotaxis. It can also be applied to other cell types such as human monocytes. When attempting this technique for the first time, it is best to practice filling the chemotaxis chambers before working with cells.
One of the most important aspects of the technique is the avoidance of air bubbles. Start by pre-filling the connecting channels of one or two chemotaxis slides with modified RPMI 1640 HEPES Medium prepared according to manuscript directions. Place a slide into a cell culture dish, and set the dish onto an aluminum block heated to 37 degrees Celsius.
Then insert plugs into ports one and four. Use a 200 microliter beveled pipet tip to deposit 15 microliters of the modified RPMI 1640 HEPES Medium into filling port three. Next, insert the pipet tip into port two, and aspirate 15 microliters at a moderately fast rate, which will pre-fill the connecting channel as well as the two flanking supply channels.
Cover the filling ports two and three with caps and place the chemotaxis slide on a rack in a closed humidity chamber within an otherwise dry and carbon dioxide-free incubator at 37 degrees Celsius. To isolate the macrophages, insert a 24 gauge plastic catheter into the peritoneal cavity of a sacrificed three to four month old mouse, and use a five milliliter plastic syringe to lavage the cavity with ice cold Hank's buffered salt solution without calcium or magnesium. After collecting the lavaged medium in a tube, centrifuge it at 300 times G for 6.5 minutes.
Discard the supernatant, and re-suspend the cells in 200 microliters of modified RPMI 1640 medium. Dilute an aliquot of the cells suspension one to 20. Then use a counting device to count the cells.
Dilute the cells to a final concentration of 10 times 10 to the six cells per milliliter, and maintain them at 37 degrees Celsius in a heated aluminum block. Pipet the cell suspension up and down five times to reduce clumping, and gently deposit 10 microliters onto port three of chemotaxis chamber. Place the pipet tip in port two, and slowly draw the cell suspension into the connecting channel.
As soon as the cell suspension has been introduced, remove the plugs at ports one and four to arrest the flow, and place caps on all four filling ports. Then, place the chemotaxis slides in the 37 degree Celsius humidity chamber for two to three hours. Inspect the observation area with an inverted microscope.
Then place plugs into filling ports one and two, and check whether filling port three is filled to the top with medium, and free of air bubbles. If necessary, use a sterile 27 gauge syringe needle to dislodge air bubbles. Next, aspirate 60 microliters of medium with a 100 microliter mechanical pipet, and place the tip into filling port three.
Use the volume setting ring to slowly and steadily inject the medium into the reservoir until it reaches the top of filling port four after one to two minutes. To fill the second reservoir, move the plug from port one and slowly insert it into port three. Aspirate 50 microliters of medium, and place the pipet tip into port four, and slowly inject the medium into the second reservoir so that it reaches the top of filling port one after one to two minutes.
Add 495 microliters of medium to a two milliliter micro-centrifuge tube, and add five microliters of patent blue five. Briefly vortex the mixture, then add 5.4 microliters of Recombinant Mouse Complement C5a and vortex again to mix. Make sure that the shallow depression at the top of port one is medium-free, and deposit 15 microliters of the blue Complement C5a containing medium.
Then insert a 200 microliter pipet tip into filling port four, and slowly rotate the volume setting ring to draw the medium into the opposite reservoir. Draw air into the short vertical column of filling port one until the fluid-air interface is midway in the column. Then plug port one before gently lifting the pipet from port four, making sure that the slide remains in place, and slowly plug port four.
To perform time lapse imaging, place the chemotaxis slide on the stage of an inverted microscope fitted with a stage incubator, and image the observation area with a 10X phase contrast objective lens, focusing on the macrophage lamellipodia. The chemotaxis slide used for time lapse video microscopy of mouse peritoneal macrophages has three chemotaxis chambers, each of which has four filling ports. Cells were seeded in the observation area of each chamber.
And after a two to three hour incubation, the chambers were slowly filled with medium. When the cells in the observation area were inspected, up to two thirds of the cells had been washed out. Generally, weakly adherent B-1 cells were washed out, and the remaining cells were predominantly macrophages.
To distinguish the macrophages from the B cells, freshly isolated cells were labeled with specific antibodies. The macrophages were labeled with green fluorescence, the B cells with red, and the cell nuclei with blue. Macrophage migration was investigated by introducing Complement C5a, a chemoattractant, to one of the two reservoirs.
The migration tracks of macrophages migrating in a Complement C5 gradient were recorded. The start point of each migration track was normalized to X equals zero and Y equals zero to create a migration plot. Then, cell velocity and chemotactic efficiency of individual macrophages were calculated.
This technique has been useful to study the roles of G protein subunits, and Rho GTPases, and macrophage motility in chemotaxis.
