Quantifying Human Monocyte Chemotaxis In Vitro and Murine Lymphocyte Trafficking In Vivo

All procedures were approved by Rockefeller University's Institutional Animal Care and Use Committee. All animals were housed under specific pathogen-free conditions.

1. In Vitro Chemotaxis

1. Culture THP-1 cells⁶ in Roswell Park Memorial Institute 1640 (RPMI 1640) supplemented with 10% fetal bovine serum (FBS) and 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES; 25 mM) (complete media), maintaining the density between 0.5-1.0 x 10⁶ cells/mL.
2. In a 35 mm cell culture dish, label 3.5 x 10⁵ THP-1 cells per condition with calcein acetoxymethyl (calcein-AM; 2.5 µM) or other suitable fluorescent dye in complete media at 37 °C for 30 min.
   NOTE: Dyes must be cell membrane-permeable and compatible with live-cell labeling.
3. Pre-warm a plate reader to 37 °C.
   NOTE: In addition to temperature control, bottom-reading functionality is also required. See step 1.10.2 for an alternative to a plate reader for quantification of migrating cells.
4. While cells are labeling, prepare the assay plate and lower chamber.
   1. Use a 24-well tissue culture plate, with one condition per well. Perform each condition in triplicate.
   2. Add 750 µL of Hank's Buffered Salt Solution (HBSS) buffered with 25 mM HEPES and 0.1 % bovine serum albumin (BSA) to the 24-well plate. This serves as the basal control condition.
      1. Use other wells for comparator conditions, always maintaining the total volume in the bottom chamber at 750 µL.
         NOTE: In this example, human monocyte chemoattractant protein-1 (MCP-1) serves as the positive control for chemotaxis⁷.
         1. Assemble the reagent mixture by combining HBSS with 25 mM HEPES, BSA (0.1 %), and MCP-1 (75 ng/mL). Use adult bovine serum (2.7 % v / v) with MCP-1 as an additional positive control.
            NOTE: Chemokine and chemokine concentration will depend on the chemotactic axis under study. Lyophilized MCP-1 is resuspended in distilled water with BSA (0.1%) to make a stock solution with a final concentration of 50 μg/mL. This can be diluted in water to a working solution of 10 μg/mL. Add 5.6 μL of this solution to the lower chamber.
   3. After the composition of the bottom chambers is complete, place a porous insert into each well, ensuring that the tabs of the insert align with the notches of the plate.
      NOTE: Choose the appropriate pore size for the inserts. For monocytes and lymphocytes, a pore size of 3 μm performs well. Some cell types and/or chemokine systems may require a matrix-coated surface for optimal migration. For example, for measuring THP-1 monocyte chemotaxis towards MCP-1, fibronectin or collagen-coated inserts are recommended. For plate reader for quantitation, the insert must have a light-impermeant coating, such as polyethylene terephthalate, to prevent detection of non-migrating cells in the upper chamber⁸.
5. After incubating cells for 30 min with calcein-AM, transfer cell suspension to a 15mL conical tube. Centrifuge cells at 1,000 x g for 2 min. Visually inspect cells to confirm uptake of calcein-AM (Figure 1).
6. Carefully remove the supernatant using a vacuum aspirator. Wash labeled cells in serum-free RPMI 1640 supplemented with HEPES 25 mM. Centrifuge cells at 1,000 x g for 2 min and discard the supernatant.
7. Resuspend cells in serum-free RPMI 1640 supplemented with HEPES 25 mM. Count cells and adjust the volume so that the cells are at a final concentration of 1.0-1.5 x 10⁶ cells/mL.
8. Dispense 250 μL of the cell suspension into the insert (upper chamber).
9. After all upper chambers have been filled, gently lift the plate, and inspect the underside for the presence of air bubbles that may interfere with migration and detection. To remove a bubble, first try removing then re-positioning the insert. If unsuccessful, use a 27 G needle to puncture the bubble and then re-position the insert.
10. Place the plate into the pre-warmed 37 °C plate reader.
    1. Acquire fluorescence readings from the bottom at 1-3 min intervals. When using calcein-AM, set the fluorescence excitation and emission wavelengths at 485 nm and 520 nm, respectively. Depending on the cell type and chemokine system, chemotaxis typically occurs over 1-4 h (Figure 2A).
    2. As an alternative to continuous measurements using a plate-reader, image the wells using an inverted fluorescent microscope (Figure 2B) capable of excitation and detection of light at 485 nm and 520 nm, respectively. Use a low magnification to capture the majority of the underside of the insert. Quantify cells by processing the images with ImageJ or similar software⁹.

2. Adoptive Transfer of Murine Lymphocytes

1. Preparation of donor cells
   1. Anesthetize 8-12 week-old C57BL/6J male donor mice with isoflurane anesthesia (1-3 %) using a vaporizer. Confirm appropriate depth of anesthesia by toe-pinch.
   2. Place the mouse in the supine position. Make a midline incision with a scalpel and locate the spleen in the upper left peritoneal space. Excise the whole spleen and place into complete media (RPMI supplemented with 10% FBS and 25 mM HEPES; 1 mL/spleen) on ice. Euthanize anesthetized donor animals by decapitation.
   3. Grind spleen tissue gently through 40 µm nylon mesh into complete media (1 mL /spleen) using the end of a syringe plunger to make a single-cell suspension and transfer to a 15 mL conical tube.
   4. Add 4 volumes of ammonium chloride potassium (ACK) lysis buffer and incubate for 5 min at room temperature to lyse erythrocytes.
   5. Centrifuge the cells at 1,000 x g for 2 min at room temperature and discard supernatant. If erythrocytes remain in the pellet, resuspend in ACK lysis buffer, incubate for 5 min at room temperature, centrifuge cells at 1,000 x g for 2 min and discard supernatant.
   6. Resuspend the cells in complete media at a concentration of 2-5×10⁷ cells/mL, then strain the cell suspension through a 40 µm nylon mesh to remove cell debris.
   7. Label 1 x 10⁷ cells/recipient mouse with a fluorescent dye compatible with live cells and that will be retained upon subsequent fixation, such as 5-chloromethylfluorescein diacetate (final concentration 2 μM)¹⁰. Incubate at 37 °C for 30 min.
      NOTE: To track more than one cell population, use other fluorescent dyes with additional cell populations. For example, a portion of donor cells can be labeled with 5-(and-6)-(((4-chloromethyl) benzoyl) amino) tetramethylrhodamine (final concentration 2 μM)¹¹.
   8. Add 4 volumes HBSS supplemented with 25 mM HEPES, then centrifuge at 1,000 x g for 2 min and discard supernatant. Resuspend pellet in HBSS supplemented with HEPES 25 mM to a final concentration of 1 x 10⁸ cells/mL.
      1. If using more than one fluorescent dye, keep cell populations separated until immediately prior to injection.
      2. Transfer a small aliquot (10 μL) of cells of each color to a fixative solution (1 mL) for subsequent flow cytometry compensation.
2. Transfer of lymphocytes
   1. Anesthetize 8-12 week-old C57BL/6J male recipient mice with isoflurane (1-3 %) using a vaporizer; confirm depth of anesthesia by toe-pinch. Apply veterinary ointment to animal's eyes to prevent drying.
   2. Briefly vortex donor cells (1 s) and transfer to 1 mL injection syringe with 27 G, 0.5 in needle, combining differentially labeled cell populations, if applicable.
      1. Transfer a small aliquot (10 μL) of mixed cells to a fixative solution (1 mL) for subsequent flow cytometry analyses.
   3. Place donor animals in the prone position. Apply mild caudal pressure to the skin dorsal to the eye, causing the eyeball to protrude slightly.
   4. Direct the needle medially and inject 100 µL of donor cell suspension retro-orbitally into each recipient mouse. Cells can also be injected via the tail vein.
   5. Gradually withdraw the injection needle and place mouse alone in a recovery cage. Monitor animal until it has regained consciousness; once fully recovered return the animal to social housing.
3. Collection and Analysis
   1. After 1 h, anesthetize recipient mice with isoflurane anesthesia (1-3%) using a vaporizer; confirm depth of anesthesia by toe-pinch. Harvest the spleen (or other tissue of interest) (step 2.1.2.) from each recipient mouse and place into complete media (1 mL/recipient). Euthanize anesthetized animals by decapitation.
      NOTE: Longer intervals prior to tissue harvest will increase tissue accumulation through at least 24 h.
   2. Grind spleen tissue gently through 40 µm nylon mesh into complete media using the end of a syringe plunger to make a single-cell suspension and transfer to 15 mL conical tube.
   3. Add 4 volumes of ACK lysis buffer and incubate 5 min at room temperature. Centrifuge the cell suspension at 1,000 x g for 2 min at room temperature and discard supernatant.
   4. Resuspend cells in staining buffer (phosphate-buffered saline supplemented with 1% FBS). Count cells and adjust to a final concentration of 3 x 10 ⁷ cells/mL. Transfer 100 µL of cell suspension to a 96-well round bottom plate.
   5. Stain cells for desired markers (see the Table of Materials) for analysis by flow cytometry. Add fluorophore-conjugated antibodies (see the Table of Materials) against desired markers and incubate at 4 °C for 20 min in the dark. Add 100 µL staining buffer, centrifuge cells at 1,000 x g for 3 min and discard the supernatant.
   6. Resuspend the pellet in 200 µL staining buffer, centrifuge cells at 1,000 x g for 3 min and discard the supernatant.
   7. Resuspend the pellet in 125 µL staining buffer and acquire samples using a flow cytometer using standard procedures.
      NOTE: The green dye is excited by a 488 nm laser and the emission is detected with 505 and 530/30 dichroic filters. The orange dye is excited by a 561 nm and the emission is detected by a 582/15 dichroic filter.
      1. Use saved cells from 2.1.8.2 to compensate for each fluorescent dye.
         NOTE: Using bead-based compensation with fluorophores matching the excitation and emission spectra of the labeling dyes will result in suboptimal compensation.
      2. Use the saved input cells from step 2.2.2.1 to accurately determine the ratio of differentially labeled input cells if using more than one labeled population (Figure 3). Calculate tissue specific migration based on the ratio of labeled to unlabeled cells (Figure 4).