This work was supported by the Bernard L. Schwartz Program for Physician Scientists at The Rockefeller University, the Robertson Therapeutic Development Fund at The Rockefeller University, and the Sackler Center for Biomedicine and Nutrition Research at The Rockefeller University.

All procedures were approved by Rockefeller University&#39;s Institutional Animal Care and Use Committee. All animals were housed under specific pathogen-free conditions.
1. In Vitro Chemotaxis
<ol>
	<li>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.</li>
	<li>In a 35 mm cell culture dish, label 3.5 x 105 THP-1 cells per condition with calcein acetoxymethyl (calcein-AM; 2.5 &#181;M) or other suitable fluorescent dye in complete media at 37 &#176;C for 30 min. 
	NOTE: Dyes must be cell membrane-permeable and compatible with live-cell labeling.</li>
	<li>Pre-warm a plate reader to 37 &#176;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.</li>
	<li>While cells are labeling, prepare the assay plate and lower chamber.
	<ol>
		<li>Use a 24-well tissue culture plate, with one condition per well. Perform each condition in triplicate.</li>
		<li>Add 750 &#181;L of Hank&#39;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.
		<ol>
			<li>Use other wells for comparator conditions, always maintaining the total volume in the bottom chamber at 750 &#181;L. 
			NOTE: In this example, human monocyte chemoattractant protein-1 (MCP-1) serves as the positive control for chemotaxis7.
			<ol>
				<li>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 &#956;g/mL. This can be diluted in water to a working solution of 10 &#956;g/mL. Add 5.6 &#956;L of this solution to the lower chamber.</li>
			</ol>
			</li>
		</ol>
		</li>
		<li>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 &#956;m 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.</li>
	</ol>
	</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>Dispense 250 &#956;L of the cell suspension into the insert (upper chamber).</li>
	<li>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.</li>
	<li>Place the plate into the pre-warmed 37 &#176;C plate reader.
	<ol>
		<li>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).</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
2. Adoptive Transfer of Murine Lymphocytes
<ol>
	<li>Preparation of donor cells
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Grind spleen tissue gently through 40 &#181;m 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.</li>
		<li>Add 4 volumes of ammonium chloride potassium (ACK) lysis buffer and incubate for 5 min at room temperature to lyse erythrocytes.</li>
		<li>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.</li>
		<li>Resuspend the cells in complete media at a concentration of 2-5x107 cells/mL, then strain the cell suspension through a 40 &#181;m nylon mesh to remove cell debris.</li>
		<li>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 &#956;M)10. Incubate at 37 &#176;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 &#956;M)11.</li>
		<li>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.
		<ol>
			<li>If using more than one fluorescent dye, keep cell populations separated until immediately prior to injection.</li>
			<li>Transfer a small aliquot (10 &#956;L) of cells of each color to a fixative solution (1 mL) for subsequent flow cytometry compensation.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Transfer of lymphocytes
	<ol>
		<li>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&#39;s eyes to prevent drying.</li>
		<li>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.
		<ol>
			<li>Transfer a small aliquot (10 &#956;L) of mixed cells to a fixative solution (1 mL) for subsequent flow cytometry analyses.</li>
		</ol>
		</li>
		<li>Place donor animals in the prone position. Apply mild caudal pressure to the skin dorsal to the eye, causing the eyeball to protrude slightly.</li>
		<li>Direct the needle medially and inject 100 &#181;L of donor cell suspension retro-orbitally into each recipient mouse. Cells can also be injected via the tail vein.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Collection and Analysis
	<ol>
		<li>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.</li>
		<li>Grind spleen tissue gently through 40 &#181;m 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.</li>
		<li>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.</li>
		<li>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 &#181;L of cell suspension to a 96-well round bottom plate.</li>
		<li>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 &#176;C for 20 min in the dark. Add 100 &#181;L staining buffer, centrifuge cells at 1,000 x g for 3 min and discard the supernatant.</li>
		<li>Resuspend the pellet in 200 &#181;L staining buffer, centrifuge cells at 1,000 x g for 3 min and discard the supernatant.</li>
		<li>Resuspend the pellet in 125 &#181;L 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.
		<ol>
			<li>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.</li>
			<li>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).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Griffith, J. W., Sokol, C. L., Luster, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemokines+and+chemokine+receptors:+positioning+cells+for+host+defense+and+immunity.">Chemokines and chemokine receptors: positioning cells for host defense and immunity.</a> Annu Rev Immunol. 32, 659-702 (2014).</li><li>Springer, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Traffic+signals+for+lymphocyte+recirculation+and+leukocyte+emigration:+the+multistep+paradigm.">Traffic signals for lymphocyte recirculation and leukocyte emigration: the multistep paradigm.</a> Cell. 76 (2), 301-314 (1994).</li><li>Ponda, M. P., Breslow, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Serum+stimulation+of+CCR7+chemotaxis+due+to+coagulation+factor+XIIa-dependent+production+of+high-molecular-weight+kininogen+domain+5.">Serum stimulation of CCR7 chemotaxis due to coagulation factor XIIa-dependent production of high-molecular-weight kininogen domain 5.</a> Proc Natl Acad Sci U S A. 113 (45), E7059-E7068 (2016).</li><li>Boyden, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+chemotactic+effect+of+mixtures+of+antibody+and+antigen+on+polymorphonuclear+leucocytes.">The chemotactic effect of mixtures of antibody and antigen on polymorphonuclear leucocytes.</a> J Exp Med. 115, 453-466 (1962).</li><li>Chen, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Boyden+chamber+assay.">Boyden chamber assay.</a> Methods Mol Biol. 294, 15-22 (2005).</li><li>Tsuchiya, S., Yamabe, M., Yamaguchi, Y., Kobayashi, Y., Konno, T., Tada, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+and+characterization+of+a+human+acute+monocytic+leukemia+cell+line+(THP-1).">Establishment and characterization of a human acute monocytic leukemia cell line (THP-1).</a> Int J Cancer. 26 (2), 171-176 (1980).</li><li>Leonard, E. J., Yoshimura, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+monocyte+chemoattractant+protein-1+(MCP-1).">Human monocyte chemoattractant protein-1 (MCP-1).</a> Immunol Today. 11 (3), 97-101 (1990).</li><li>Crouch, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+automated+fluorescence+based+assay+of+neurite+formation.">An automated fluorescence based assay of neurite formation.</a> J Neurosci Methods. 104 (1), 87-91 (2000).</li><li>Schneider, C. A., Rasband, W. S., Eliceiri, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NIH+Image+to+ImageJ:+25+years+of+image+analysis.">NIH Image to ImageJ: 25 years of image analysis.</a> Nat Methods. 9 (7), 671-675 (2012).</li><li>Poot, M., Kavanagh, T. J., Kang, H. C., Haugland, R. P., Rabinovitch, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+cytometric+analysis+of+cell+cycle-dependent+changes+in+cell+thiol+level+by+combining+a+new+laser+dye+with+Hoechst+33342.">Flow cytometric analysis of cell cycle-dependent changes in cell thiol level by combining a new laser dye with Hoechst 33342.</a> Cytometry. 12 (2), 184-187 (1991).</li><li>West, C. A., He, C., Su, M., Swanson, S. J., Mentzer, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aldehyde+fixation+of+thiol-reactive+fluorescent+cytoplasmic+probes+for+tracking+cell+migration.">Aldehyde fixation of thiol-reactive fluorescent cytoplasmic probes for tracking cell migration.</a> J Histochem Cytochem. 49 (4), 511-518 (2001).</li><li>Kim, C. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+greater+chemotactic+network+for+lymphocyte+trafficking:+chemokines+and+beyond.">The greater chemotactic network for lymphocyte trafficking: chemokines and beyond.</a> Curr Opin Hematol. 12 (4), 298-304 (2005).</li></ol>

The authors have nothing to disclose

Quantification of immune cell migration can be accomplished using simple and rapid assays both in vitro and in vivo. We demonstrate the in vitro chemotaxis of human monocytes in response to a MCP-1 gradient and augmentation by serum. In vivo, donor murine splenocytes were differentially labeled and following adoptive transfer, were recovered from recipient animals.
Using a plate reader has the advantage of sampling several time points (as frequently as every...

Robust immunity requires the complex temporal and spatial coordination of a myriad of cell types in order to respond appropriately to injury, infection and generate self-tolerance. Several dozen chemokine receptors and their corresponding ligands have been discovered and characterized providing molecular mechanisms by which specific cells can be directed into a specific tissue at a specific time. Thus, studying chemotaxis and migration is an indispensable component of immunology research. Indeed, the described in vitro assay was recently used as a screening tool to identify a chemotactic cofactor that accelerates chemotaxis of T-cells toward C-C chemokines 19 and 213. The purpose of the methods described here are to permit quantitative assessments of immune cell chemotaxis in vitro and in vivo.
The Boyden (cell migration and invasion) chamber assay is an inexpensive, reproducible, and rapid method for assessing cell migration4,5. In the standard assay, the upper chamber is seeded with cells, and is separated by a porous insert from a lower chamber, into which the cells migrate. At the desired time, cells that have migrated to the underside of the insert can be fixed and stained for quantitation by light microscopy. However, such measurements constrain data collection to a single end point, which precludes dynamic data collection and can require extensive optimization to determine the optimal time point for analysis. Here, several adaptations are described that permit real-time, quantitative and multiplexed measurements of chemotaxis in vitro.
For in vivo studies, a functional end point is used, namely the specific accumulation of cells in a given tissue compartment. Pre-labeled donor cells are introduced into recipient animals through adoptive transfer. These donor cells can subsequently be identified by flow cytometry after recipient tissue harvest. Also presented is a co-labeling strategy that allows for the determination of trafficking of different cell types within a single recipient animal. This method eliminates the inter-animal variation from cell injection, and accounts for physiologic inter-animal variability.

<table>
	<tbody>
		<tr>
			<td>RPMI-1640</td>
			<td>Thermo Fisher Scientific</td>
			<td>11875-093</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Thermo Fisher Scientific</td>
			<td>15630-080</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>ATCC</td>
			<td>30-2020</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture flask</td>
			<td>Corning</td>
			<td>353136</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcein AM</td>
			<td>Thermo Fisher Scientific</td>
			<td>C1430</td>
			<td>Excitation: 485nm, Emission: 520nm</td>
		</tr>
		<tr>
			<td>Cell culture dish</td>
			<td>Corning</td>
			<td>1007</td>
			<td></td>
		</tr>
		<tr>
			<td>24 well plate</td>
			<td>Corning</td>
			<td>353504</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoroblok Fibronectin Insert</td>
			<td>Corning</td>
			<td>CB354597</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL conical tube</td>
			<td>Corning</td>
			<td>352097</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Cell Signaling Technology</td>
			<td>9998S</td>
			<td></td>
		</tr>
		<tr>
			<td>Human Recombinant MCP-1</td>
			<td>Peprotech</td>
			<td>300-04</td>
			<td></td>
		</tr>
		<tr>
			<td>Adult bovine Serum</td>
			<td>Sigma</td>
			<td>B9433</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS with calcium and magnesium; no phenol red</td>
			<td>Thermo Fisher Scientific</td>
			<td>14025-092</td>
			<td></td>
		</tr>
		<tr>
			<td>SpectraMax M2e plate reader</td>
			<td>Molecular Devices</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus IX71 inverted fluorescence microscope</td>
			<td>Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isothesia</td>
			<td>Henry Schein animal health</td>
			<td>11695-6776-2</td>
			<td>Isofluorane anesthesia</td>
		</tr>
		<tr>
			<td>Cell strainer 40um nylon</td>
			<td>Falcon Corning</td>
			<td>352340</td>
			<td></td>
		</tr>
		<tr>
			<td>ACK lysing buffer</td>
			<td>Quality Biological</td>
			<td>118-156-101</td>
			<td></td>
		</tr>
		<tr>
			<td>CellTracker Orange CMTMR Dye</td>
			<td>Thermo Fisher Scientific</td>
			<td>C2927</td>
			<td>Excitation: 541nm, Emission: 565nm</td>
		</tr>
		<tr>
			<td>CellTracker Green CMFDA Dye</td>
			<td>Thermo Fisher Scientific</td>
			<td>C2925</td>
			<td>Excitation: 485nm, Emission: 520nm</td>
		</tr>
		<tr>
			<td>5ml syringe</td>
			<td>BD Syringe</td>
			<td>309646</td>
			<td></td>
		</tr>
		<tr>
			<td>1ml TB syringe</td>
			<td>BD Syringe</td>
			<td>309625</td>
			<td></td>
		</tr>
		<tr>
			<td>THP-1 cell line</td>
			<td>ATCC</td>
			<td>TIB-202</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 421 anti-mouse CD3 Antibody</td>
			<td>Biolegend</td>
			<td>100228</td>
			<td>Excitation: 405nm, Emission: 421nm</td>
		</tr>
		<tr>
			<td>Brilliant Violet 605 anti-mouse CD19 Antibody</td>
			<td>Biolegend</td>
			<td>115540</td>
			<td>Excitation: 405nm, Emission: 603nm</td>
		</tr>
		<tr>
			<td>96-well round bottom plate</td>
			<td>Corning</td>
			<td>353077</td>
			<td></td>
		</tr>
		<tr>
			<td>LSRII</td>
			<td>BD Biosciences</td>
			<td></td>
			<td>Flow cytometer</td>
		</tr>
	</tbody>
</table>

quantifying human monocyte chemotaxis in vitro and murine lymphocyte trafficking in vivo

Chemotaxis is migration along a specific chemical gradient1. Chemokines are chemotactic cytokines that promote cellular trafficking with anatomic and temporal specificity2. Chemotaxis is a critical function of lymphocytes and other immune cells that can be quantitatively assessed in vitro. This manuscript describes methods that permit the evaluation of chemotaxis, both in vitro and in vivo, for diverse cell types including cell lines and native cells. The in vitro, plate-based format permits the comparison of several conditions simultaneously in real-time, and can be completed within 1-4 h. In vitro assay conditions can be manipulated to introduce agonists and antagonists, as well as differentiate chemotaxis from chemokinesis, which is random movement. For in vivo trafficking assessments, immune cells can be labeled with multiple fluorescent dyes and used for adoptive transfer. The differential labeling of cells allows for mixed cell populations to be introduced into the same animal, thereby decreasing variance and reducing the number of animals required for an adequately powered experiment. Migration into lymphoid tissue occurs in as little as 1 h, and multiple tissue compartments can be sampled. Flow cytometry following tissue harvest allows for a rapid and quantitative analysis of the migratory patterns of multiple cell types.

When using calcein-AM dye, visual inspection of cells will confirm label uptake (Figure 1). Automated fluorescent readings will track migration as cells transit onto the underside of the insert over time. These data clearly show an induction of cell migration towards MCP-1, as well as an augmentation of this response by serum (Figure 2A). Depending on the strength of the migratory stimulus, there is a lag of at least 15 min prior...

Watch this Scientific Journal Video about Quantifying Human Monocyte Chemotaxis In Vitro and Murine Lymphocyte Trafficking In Vivo at JoVE.com

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

Protocols for quantitative assessment of lymphocyte chemotaxis and migration are important tools for immunology research. Here, an in vitro protocol is described that permits real-time, multiplexed evaluation of cell migration, as well as a complementary in vivo technique enabling tracking of native cells to spleen.

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

Chemotaxis is migration along a specific chemical gradient1. Chemokines are chemotactic cytokines that promote cellular trafficking with anatomic and ...

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Immunology and Infection

7.1K Views.  The Rockefeller University. Protocols for quantitative assessment of lymphocyte chemotaxis and migration are important tools for immunology research. Here, an in vitro protocol is described that permits real-time, multiplexed evaluation of cell migration, as well as a complementary in vivo technique enabling tracking of native cells to spleen.

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

Human T Lymphocyte Isolation, Culture and Analysis of Migration In Vitro

The migration of T lymphocytes involves the adhesive interaction of cell surface integrins with ligands expressed on other cells or with extracellular matrix proteins. The precise spatiotemporal activation of integrins from a low affinity state to a high affinity state at the cell leading edge is important for T lymphocyte migration 1. Likewise, retraction of the cell trailing edge, or uropod, is a necessary step in maintaining persistent integrin-dependent T lymphocyte motility 2. Many therapeutic approaches to autoimmune or inflammatory diseases target integrins as a means to inhibit the excessive recruitment and migration of leukocytes 3. To study the molecular events that regulate human T lymphocyte migration, we have utilized an in vitro system to analyze cell migration on a two-dimensional substrate that mimics the environment that a T lymphocyte encounters during recruitment from the vasculature. T lymphocytes are first isolated from human donors and are then stimulated and cultured for seven to ten days. During the assay, T lymphocytes are allowed to adhere and migrate on a substrate coated with intercellular adhesion molecule-1 (ICAM-1), a ligand for integrin LFA-1, and stromal cell-derived factor-1 (SDF-1). Our data show that T lymphocytes exhibit a migratory velocity of ~15 &mu;m/min. T lymphocyte migration can be inhibited by integrin blockade 1 or by inhibitors of the cellular actomyosin machinery that regulates cell migration 2.

Human T Lymphocyte Isolation, Culture and Analysis of Migration In Vitro

T lymphocyte migration occurs during homing to lymphoid organs, exit from the vasculature, and entering into peripheral tissues. Here, we describe a protocol that can be used to analyze T lymphocyte migration in vitro.

The migration of T lymphocytes involves the adhesive interaction of cell surface integrins with ligands expressed on other cells or with extracellular ...

An In vitro Co-infection Model to Study Plasmodium falciparum-HIV-1 Interactions in Human Primary Monocyte-derived Immune Cells

Plasmodium falciparum, the causative agent of the deadliest form of malaria, and human immunodeficiency virus type-1 (HIV-1) are among the most important health problems worldwide, being responsible for a total of 4 million deaths annually1. Due to their extensive overlap in developing regions, especially Sub-Saharan Africa, co-infections with malaria and HIV-1 are common, but the interplay between the two diseases is poorly understood. Epidemiological reports have suggested that malarial infection transiently enhances HIV-1 replication and increases HIV-1 viral load in co-infected individuals2,3. Because this viremia stays high for several weeks after treatment with antimalarials, this phenomenon could have an impact on disease progression and transmission.
The cellular immunological mechanisms behind these observations have been studied only scarcely. The few in vitro studies investigating the impact of malaria on HIV-1 have demonstrated that exposure to soluble malarial antigens can increase HIV-1 infection and reactivation in immune cells. However, these studies used whole cell extracts of P. falciparum schizont stage parasites and peripheral blood mononuclear cells (PBMC), making it hard to decipher which malarial component(s) was responsible for the observed effects and what the target host cells were4,5. Recent work has demonstrated that exposure of immature monocyte-derived dendritic cells to the malarial pigment hemozoin increased their ability to transfer HIV-1 to CD4+ T cells6,7, but that it decreased HIV-1 infection of macrophages8. To shed light on this complex process, a systematic analysis of the interactions between the malaria parasite and HIV-1 in different relevant human primary cell populations is critically needed.
Several techniques for investigating the impact of HIV-1 on the phagocytosis of micro-organisms and the effect of such pathogens on HIV-1 replication have been described. We here present a method to investigate the effects of P. falciparum-infected erythrocytes on the replication of HIV-1 in human primary monocyte-derived macrophages. The impact of parasite exposure on HIV-1 transcriptional/translational events is monitored by using single cycle pseudotyped viruses in which a luciferase reporter gene has replaced the Env gene while the effect on the quantity of virus released by the infected macrophages is determined by measuring the HIV-1 capsid protein p24 by ELISA in cell supernatants.

An In vitro Co-infection Model to Study Plasmodium falciparum-HIV-1 Interactions in Human Primary Monocyte-derived Immune Cells

We have developed an in vitro malaria-HIV-1 co-infection model to study the impact of Plasmodium falciparum on the HIV-1 replicative cycle in human primary monocyte-derived macrophages. This versatile system can easily be adapted to other primary cell types susceptible to HIV-1 infection.

Plasmodium falciparum, the causative agent  of the deadliest form of malaria, and human immunodeficiency virus type-1 (HIV-1) are among the most important ...

Quantitative In vitro Assay to Measure Neutrophil Adhesion to Activated Primary Human Microvascular Endothelial Cells under Static Conditions

The vascular endothelium plays an integral part in the inflammatory response. During the acute phase of inflammation, endothelial cells (ECs) are activated by host mediators or directly by conserved microbial components or host-derived danger molecules. Activated ECs express cytokines, chemokines and adhesion molecules that mobilize, activate and retain leukocytes at the site of infection or injury. Neutrophils are the first leukocytes to arrive, and adhere to the endothelium through a variety of adhesion molecules present on the surfaces of both cells. The main functions of neutrophils are to directly eliminate microbial threats, promote the recruitment of other leukocytes through the release of additional factors, and initiate wound repair. Therefore, their recruitment and attachment to the endothelium is a critical step in the initiation of the inflammatory response. In this report, we describe an in vitro neutrophil adhesion assay using calcein AM-labeled primary human neutrophils to quantitate the extent of microvascular endothelial cell activation under static conditions. This method has the additional advantage that the same samples quantitated by fluorescence spectrophotometry can also be visualized directly using fluorescence microscopy for a more qualitative assessment of neutrophil binding.

Quantitative In vitro Assay to Measure Neutrophil Adhesion to Activated Primary Human Microvascular Endothelial Cells under Static Conditions

Neutrophil adherence to the activated endothelium at sites of infection is an integral component of the host's inflammatory response. Described in this report is a neutrophil binding assay that allows for the in vitro quantitation of primary human neutrophil binding to endothelial cells activated by inflammatory mediators under static conditions.

The vascular  endothelium plays an integral part in the inflammatory response. During the  acute phase of inflammation, endothelial cells (ECs) are ...

Lymphocyte Isolation from Human Skin for Phenotypic Analysis and Ex Vivo Cell Culture

Human skin has an important barrier function and contains various immune cells that contribute to tissue homeostasis and protection from pathogens. As the skin is relatively easy to access, it provides an ideal platform to study peripheral immune regulatory mechanisms. Immune resident cells in healthy skin conduct immunosurveillance, but also play an important role in the development of inflammatory skin disorders, such as psoriasis. Despite emerging insights, our understanding of the biology underlying various inflammatory skin diseases is still limited. There is a need for good quality (single) cell populations isolated from biopsied skin samples. So far, isolation procedures have been seriously hampered by a lack of obtaining a sufficient number of viable cells. Isolation and subsequent analysis have also been affected by the loss of immune cell lineage markers, due to the mechanical and chemical stress caused by the current dissociation procedures to obtain single cell suspension. Here, we describe a modified method to isolate T cells from both healthy and involved psoriatic human skin by combining mechanical skin dissociation using an automated tissue dissociator and collagenase treatment. This methodology preserves expression of most immune lineage markers such as CD4, CD8, Foxp3 and CD11c upon the preparation of single cell suspensions. Examples of successful CD4+ T cell isolation and subsequent phenotypic and functional analysis are shown.

Lymphocyte Isolation from Human Skin for Phenotypic Analysis and Ex Vivo Cell Culture

We describe a protocol to efficiently isolate skin resident T cells from human skin biopsies. This protocol yields sufficient numbers of viable human skin resident lymphocytes for flow cytometric analysis and ex vivo culture.

Human skin has an important barrier function and contains various immune cells that contribute to tissue homeostasis and protection from pathogens. As the ...

Measuring Granulocyte and Monocyte Phagocytosis and Oxidative Burst Activity in Human Blood

The granulocyte and monocyte phagocytosis and oxidative burst (OB) activity assay can be used to study the innate immune system. This manuscript provides the necessary methodology to add this assay to an exercise immunology arsenal. The first step in this assay is to prepare two aliquots ("H" and "F") of whole blood (heparin). Then, dihydroethidium is added to the H aliquot, and both aliquots are incubated in a warm water bath followed by a cold water bath. Next, Staphylococcus aureus (S. aureus) is added to the H aliquot and fluorescein isothiocyanate-labeled S. aureus is added to the F aliquot (bacteria:phagocyte = 8:1), and both aliquots are incubated in a warm water bath followed by a cold water bath. Then, trypan blue is added to each aliquot to quench extracellular fluorescence, and the cells are washed with phosphate-buffered saline. Next, the red blood cells are lysed, and the white blood cells are fixed. Finally, a flow cytometer and appropriate analysis software are used to measure granulocyte and monocyte phagocytosis and OB activity. This assay has been used for over 20 years. After heavy and prolonged exertion, athletes experience a significant but transient increase in phagocytosis and an extended decrease in OB activity. The post-exercise increase in phagocytosis is correlated with inflammation. In contrast to normal weight individuals, granulocyte and monocyte phagocytosis is chronically elevated in overweight and obese participants, and is modestly correlated with C-reactive protein. In summary, this flow cytometry-based assay measures the phagocytosis and OB activity of phagocytes and can be used as an additional measure of exercise- and obesity-induced inflammation.

The purpose of this manuscript is to present a method for measuring monocyte and granulocyte phagocytosis and oxidative burst activity in human blood samples.

The granulocyte and monocyte phagocytosis and oxidative burst (OB) activity assay can be used to study the innate immune system. This manuscript provides ...

Murine Lymphocyte Labeling by 64Cu-Antibody Receptor Targeting for In Vivo Cell Trafficking by PET/CT

This protocol illustrates the production of 64Cu and the chelator conjugation/radiolabeling of a monoclonal antibody (mAb) followed by murine lymphocyte cell culture and 64Cu-antibody receptor targeting of the cells. In vitro evaluation of the radiolabel and non-invasive in vivo cell tracking in an animal model of an airway delayed-type hypersensitivity reaction (DTHR) by PET/CT are described.
In detail, the conjugation of a mAb with the chelator 1,4,7,10-tetraazacyclododecane-1,4,7,10-tetraacetic acid (DOTA) is shown. Following the production of radioactive 64Cu, radiolabeling of the DOTA-conjugated mAb is described. Next, the expansion of chicken ovalbumin (cOVA)-specific CD4+ interferon (IFN)-&#947;-producing T helper cells (cOVA-TH1) and the subsequent radiolabeling of the cOVA-TH1 cells are depicted. Various in vitro techniques are presented to evaluate the effects of 64Cu-radiolabeling on the cells, such as the determination of cell viability by trypan blue exclusion, the staining for apoptosis with Annexin V for flow cytometry, and the assessment of functionality by IFN-&#947; enzyme-linked immunosorbent assay (ELISA). Furthermore, the determination of the radioactive uptake into the cells and the labeling stability are described in detail. This protocol further describes how to perform cell tracking studies in an animal model for an airway DTHR and, therefore, the induction of cOVA-induced acute airway DHTR in BALB/c mice is included. Finally, a robust PET/CT workflow including image acquisition, reconstruction, and analysis is presented.
The 64Cu-antibody receptor targeting approach with subsequent receptor internalization provides high specificity and stability, reduced cellular toxicity, and low efflux rates compared to common PET-tracers for cell labeling, e.g.64Cu-pyruvaldehyde bis(N4-methylthiosemicarbazone) (64Cu-PTSM). Finally, our approach enables non-invasive in vivo cell tracking by PET/CT with an optimal signal-to-background ratio for 48 h. This experimental approach can be transferred to different animal models and cell types with membrane-bound receptors that are internalized.

Murine Lymphocyte Labeling by 64Cu-Antibody Receptor Targeting for In Vivo  Cell Trafficking by PET/CT

Following the preparation of a 64Cu-modified monoclonal antibody binding to a transgenic murine T cell receptor, T cells are radiolabeled in vivo, analyzed for viability, functionality, labeling stability and apoptosis, and adoptively transferred into mice with an airway delayed-type hypersensitivity reaction for non-invasive imaging by positron emission tomography/computed tomography (PET/CT).

This protocol illustrates the production of 64Cu and the chelator conjugation/radiolabeling of a monoclonal antibody (mAb) followed by murine lymphocyte ...

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte extravasation might result in pathophysiological changes in the CNS. Thus, investigation of lymphocyte migration into the CNS is important to understand inflammatory CNS diseases and to develop new therapy approaches. Here we present an in vitro model of the human blood-brain barrier to study lymphocyte extravasation. Human brain microvascular endothelial cells (HBMEC) are confluently grown on a porous polyethylene terephthalate transwell insert to mimic the endothelium of the blood-brain barrier. Barrier function is validated by zonula occludens immunohistochemistry, transendothelial electrical resistance (TEER) measurements as well as analysis of evans blue permeation. This model allows investigation of the diapedesis of rare lymphocyte subsets such as CD56brightCD16dim/- NK cells. Furthermore, the effects of other cells, cytokines and chemokines, disease-related alterations, and distinct treatment regimens on the migratory capacity of lymphocytes can be studied. Finally, the impact of inflammatory stimuli as well as different treatment regimens on the endothelial barrier can be analyzed.

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Here, we describe a human blood-brain barrier model enabling to investigate lymphocyte transmigration into the central nervous system in vitro.

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte ...

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and their important role in lung development and function, as well as their lung-localized responses to infection and inflammation. To date, no unified method for identification, isolation, and handling of alveolar macrophages from humans and mice exists. Such a method is needed for studies on these important innate immune cells in various experimental settings. The method described here, which can be easily adopted by any laboratory, is a simplified approach to harvesting alveolar macrophages from bronchoalveolar lavage fluid or from lung tissue and maintaining them in vitro. Because alveolar macrophages primarily occur as adherent cells in the alveoli, the focus of this method is on dislodging them prior to harvest and identification. The lung is a highly vascularized organ, and various cell types of myeloid and lymphoid origin inhabit, interact, and are influenced by the lung microenvironment. By using the set of surface markers described here, researchers can easily and unambiguously distinguish alveolar macrophages from other leukocytes, and purify them for downstream applications. The culture method developed herein supports both human and mouse alveolar macrophages for in vitro growth, and is compatible with cellular and molecular studies.

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

This communication describes methodologies for isolation and culture of alveolar macrophages from humans and murine models for experimental purposes.

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and ...

Simultaneous Study of the Recruitment of Monocyte Subpopulations Under Flow In Vitro

The recruitment of monocytes from the blood to targeted peripheral tissues is critical to the inflammatory process during tissue injury, tumor development and autoimmune diseases. This is facilitated through a process of capture from free flow onto the luminal surface of activated endothelial cells, followed by their adhesion and transendothelial migration (transmigration) into the underlying affected tissue. However, the mechanisms that support the preferential and context-dependent recruitment of monocyte subpopulations are still not fully understood. Therefore, we have developed a method that allows the recruitment of different monocyte subpopulations to be simultaneously visualized and measured under flow. This method, based on time-lapse confocal imaging, allows for the unambiguous distinction between adherent and transmigrated monocytes. Here, we describe how this method can be used to simultaneously study the recruitment cascade of pro-angiogenic and non-angiogenic monocytes in vitro. Furthermore, this method can be extended to study the different steps of recruitment of up to three monocyte populations.

Here, we present an integrated protocol that measures monocyte subpopulation trafficking under flow in vitro by use of specific surface markers and confocal fluorescence microscopy. This protocol can be used to explore sequential recruitment steps as well as to profile other leukocyte subtypes using other specific surface markers.

The recruitment of monocytes from the blood to targeted peripheral tissues is critical to the inflammatory process during tissue injury, tumor development ...

Quantification of Monocyte Chemotactic Activity In Vivo and Characterization of Blood Monocyte Derived Macrophages

Tissue homeostasis and repair are critically dependent on the recruitment of monocyte-derived macrophages. Both under- and over-recruitment of monocyte-derived macrophages can impair wound healing. We showed that high fat and high sugar diets promote monocyte priming and dysfunction, converting healthy blood monocytes into a hyper chemotactic phenotype poised to differentiate into macrophages with dysregulated activation profiles and impaired phenotypic plasticity. The over-recruitment of monocyte-derived macrophages and recruitment of macrophages with dysregulated activation profiles is believed to be a major contributor to the development of chronic inflammatory diseases associated with metabolic disorders, including atherosclerosis and obesity. The goal of this protocol is to quantify the chemotactic activity of blood monocytes as a biomarker for monocyte priming and dysfunction and to characterize the macrophage phenotype blood monocytes are poised to differentiate into in these mouse models. Using single cell Western blot analysis, we show that after 24 h 33%of cells recruited into MCP-1-loaded basement membrane-derived gel plugs injected into mice are monocytes and macrophages; 58% after day 3. However, on day 5, monocyte and macrophage numbers were significantly decreased. Finally, we show that this assays also allows for the isolation of live macrophages from the surgically retrieved basement membrane-derived gel plugs, which can then be subjected to subsequent characterization by single cell Western blot analysis.

Here we present a protocol to quantify the chemotactic activity of blood monocytes in mouse models, to assess the effects of nutritional, pharmacological and genetic interventions on blood monocyte and to characterize the blood monocytes derived macrophages in mouse models using monocyte-chemoattractant protein-1 (MCP-1)-loaded basement membrane-derived gel plugs.