This work was supported by a grant (HA 3271/3-2) from the DFG (Deutsche Forschungsgemeinschaft).

The protocols follow the guidelines of our local research ethics committee, as well as the animal care guidelines.
NOTE: Figure 1 shows a workflow of the chemotaxis assay.
1. Prefilling Chemotaxis Slides
<ol>
	<li>Prefill the 1 mm wide and 2 mm long connecting channels of one or two chemotaxis slides using modified RPMI 1640 HEPES medium, consisting of bicarbonate-free RPMI 1640 medium containing 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 10% heat-inactivated fetal bovine serum (FBS), and antibiotics such as penicillin (100 U/mL) and streptomycin (100 &#181;g/mL), prepared by diluting 100x penicillin/streptomycin, and 1 &#181;g/mL lipopolysaccharide (from E. coli), and a Toll-like receptor 4 ligand used to activate the cells.
	<ol>
		<li>Place a chemotaxis slide (Figure 2A) into a round (10 cm diameter) cell culture dish, both preheated to 37 &#176;C, and set the dish onto a heated (37 &#176;C) aluminum block. Insert plugs into ports 1 and 4 (Figure 2B). 
		NOTE: An aluminum block maintained at 37 &#176;C and placed inside the laminar flow hood is useful for preparing chemotaxis chambers. Ideally, the heated block should provide a flat working area and wells for various tubes, such as 50 mL tubes and 2 mL microcentrifuge tubes.</li>
		<li>Using a 10&#8211;200 &#181;L pipette tip with a beveled tip, deposit 15 &#181;L of modified RPMI 1640 HEPES medium into filling port 3 (Figure 2B). Next, with the volume still set at 15 &#181;L and the control button of the (2&#8211;20 &#181;L volume) pipette depressed, insert the pipette tip into port 2 and aspirate 15 &#181;L at a moderately fast rate (Figure 2B). This will prefill the 1 mm x 2 mm connecting channel (observation area), as well as the two flanking supply channels (between the central observation area and ports 2 and 3, respectively). Cover filling ports 2 and 3 with caps.</li>
		<li>After prefilling, place the chemotaxis slides onto a rack kept in a closed humidity chamber within an otherwise dry and CO2-free incubator at 37 &#176;C. 
		NOTE: It is important to use the correct pipette tip for filling the chemotaxis slide. A beveled pipette tip wedges into the top of the filling port, whereas commonly used pointed pipette tips can be inserted more deeply into the filling port and may greatly increase resistance to fluid flow.</li>
	</ol>
	</li>
</ol>
2. Isolation of Mouse Resident Peritoneal Macrophages
<ol>
	<li>Sacrifice a 3&#8211;4 month-old mouse using a high concentration of the volatile anesthetic isoflurane (&#62;5% in air) or carbon dioxide25, followed by cervical dislocation. Loss of the righting reflex in rodents correlates with loss of consciousness in humans26. Clean the abdomen of the mouse with 80% ethanol in water and then make a 1&#8211;2 cm midline skin incision using surgical scissors with blunt tips. Peel back the skin to expose the underlying abdominal wall.</li>
	<li>Insert a 24 G plastic catheter into the peritoneal cavity. Using a 5 mL plastic syringe, lavage the cavity using 2 x 4.5 mL ice-cold Hank&#8217;s buffered salt solution (HBSS), without Ca2+ and Mg2+. Leave around 0.5 mL of residual HBSS in the syringe so that tissue inadvertently sucked onto the tip of the catheter can be expelled.</li>
	<li>Transfer the lavaged medium, typically 8&#8211;8.5 mL in total, into a 14 mL polypropylene round bottom tube. Centrifuge the tube at 300 x g for 6.5 min at room temperature. 
	NOTE: The round bottom tube allows the supernatant to be fully decanted and reduces cell clumping.</li>
	<li>Discard the supernatant and resuspend the peritoneal cells (typically ~4 x 106 cells per mouse) in 200 &#181;L of modified RPMI 1640 HEPES medium. Dilute a sample of the cell suspension 1:20 and use a counting device, such as a Neubauer improved counting chamber, to count the cells. Next, dilute the cell suspension to a final concentration of 10 x 106 cells/mL and maintain the cells in a round bottom 2 mL polypropylene microcentrifuge tube at 37 &#176;C using a heated aluminum block (see NOTE in step 1.1.1).</li>
</ol>
3. Seeding Peritoneal Cells into Chemotaxis Slides
<ol>
	<li>After pipetting the cell suspension up and down 5x with the pipette volume set at 100 &#181;L (or at half the suspension volume) to reduce clumping, gently deposit 10 &#181;L of the cell suspension into port 3 of a chemotaxis chamber (Figure 2C). Place the pipette tip into port 2 and slowly draw the cell suspension into the connecting channel (Figure 2C). As soon as the cell suspension has been introduced, remove the plugs at ports 1 and 4, which will help to arrest the flow of the cell suspension. Place caps on all four filling ports.</li>
	<li>Repeat step 3.1 for all chemotaxis chambers. Using a small inverted microscope and a 10x phase-contrast objective lens, inspect the chemotaxis slides for unwanted air bubbles.</li>
	<li>Place the chemotaxis slides seeded with peritoneal cells in a humidity chamber at 37 &#176;C for 2&#8211;3 h.</li>
</ol>
4. Filling the Reservoirs and Adding Chemoattractant
<ol>
	<li>Inspect the observation area (channel connecting the two 40 &#181;L reservoirs) using an inverted microscope. 
	NOTE: At this stage, the cell density will be higher than after filling the reservoirs, because weakly adherent cells, predominantly CD19+ cells (B1 cells), will be washed out of the observation area during the filling procedure (Figure 2C&#8211;E).</li>
	<li>Place plugs into filling ports 1 and 2 (Figure 2D). Ensure that filling port 3 is filled to the top with medium and free of air bubbles. Use a sterile 27 G syringe needle to dislodge unwanted air bubbles, if required.</li>
	<li>Using a 10&#8211;100 &#181;L volume mechanical pipette, aspirate ~60 &#181;L of modified RPMI 1640 HEPES medium and place the pipette tip into filling port 3. Use the volume setting ring of the pipette to slowly and steadily inject medium into the reservoir such that medium reaches the top of filling port 4 after 1&#8211;2 min (Figure 2D).</li>
	<li>Fill the second reservoir. Move the plug from port 1 and slowly insert it into port 3 (Figure 2D). Next, aspirate ~50 &#181;L of modified RPMI 1640 HEPES medium and place the pipette tip into filling port 4. Use the volume setting ring of the 10&#8211;100 &#181;L volume pipette to slowly and steadily inject medium into the second reservoir such that medium reaches the top of filling port 1 after 1&#8211;2 min (Figure 2D).</li>
	<li>Place 495 &#181;L of modified RPMI 1640 HEPES medium into a (round bottom) 2 mL microcentrifuge tube and add 5 &#181;L of Patent Blue V (stock solution: 10 mg/mL in phosphate-buffered saline [PBS]), a blue dye used as a visual indicator of concentration gradient formation. Mix by brief vortexing. Add 5.4 &#181;L of recombinant mouse complement C5a (stock solution: 50 &#181;g/mL in PBS with 0.1% bovine serum albumin) and mix by brief vortexing.</li>
	<li>Deposit 15 &#181;L of blue, complement C5a-containing medium into filling port 1 (Figure 3A), after making sure that the shallow depression at the top of the port is medium free (otherwise the drop may spillover).</li>
	<li>Insert a 10&#8211;200 &#181;L pipette tip into filling port 4 and slowly and steadily rotate the volume setting ring of the 10&#8211;100 &#181;L volume pipette to draw the drop of blue, complement C5a-containing medium into the opposite reservoir (Figure 3B). Air will start to enter the short vertical column of filling port 1. Draw air in until the fluid-air interface is midway in the vertical column, and then slowly insert a plug into the port.</li>
	<li>Gently lift the pipette from port 4, using the other hand to ensure that the slide remains fixed in place. Finally, slowly plug port 4 (Figure 3B).</li>
	<li>Inspect the chemotaxis slide on an inverted microscope. 
	NOTE: The remaining adherent cells in the observation area should be predominantly macrophages. This can be confirmed using fluorescently tagged anti-F4/80 antibodies (F4/80 is a specific marker for mouse macrophages). B cells can be identified using fluorescently tagged anti-CD19 antibodies and F4/80-/CD19- cells can be detected using a blue fluorescent nucleic acid stain (Figure 4).</li>
</ol>
5. Imaging Macrophage Migration by Time-lapse, Phase-contrast Microscopy
<ol>
	<li>Place a chemotaxis slide on the stage of an inverted microscope fitted with a stage incubator. Maintain the temperature at 37 &#176;C.</li>
	<li>Image the 1 mm x 2 mm observation area using a 10x phase-contrast objective lens and focus on the macrophage lamellipodia: thin, sheet-like membrane protrusions. Capture images for 14 h at a rate of 1 frame every 2 min.</li>
</ol>
6. Analysis of Time-lapse Images
<ol>
	<li>Analyze the time-lapse, phase-contrast images using automated image analysis software or the Manual Tracking plugin, produced by Fabrice P. Cordeli&#232;res, for ImageJ. 
	NOTE: Automated tracking programs can be used to analyze cells imaged by either time-lapse, phase-contrast, or fluorescence microscopy. For example, the Java-based software iTrack4U produced by Cordeli&#232;res et al.27 can be used for automated cell tracking and analysis using time-lapse, phase-contrast, or fluorescence images as input. Manual tracking is more time consuming, but the tracks generated by the ImageJ plugin Manual Tracking can be directly imported and automatically analyzed by the ImageJ plugin Chemotaxis and Migration Tool28,29.</li>
</ol>

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G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+production+of+macrophages+or+neutrophils+ex+vivo+using+conditional+Hoxb8.">Quantitative production of macrophages or neutrophils ex vivo using conditional Hoxb8.</a> Nature Methods. 3, 287-293 (2006).</li><li>Gran, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging,+myeloid+precursor+immortalization,+and+genome+editing+for+defining+mechanisms+of+leukocyte+recruitment+in+vivo.">Imaging, myeloid precursor immortalization, and genome editing for defining mechanisms of leukocyte recruitment in vivo.</a> Theranostics. 8, 2407-2423 (2018).</li><li>Magidson, V., Khodjakov, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circumventing+photodamage+in+live-cell+microscopy.">Circumventing photodamage in live-cell microscopy.</a> Methods in Cell Biology. 114, 545-560 (2013).</li><li>Icha, J., Weber, M., Waters, J. C., Norden, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phototoxicity+in+live+fluorescence+microscopy,+and+how+to+avoid+it.">Phototoxicity in live fluorescence microscopy, and how to avoid it.</a> BioEssays : News and Reviews in Molecular, Cellular and Developmental Biology. 39 (8), 1700003 (2017).</li><li>Isfort, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+imaging+reveals+that+P2Y2+and+P2Y12+receptor+agonists+are+not+chemoattractants+and+macrophage+chemotaxis+to+complement+C5a+is+phosphatidylinositol+3-kinase+(PI3K)-+and+p38+mitogen-activated+protein+kinase+(MAPK)-independent.">Real-time imaging reveals that P2Y2 and P2Y12 receptor agonists are not chemoattractants and macrophage chemotaxis to complement C5a is phosphatidylinositol 3-kinase (PI3K)- and p38 mitogen-activated protein kinase (MAPK)-independent.</a> The Journal of Biological Chemistry. 286, 44776-44787 (2011).</li></ol>

The authors have nothing to disclose.

Intravital imaging dates back to the 19th century and provides a means to study the behavior of living immune cells in their natural environment. However, even with today&#8217;s advanced microscopy and genetic techniques it is difficult to study the response of cells to specific chemoattractants in vivo. To circumvent this problem, Boyden18 developed Transwell assays in the 1960s, but these end-point assays did not provide visualization of how cells actually migrated towards chemoattractants, making it difficult to distinguish chemokinesis, stimulated random migration by a chemical cue32, and chemotaxis, migration towards higher concentrations of chemical stimuli from each other33. This problem was solved by designing various open chambers with a bridge, typically 1 mm wide, situated between two reservoirs and accessible by an objective lens21,22,23. Applying an inverted cover slip, seeded with adherent cells, closes the chambers and chemoattractant added to one of the reservoirs diffuses across the bridge to the opposing reservoir, creating a concentration gradient. Here we describe a chemotaxis assay using the same principle but using a closed chamber featuring four filling ports. Using this system and time-lapse, phase-contrast microscopy, we developed an assay to image mouse resident peritoneal macrophages migrating in a chemotactic complement C5a gradient31,34,35,36. This assay, combined with knockout mouse models, proved instrumental in the investigation of the roles of various Rho GTPases and motor proteins in macrophage morphology, motility, and chemotaxis31,34,35,36,37. We have also used this approach to image human peripheral blood monocytes migrating on a 2D surface or in a 3D collagen type I matrix38. Moreover, the assay is suitable for mouse bone marrow-derived macrophages or macrophages derived from conditionally immortalized myeloid precursor cells39,40. We have previously used polytetrafluoroethylene (PTFE) bags with luer adapters to culture bone marrow cells and obtain macrophages34. The advantage of PTFE bags is that the cells can be readily resuspended and ready for use after placing the bag on ice for 20&#8211;30 min. Note that we prefill the chemotaxis slide observation area before introducing the cells. This approach has the advantage that unwanted air bubbles can be subsequently flushed out (with variable success) and the presoaked observation area enables the slow introduction of a cell suspension by pipetting. Prefilling, though, increases the likelihood that the medium will partially flow into one or both of the flanking reservoirs, which will promote the seeding of cells beyond the observation area. Alternatively, the cell suspension can be directly pipetted into a dry observation area, but unwanted air bubbles cannot be subsequently expelled.
The peritoneal cavity of the mouse contains two main populations of cells: F4/80+ macrophages and (smaller) CD19+ B cells, at a ratio of about 1:2 (Figure 4A). These two cell populations account for over 95% of peritoneal cavity cells, whereas the remaining F4/80-/CD19- cells can usually be identified as CD11c+ cells (dendritic cells) or CD3+ cells (T cells). Weakly adherent B cells are washed out of the observation area during the filling of the reservoirs with the medium (Figure 2). After adding chemoattractant to one of the two reservoirs, time-lapse, phase-contrast microscopy can be used to image the remaining cells (macrophages) migrating in an evolving chemoattractant gradient. The formation of the complement C5a gradient in the observation area, via diffusion from one reservoir to the other, can be simulated using a fluorescent dye with similar molecular weight. A good substitute for recombinant mouse complement C5a (predicted molecular weight, 9.0 kDa) is fluorescently labeled dextran (10 kDa)31. Using confocal microscopy, the fluorescence gradient in the narrow channel (observation area) connecting the two reservoirs of the chemotaxis slide can be measured at fixed intervals and concentration profiles at the different time points can be plotted24,31. We routinely add a nonfluorescent, blue dye (Patent Blue V) to the chemoattractant medium to provide a convenient visual indicator of diffusion and gradient formation. Within 1 h of introducing 15 &#181;L of blue, chemoattractant-containing medium into a reservoir, the reservoir appears uniformly blue and, according to Fick&#8217;s laws of diffusion, a gradient will form across the narrow observation area connecting the reservoirs (Figure 3B). Several days are required for the solute (blue dye or chemoattractant) to become uniformly distributed.
Fluorescence microscopy can be substituted for phase-contrast microscopy, which offers advantages for automated cell tracking, because fluorescently labeled cells can be readily distinguished from the background. Another advantage is that specific populations of immune cells can be selectively tracked after labeling surface markers with fluorescent antibodies. We used this approach to image human peripheral blood CD14+ cells (monocytes) migrating in a chemotactic fMLP (N-formylmethionine-leucyl-phenylalanine) gradient38. Similarly, fluorescent anti-F4/80 antibodies could be used to image mouse macrophages migrating in a chemotactic complement C5a gradient. Phototoxicity is a potential disadvantage of using fluorescence imaging41. This can be reduced by various means42, including using fluorophores excited with longer wavelengths and adding antioxidants to the medium. Alternatively, labeled cells could initially be identified by fluorescence microscopy and subsequently imaged by time-lapse, phase-contrast microscopy. However, in practice, cells moving at moderately low velocities, such as ~1 &#181;m/min (macrophages) or ~4 &#181;m/min (monocytes), can be intermittently imaged by fluorescence microscopy at intervals of minutes, which is well tolerated38. We previously used fluorescence microscopy and the chemotaxis slide described here for 3D chemotaxis assays38,43. In this case, both reservoirs were prefilled with medium and 15 &#181;L chemoattractant-containing medium was drawn into one of the reservoirs immediately before slowly pipetting fluorescently labeled cells suspended in medium containing collagen type I into the observation area. The difficult part of this procedure is the handling of collagen type I, which is concentrated in acidic solution. The pH of the collagen solution needs to be neutralized by addition of alkaline solution before mixing the ice-cold collagen solution with the cell suspension. Transfer of the collagen-cell mixture to an incubator at 37 &#176;C will initiate collagen polymerization. During incubation, the slide should be slowly rotated around its long axis so that the cells remain evenly distributed in the X-, Y- and Z-axis directions while the collagen polymerizes into a gel. A related closed chemotaxis slide suitable for 3D chemotaxis assays, with six plugs instead of four plugs, has recently been described29. This system allows the collagen-cell mixture to be introduced into the observation area before independently filling each of the flanking reservoirs, because each reservoir has two filling ports, rather than a single port.
In summary, we describe a real-time chemotaxis assay that allows the visualization of cells navigating in a chemotactic gradient over a period of 6 or more hours. Herein we focus on macrophages, which play major roles in inflammatory diseases but have been underrepresented in real-time chemotaxis assays compared to faster moving cells like neutrophils and Dictyostelium amoebae.

Immune cells typically migrate singly on a 2D surface in an amoeboid fashion1,2, which involves repeated cycles of protrusion of the front, integrin-mediated cell adhesion, and retraction of the rear. A prerequisite step is cell polarization, in which cells form front and rear ends3. Chemotaxis starts with the detection of chemoattractants by G protein-coupled receptors and a complex signaling network mediated by membrane-anchored heterotrimeric G proteins and small monomeric G proteins, as well as phospholipid-bound guanine nucleotide exchange factors (GEFs)4,5. Activation of Rho GTPases of the Cdc42 and Rac subfamilies induce protrusions at the front6 and members of the Rho subfamily, especially RhoA, activate contraction of the rear5,7. In a three-dimensional (3D) environment, integrins are largely redundant for leukocyte migration and RhoA becomes more important for squeezing cells through narrow passages8, whereas Cdc42- or Rac-induced Arp2/3 activation remains important for chemotactic steering9,10.
Immune cells may face different chemoattractants, especially in the settings of tissue injury, pathogen invasion, and inflammation. The endogenous chemoattractants expressed on phagocytes complement C3a and C5a, are rapidly generated by activation of the complement cascade, and are recognized by complement C3a and C5a receptors. Similarly, necrotic cells recruit phagocytes via formyl peptide receptors, which recognize mitochondria-derived as well as bacteria-derived formyl peptides11. Immune cells also express G protein-coupled receptors for chemokines, a large family of chemoattractant peptides involved in the regulation of immune cell trafficking during both homeostasis and inflammation. Chemokines are classified into four groups depending on the spacing of the first two cysteine (C) residues: C, CC, CXC, and CX3C cytokines, where X is an amino acid. Thus, in vivo immune cells need to appropriately respond to highly complex spatial and temporal signals, making the study of chemotaxis a daunting task. Below we provide a brief history of chemotaxis, which began with intravital imaging approaches.
The study of leukocyte chemotaxis dates back to 188812, when the ophthalmologist Theodor Karl Gustav Leber clearly described the directed migration of leukocytes to, and the accumulation at, sites of inflammation in a model of mycotic (fungal) keratitis. Leber stressed that the attraction of excess leukocytes by pathogen-derived substances is important for elimination of harmful microorganisms via phagocytosis, which had been described by Metchnikoff (also known as Metschnikoff) earlier in the same decade13. In vivo experiments were also performed in the 1920s by Clark and Clark14,15, who took advantage of the transparency of tadpoles and showed that sterile inflammation induced by croton oil14 or other irritants15 caused leukocytes to adhere to blood vessels, followed by diapedesis (transendothelial migration) and rapid migration through the tissue spaces towards the irritant. In vitro experiments using the microcinematography method developed by Jean Comandon16 showed that leukocytes migrated towards a particulate chemoattractant source such as bacteria17. At that time, the molecular identities of chemotactic factors were unknown. In the 1960s, Stephen Boyden18 recognized that techniques to study the chemotactic activity of soluble substances were lacking. He devised a chamber, subsequently known as the Boyden chamber, with two compartments separated by a filter paper membrane. A cell suspension is added to the upper compartment and the test substance is added to either both compartments or only to the lower compartment. After an incubation period, the filter membrane is removed, and the cells are fixed and stained. By comparing how many cells migrate across the filter membrane towards the lower well with the test substance in both compartments, in neither compartment, or only in the lower compartment, chemotactic activity can be determined. Transwell assays are still popular today and have been modified in various ways, including the use of different polycarbonate membranes with defined pore sizes and densities19,20. A major drawback of Transwell assays is that it is impractical to directly visualize cells migrating and the migration path across the membrane typically does not exceed the diameter of an immune cell.
Sally H. Zigmond developed a chemotaxis chamber21 that enabled visualization of both gradient formation and cell morphology using fluorescent dyes. The chamber consists of a plexiglass (acrylic) slide with two parallel linear wells, each with a volume of ~100 &#181;L, separated by a 1 mm wide bridge 3&#8211;10 &#181;m below the upper plane of the slide. A coverslip seeded with cells is inverted and placed onto the slide such that it spans the two wells. After addition of a chemoattractant to one of the wells, a steep chemoattractant gradient forms across the bridge, typically within 30&#8211;90 min. Human polymorphonuclear leukocytes (granulocytes) in the Zigmond chamber are observed orienting themselves towards the chemoattractant. Variations of the Zigmond chamber have been reported, including the Dunn22 and Insall23 chambers, both of which use a coverslip seeded with cells placed across two wells separated by a 1 mm wide bridge. The Dunn chamber consists of concentric wells separated by a circular bridge, whereas the Insall chamber is more closely related to the Zigmond chamber, but provides bridges of two different widths, 0.5 mm and 1 mm. A novel chemotaxis chamber, termed &#181;-Slide Chemotaxis and manufactured by plastic injection molding, was described by Zemgel et al.24. The chemotaxis chamber consists of two 40 &#181;L reservoirs separated by a 1 mm wide channel with a length of 2 mm and a height of 70 &#181;m. The bottom of the chamber is formed by a gas permeable, thin plastic sheet with the same thickness and optical properties of a No. 1.5 glass coverslip24. Here we describe a chemotaxis assay using the &#181;-Slide Chemotaxis chamber to visualize the migration of mouse resident peritoneal macrophages for up to 14 h in a chemotactic (complement C5a) gradient.

<table>
	<tbody>
		<tr>
			<td>&#181;-Slide (anodized aluminium) rack</td>
			<td>Ibidi, Martinsried, Germany</td>
			<td>80003</td>
			<td>Autoclavable stackable rack for channel slides</td>
		</tr>
		<tr>
			<td>&#181;-Slide Chemotaxis 2D (chemotaxis slide)</td>
			<td>Ibidi, Martinsried, Germany</td>
			<td>80306</td>
			<td>Slide containing chemotaxis chambers (tissue culture treated)</td>
		</tr>
		<tr>
			<td>100x penicillin/streptomycin</td>
			<td>Thermo Fisher Scientific</td>
			<td>15140122</td>
			<td>Used as supplement for RPMI 1640 media</td>
		</tr>
		<tr>
			<td>10-100 &#181;L pipette with volume control ring</td>
			<td>Eppendorf</td>
			<td>3123000047</td>
			<td>Eppendorf Research plus pipette</td>
		</tr>
		<tr>
			<td>10-200 &#181;L pipette tips</td>
			<td>Greiner Bio-One International</td>
			<td>739261</td>
			<td>Pipette tips with beveled tips (96 pieces per rack: sterile)</td>
		</tr>
		<tr>
			<td>14 ml polypropylene round bottom tubes</td>
			<td>BD Falcon</td>
			<td>352059</td>
			<td>Used to collect peritoneal cells</td>
		</tr>
		<tr>
			<td>14-bit Hamamatsu C9100-50 Electron Multiplying-Charged Couple Device (EM-CCD) peltier-cooled camera</td>
			<td>Hamamatsu Photonics Inc., Japan</td>
			<td></td>
			<td>EM-CCD camera of the spinning disk confocal microscope system</td>
		</tr>
		<tr>
			<td>2-20 &#181;L pipette with volume control ring</td>
			<td>Eppendorf</td>
			<td>3123000039</td>
			<td>Eppendorf Research plus pipette</td>
		</tr>
		<tr>
			<td>24-G plastic catheter</td>
			<td>B Braun Mesungen AG, Germany</td>
			<td>4254503-01</td>
			<td>Used for peritoneal lavage</td>
		</tr>
		<tr>
			<td>405 nm solid state laser, 50 mW</td>
			<td>Perkin Elmer, Rodgau, Germany</td>
			<td></td>
			<td>Laser (405 nm) source of spinning disk confocal microscope system</td>
		</tr>
		<tr>
			<td>488 nm solid state laser, 50 mW</td>
			<td>Perkin Elmer, Rodgau, Germany</td>
			<td></td>
			<td>Laser (488 nm) source of spinning disk confocal microscope system</td>
		</tr>
		<tr>
			<td>561 nm solid state laser, 50 mW</td>
			<td>Perkin Elmer, Rodgau, Germany</td>
			<td></td>
			<td>Laser (561 nm) source of spinning disk confocal microscope system</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488-conjugated rat (IgG2a) monoclonal (clone BM8) anti-mouse F4/80 antibody</td>
			<td>Thermo Fisher Scientific</td>
			<td>MF48020</td>
			<td>Mouse macrophage marker and plasma membrane label</td>
		</tr>
		<tr>
			<td>Alexa Fluor 594-conjugated rat (IgG2a) monoclonal (clone 6D5) anti-mouse CD19 antibody</td>
			<td>BioLegend</td>
			<td>115552</td>
			<td>Mouse B cell marker</td>
		</tr>
		<tr>
			<td>C-Chip disposable (improved Neubauer) hemocytometer</td>
			<td>NanoEnTek (distributed by VWR International)</td>
			<td>631-1098</td>
			<td>Used to count cells</td>
		</tr>
		<tr>
			<td>CSU-X1 spinning disk scanner</td>
			<td>Yokogawa Electric Corporation, Japan</td>
			<td></td>
			<td>Nipkow spinning disk unit</td>
		</tr>
		<tr>
			<td>Hank&#8217;s buffered salt solution without Ca2+ and Mg2+</td>
			<td>Thermo Fisher Scientific</td>
			<td>14170120</td>
			<td>Used for peritoneal lavage</td>
		</tr>
		<tr>
			<td>Heat-inactivated fetal bovine serum</td>
			<td>Thermo Fisher Scientific</td>
			<td>10082139</td>
			<td>Used as supplement for RPMI 1640 media</td>
		</tr>
		<tr>
			<td>Hoechst 34580</td>
			<td>Thermo Fisher Scientific</td>
			<td>H21486</td>
			<td>Cell permeable, blue fluorescent nucleic acid stain</td>
		</tr>
		<tr>
			<td>ImageJ (image processing and analysis in Java)</td>
			<td>National Institutes of Health (NIH)</td>
			<td></td>
			<td>Image analysis software</td>
		</tr>
		<tr>
			<td>Lipopolysaccharides from Escherichia coliO111:B4</td>
			<td>Sigma-Aldrich</td>
			<td>L4391-1MG</td>
			<td>Toll-like receptor 4 ligand</td>
		</tr>
		<tr>
			<td>Nikon Eclipse Ti inverse microscope</td>
			<td>Nikon, Japan</td>
			<td></td>
			<td>Inverted microscope</td>
		</tr>
		<tr>
			<td>Patent Blue V, sodium salt</td>
			<td>Sigma-Aldrich</td>
			<td>21605-10G</td>
			<td>Blue-colored dye used as visual indicator of gradient formation</td>
		</tr>
		<tr>
			<td>Recombinant mouse complement C5a protein</td>
			<td>R&#38;D Systems</td>
			<td>2150-C5-025</td>
			<td>Chemoattractant for mouse macrophages</td>
		</tr>
		<tr>
			<td>RPMI 1640 medium containing 20 mM Hepes</td>
			<td>Sigma-Aldrich</td>
			<td>R7388</td>
			<td>Basis medium for assays</td>
		</tr>
		<tr>
			<td>UltraVIEW Vox 3D live cell imaging system + Volocity software</td>
			<td>Perkin Elmer, Rodgau, Germany</td>
			<td></td>
			<td>Spinning disk confocal microscope system</td>
		</tr>
		<tr>
			<td>Zeiss LSM 510 + Axiovision software</td>
			<td>Carl Zeiss Microscopy, Oberkochen, Germany</td>
			<td></td>
			<td>Confocal laser scanning microscope (LSM) adapted for phase-contrast microscopy</td>
		</tr>
	</tbody>
</table>

time-lapse imaging of mouse macrophage chemotaxis

Chemotaxis is receptor-mediated guidance of cells along a chemical gradient, whereas chemokinesis is the stimulation of random cell motility by a chemical. Chemokinesis and chemotaxis are fundamental for the mobilization and deployment of immune cells. For example, chemokines (chemotactic cytokines) can rapidly recruit circulating neutrophils and monocytes to extravascular sites of inflammation. Chemoattractant receptors belong to the large family of G protein-coupled receptors. How chemoattractant (i.e., ligand) gradients direct cell migration via G protein-coupled receptor signaling is not yet fully understood. In the field of immunology, neutrophils are popular model cells for studying chemotaxis in vitro. Here we describe a real-time two-dimensional (2D) chemotaxis assay tailored for mouse resident macrophages, which have traditionally been more difficult to study. Macrophages move at a slow pace of ~1 &#181;m/min on a 2D surface and are less well suited for point-source migration assays (e.g., migration towards the tip of a micropipette filled with chemoattractant) than neutrophils or Dictyostelium discoideum, which move an order of magnitude faster. Widely used Transwell assays are useful for studying the chemotactic activity of different substances, but do not provide information on cell morphology, velocity, or chemotactic navigation. Here we describe a time-lapse microscopy-based macrophage chemotaxis assay that allows quantification of cell velocity and chemotactic efficiency and provides a platform to delineate the transducers, signal pathways, and effectors of chemotaxis.

A schematic diagram of the chemotaxis slide used for time-lapse video microscopy of mouse peritoneal macrophages migrating in a chemotactic gradient is shown in Figure 2A. The slide contains three chemotaxis chambers, each of which has four filling ports. Ports can be individually closed using the plugs shown above the slide. Alternatively, a non-sealing cap can be placed over an unplugged port to maintain sterility. After plugging ports 1 and 4, the observation area (1 mm wide x 2 mm long x 70 &#181;m high channel connecting the two reservoirs) between ports 2 and 3 can be prefilled with medium by placing a 15 &#181;L drop into port 3 and aspirating with a 2&#8211;20 &#181;L volume pipette at port 2 (Figure 2B). A suspension of mouse resident peritoneal cells (10 x 106 cells/mL) was seeded into the observation area by placing a 10 &#181;L drop of suspension into port 3 and slowly aspirating at port 2 (Figure 2C). A typical image of cells seeded in the observation area taken by phase-contrast microscopy using a 10x objective lens is shown in Figure 2C. After incubating for 2&#8211;3 h, the chemotaxis slide was slowly filled with medium (Figure 2D). After plugging ports 1 and 2, medium was slowly injected via port 3 until it emerged from port 4. Next, the plug was switched from port 1 to port 3, and then the second reservoir was filled by slowly injecting medium via port 4 until it emerged at port 1. At this stage, cells in the observation area were reinspected using an inverted microscope (Figure 2E). By comparing images shortly before (Figure 2C) and after (Figure 2E) filling of the reservoirs, up to two-thirds of the cells had been washed out of the observation area. Generally, weakly adherent CD19+ cells (B1 cells) were washed out and the remaining cells were predominantly F4/80+ cells (macrophages). This was demonstrated by fluorescence microscopy after labeling each cell type with fluorescently labeled specific antibodies (Figure 4). In Figure 4A, freshly isolated mouse resident peritoneal cells were labeled with green fluorescent fluorophore-conjugated anti-F4/80 antibodies and red fluorescent fluorophore-conjugated anti-CD19 antibodies, and the nuclei of cells were labeled with a blue fluorescent nucleic acid stain. F4/80 is a specific marker for mouse macrophages30, whereas CD19 is a B cell marker. Figure 4B shows F4/80+ cells imaged by spinning disk confocal microscopy in the observation area of a chemotaxis chamber. The cells were labeled after an overnight chemotaxis assay recorded by time-lapse, phase-contrast microscopy.
Complement C5a (chemoattractant) was introduced to one of the two reservoirs by placing a 15 &#181;L drop of medium containing 0.54 &#181;g/mL (recombinant mouse) complement C5a and 10 &#181;g/mL Patent Blue V into filling port 1 (Figure 3A) after plugging ports 2 and 3. The chemoattractant medium was slowly drawn into the reservoir by slow aspiration with a pipette via port 4. Figure 3B shows the diffusion of the blue dye after drawing the 15 &#181;L drop into a reservoir. Patent Blue V was used as an indirect visual indicator of chemoattractant diffusion. Complement C5a molecules are considerably larger than those of Patent Blue V (9.0 kDa versus 0.57 kDa) and diffuse more slowly. After diffusion of complement C5a in the reservoir, its concentration was ~0.2 &#181;g/mL (15 &#181;L/40 &#181;L [reservoir volume] x 0.54 &#181;g/mL = 0.2 &#181;g/mL), equivalent to ~22.5 nM. A modestly steep gradient formed across the observation area after 3 h and continued to increase, reaching a maximum at around 12 h31. Figure 3C shows the migration tracks of macrophages migrating in a complement C5a gradient, between 6&#8211;12 h after adding the chemoattractant. Cell velocity and chemotactic efficiency, indexed as y-FMI (y-forward migration index; range: -1 to +1) and x-FMI, of individual macrophages was calculated from the migration plots (Figure 3D). Figure 3D also shows a migration plot produced after normalizing the start point of each migration track to X = 0 and Y = 0 below the box plots. The inset in the migration plot shows how the y-FMI was calculated for each migration track.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60750/60750fig01.jpg" /> 
Figure 1: The workflow of the chemotaxis assay. <a href="https://www.jove.com/files/ftp_upload/60750/60750fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60750/60750fig02.jpg" /> 
Figure 2: Handling of chemotaxis slides.&#160;(A) A 3D view of a chemotaxis slide with four plugs and four caps. The slide contains three chemotaxis chambers, each of which consists of two 40 &#181;L reservoirs connected by a 1 mm x 2 mm channel, which is 70 &#181;m high, termed the observation area. (B) The connecting channel extends at both ends to filling ports 2 and 3. After inserting plugs into filling ports 1 and 4, the observation area was prefilled with medium (red) by applying a drop of medium to port 3 and aspirating at port 2 with a 10&#8211;200 &#181;L pipette tip. Subsequently, caps were applied to ports 2 and 3 before incubating the slide at 37 &#176;C and preparing the cell suspension. (C) The observation area, where the chemoattractant gradient formed, was seeded with macrophages by applying a 10 &#181;L drop of mouse resident peritoneal cells at port 3 and slowly aspirating at port 2. The slide was then incubated in a humidity chamber at 37 &#176;C for 2&#8211;3 h. The phase-contrast image shown on the right, obtained via a 10x objective lens, shows peritoneal cells after seeding and incubation at 37 &#176;C for 2 h. Scale bar = 500 &#181;m. (D) Chemotaxis chambers were filled with medium by plugging ports 1 and 2 and then slowly injecting medium via port 3 until it emerged at port 4. Slow and steady filling can be achieved by turning the volume setting ring of a 20&#8211;100 &#181;L volume pipette. After filling the first reservoir, the second reservoir can be filled by plugging ports 2 and 3 and then slowly injecting medium at port 4 until it emerges at port 1. (E) Phase-contrast image of the same observation area shown above (C) after filling the two reservoirs. Scale bar = 500 &#181;m. Graphic elements provided by Elias Horn. <a href="https://www.jove.com/files/ftp_upload/60750/60750fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60750/60750fig03.jpg" /> 
Figure 3: Chemotaxis assay.&#160;(A) Chemoattractant was introduced to one of the two reservoirs of a chemotaxis chamber by applying a 15 &#181;L drop of medium containing 0.54 &#181;g/mL complement C5a and 10 &#181;g/mL Patent Blue V to filling port 1, followed by slow aspiration at port 4. (B) Initially after being drawn into the reservoir the blue, chemoattractant-containing medium had a roughly inverted drop shape, and then slowly diffused throughout the reservoir. (C) Migration tracks of macrophages migrating in a chemoattractant (complement C5a) gradient between 6&#8211;12 h after introducing chemoattractant to one of the reservoirs. The direction of the gradient is indicated on the right. The end of each migration track is indicated by a filled circle. (D) Blox plots of velocity, x-FMI (x-forward migration index) and y-FMI (y-forward migration index), an index of chemotactic efficiency that ranges from -1 to +1. Data were obtained by analysis of 25 macrophage migration tracks. Macrophages in the lower half of the observation area and showing displacement of at least one cell width over 6 h were randomly selected for analysis. Below is a plot of migration tracks after normalizing the start point to X = 0 and Y = 0. The chemotaxis index (y-FMI) was calculated by dividing the net displacement along the Y-axis (d) by the accumulated length (l) of the migration path, as schematically shown. Graphic elements in panels A and B provided by Elias Horn. <a href="https://www.jove.com/files/ftp_upload/60750/60750fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60750/60750fig04.jpg" /> 
Figure 4: Fluorescent images of living mouse resident peritoneal cells obtained by spinning disk confocal microscopy.&#160;(A) Extended focus image (brightest point merge of all Z-planes) of freshly isolated mouse peritoneal cells labeled with green fluorescent anti-F4/80 (macrophage marker) antibodies, red fluorescent anti-CD19 (B cell marker) antibodies, and a blue fluorescent nucleic acid stain. Scale bar = 10 &#181;m. (B) Snapshot (single Z-plane) of F4/80+ cells (macrophages) in the observation area of a chemotaxis chamber taken after an overnight chemotaxis assay. Cells were labeled with green fluorescent anti-F4/80 antibodies and a blue fluorescent nucleic acid stain. The complement C5a and Patent Blue V gradients were washed out by the cell labeling procedure, which explains why the upper reservoir in the schematic diagram of the chemotaxis chamber is not blue. Scale bar = 10 &#181;m. Graphic element provided by Elias Horn. <a href="https://www.jove.com/files/ftp_upload/60750/60750fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Time-lapse Imaging of Mouse Macrophage Chemotaxis at JoVE.com

Time-lapse Imaging of Mouse Macrophage Chemotaxis

Here we describe methods using time-lapse, phase-contrast microscopy to image mouse resident peritoneal macrophages in a chemotactic complement C5a gradient. The protocols can be extended to other immune cells.

Chemotaxis is receptor-mediated guidance of cells along a chemical gradient, whereas chemokinesis is the stimulation of random cell motility by a ...

time-lapse-imaging-of-mouse-macrophage-chemotaxis

Universidad San Sebastian

Research

JoVE Journal

Immunology and Infection

11.5K Views.  Institut für Molekulare Zellbiologie. Here we describe methods using time-lapse, phase-contrast microscopy to image mouse resident peritoneal macrophages in a chemotactic complement C5a gradient. The protocols can be extended to other immune cells.

Video: Time-lapse Imaging of Mouse Macrophage Chemotaxis

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not representative of the behavior, status or phenotype of single cells. Due to this new insight the number of single cell studies rises continuously (for recent reviews see 1,2,3). However, many of the single cell techniques applied do not allow monitoring the development and behavior of one specific single cell in time (e.g. flow cytometry or standard microscopy).
Here, we provide a detailed description of a microscopy method used in several recent studies 4, 5, 6, 7, which allows following and recording (fluorescence of) individual bacterial cells of Bacillus subtilis and Streptococcus pneumoniae through growth and division for many generations. The resulting movies can be used to construct phylogenetic lineage trees by tracing back the history of a single cell within a population that originated from one common ancestor. This time-lapse fluorescence microscopy method cannot only be used to investigate growth, division and differentiation of individual cells, but also to analyze the effect of cell history and ancestry on specific cellular behavior. Furthermore, time-lapse microscopy is ideally suited to examine gene expression dynamics and protein localization during the bacterial cell cycle. The method explains how to prepare the bacterial cells and construct the microscope slide to enable the outgrowth of single cells into a microcolony. In short, single cells are spotted on a semi-solid surface consisting of growth medium supplemented with agarose on which they grow and divide under a fluorescence microscope within a temperature controlled environmental chamber. Images are captured at specific intervals and are later analyzed using the open source software ImageJ.

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

This protocol provides a step-by-step procedure to monitor single cell behavior of different bacteria in time using automated fluorescence time-lapse microscopy. Furthermore, we provide guidelines how to analyze the microscopy images.

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not ...

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

Investigation of Macrophage Polarization Using Bone Marrow Derived Macrophages

The article describes a readily easy adaptive in vitro model to investigate macrophage polarization. In the presence of GM-CSF/M-CSF, hematopoietic stem/progenitor cells from the bone marrow are directed into monocytic differentiation, followed by M1 or M2 stimulation. The activation status can be tracked by changes in cell surface antigens, gene expression and cell signaling pathways.

The article describes a readily easy adaptive in vitro model to investigate macrophage polarization. In the presence of GM-CSF/M-CSF, hematopoietic stem/progenitor cells from the bone marrow are directed into monocytic differentiation, followed by M1 or M2 stimulation. The activation status can be tracked by changes in cell surface antigens, gene expression and cell signaling pathways.

The article describes a readily easy adaptive in vitro model to investigate macrophage polarization. In the presence of GM-CSF/M-CSF, hematopoietic ...

Studying Interactions of Staphylococcus aureus with Neutrophils by Flow Cytometry and Time Lapse Microscopy

We present methods to study the effect of phenol soluble modulins (PSMs) and other toxins produced and secreted by Staphylococcus aureus on neutrophils. To study the effects of the PSMs on neutrophils we isolate fresh neutrophils using density gradient centrifugation. These neutrophils are loaded with a dye that fluoresces upon calcium mobilization. The activation of neutrophils by PSMs initiates a rapid and transient increase in the free intracellular calcium concentration. In a flow cytometry experiment this rapid mobilization can be measured by monitoring the fluorescence of a pre-loaded dye that reacts to the increased concentration of free Ca2+. Using this method we can determine the PSM concentration necessary to activate the neutrophil, and measure the effects of specific and general inhibitors of the neutrophil activation.	
	To investigate the expression of the PSMs in the intracellular space, we have constructed reporter fusions of the promoter of the PSM&alpha; operon to GFP. When these reporter strains of S. aureus are phagocytosed by neutrophils, the induction of expression can be observed using fluorescence microscopy.

Studying Interactions of Staphylococcus aureus with Neutrophils by Flow Cytometry and Time Lapse Microscopy

We present methods to study the effect of PSMs and other toxins secreted by Staphylococcus aureus on neutrophils using flow cytometry and fluorescence microscopy.

We present methods to study the effect of phenol soluble modulins (PSMs)  and other toxins produced and secreted by Staphylococcus  aureus on neutrophils. ...

Bone Marrow-derived Macrophage Production

Macrophages are critical components of the innate and adaptive immune responses, and they are the first line of defense against foreign invaders because of their powerful microbicidal activities. Macrophages are widely distributed throughout the body and are present in the lymphoid organs, liver, lungs, gastrointestinal tract, central nervous system, bone, and skin. Because of their repartition, they participate in a wide range of physiological and pathological processes. Macrophages are highly versatile cells that are able to recognize microenvironmental alterations and to maintain tissue homeostasis. Numerous pathogens have evolved mechanisms to use macrophages as Trojan horses to survive, replicate in, and infect both humans and animals and to propagate throughout the body. The recent explosion of interest in evolutionary, genetic, and biochemical aspects of host-pathogen interactions has renewed scientific attention regarding macrophages. Here, we describe a procedure to isolate and cultivate macrophages from murine bone marrow that will provide large numbers of macrophages for studying host-pathogen interactions as well as other processes.

Macrophages have long been recognized as a critical component of the innate and adaptive immune responses. The recent explosion of knowledge pertaining to evolutionary, genetic, and biochemical aspects of the interaction between macrophages and microbes has renewed scientific attention to macrophages. This article describes a method to differentiate macrophages from mouse bone marrow.

Macrophages are critical components of the innate and adaptive immune responses, and they are the first line of defense against foreign invaders because ...

Fluorescence Time-lapse Imaging of the Complete S. venezuelae Life Cycle Using a Microfluidic Device

Live-cell imaging of biological processes at the single cell level has been instrumental to our current understanding of the subcellular organization of bacterial cells. However, the application of time-lapse microscopy to study the cell biological processes underpinning development in the sporulating filamentous bacteria Streptomyces has been hampered by technical difficulties. 
Here we present a protocol to overcome these limitations by growing the new model species, Streptomyces venezuelae, in a commercially available microfluidic device which is connected to an inverted fluorescence widefield microscope. Unlike the classical model species, Streptomyces coelicolor, S. venezuelae sporulates in liquid, allowing the application of microfluidic growth chambers to cultivate and microscopically monitor the cellular development and differentiation of S. venezuelae over long time periods. In addition to monitoring morphological changes, the spatio-temporal distribution of fluorescently labeled target proteins can also be visualized by time-lapse microscopy. Moreover, the microfluidic platform offers the experimental flexibility to exchange the culture medium, which is used in the detailed protocol to stimulate sporulation of S. venezuelae in the microfluidic chamber. Images of the entire S. venezuelae life cycle are acquired at specific intervals and processed in the open-source software Fiji to produce movies of the recorded time-series.

Fluorescence Time-lapse Imaging of the Complete S. venezuelae Life Cycle Using a Microfluidic Device

Streptomyces are characterized by a complex life cycle that has been experimentally challenging to study by cell biological means. Here we present a protocol to perform fluorescence time-lapse microscopy of the complete life cycle by growing Streptomyces venezuelae in a microfluidic device.

Live-cell imaging of biological processes at the single cell level has been instrumental to our current understanding of the subcellular organization of ...

Visualizing Macrophage Extracellular Traps Using Confocal Microscopy

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal microscopy methods to visualize macrophage extracellular traps (METs). These extracellular traps are defined by the presence of extracellular chromatin with co-expression of other components such as granule proteases, citrullinated histones, and peptidyl arginase deiminase (PAD). The expression of METs is generally measured after exposure to a stimulus and compared to un-stimulated samples. Samples are also included for background and isotype control. Cells are analyzed using well-defined image analysis software. Confocal microscopy may be used to define the presence of METs both in vitro and in vivo in lung tissue.

Macrophage extracellular traps are a newly described entity. This article will concentrate on confocal microscopy methods and how they are visualized in vitro and in vivo from lung samples.

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal ...

Fabricating Optical-quality Glass Surfaces to Study Macrophage Fusion

Visualizing the formation of multinucleated giant cells (MGCs) from living specimens has been challenging due to the fact that most live imaging techniques require propagation of light through glass, but on glass macrophage fusion is a rare event. This protocol presents the fabrication of several optical-quality glass surfaces where adsorption of compounds containing long-chain hydrocarbons transforms glass into a fusogenic surface. First, preparation of clean glass surfaces as starting material for surface modification is described. Second, a method is provided for the adsorption of compounds containing long-chain hydrocarbons to convert non-fusogenic glass into a fusogenic substrate. Third, this protocol describes fabrication of surface micropatterns that promote a high degree of spatiotemporal control over MGC formation. Finally, fabricating glass bottom dishes is described. Examples of use of this in vitro cell system as a model to study macrophage fusion and MGC formation are shown.

This protocol describes the fabrication of optical-quality glass surfaces adsorbed with compounds containing long-chain hydrocarbons that can be used to monitor macrophage fusion of living specimens and enables super-resolution microscopy of fixed specimens.

Visualizing the formation of multinucleated giant cells (MGCs) from living specimens has been challenging due to the fact that most live imaging ...

Time-lapse 3D Imaging of Phagocytosis by Mouse Macrophages

Phagocytosis plays a key role in host defense, as well as in tissue development and maintenance, and involves rapid, receptor-mediated rearrangements of the actin cytoskeleton to capture, envelop and engulf large particles. Although phagocytic receptors, downstream signaling pathways, and effectors, such as Rho GTPases, have been identified, the dynamic cytoskeletal remodeling of specific receptor-mediated phagocytic events remain unclear. Four decades ago, two distinct mechanisms of phagocytosis, exemplified by Fc&#947; receptor (Fc&#947;R)- and complement receptor (CR)-mediated phagocytosis, were identified using scanning electron microscopy. Binding of immunoglobulin G (IgG)-opsonized particles to Fc&#947;Rs triggers the protrusion of thin membrane extensions, which initially form a so-called phagocytic cup around the particle before it becomes completely enclosed and retracted into the cell. In contrast, complement opsonized particles appear to sink into the phagocyte following binding to complement receptors. These two modes of phagocytosis, phagocytic cup formation and sinking in, have become well established in the literature. However, the distinctions between the two modes have become blurred by reports that complement receptor-mediated phagocytosis may induce various membrane protrusions. With the availability of high resolution imaging techniques, phagocytosis assays are required that allow real-time 3D (three dimensional) visualization of how specific phagocytic receptors mediate the uptake of individual particles. More commonly used approaches for the study of phagocytosis, such as end-point assays, miss the opportunity to understand what is happening at the interface of particles and phagocytes. Here we describe phagocytic assays, using time-lapse spinning disk confocal microscopy, that allow 3D imaging of single phagocytic events. In addition, we describe assays to unambiguously image Fc&#947; receptor- or complement receptor-mediated phagocytosis.

Here we describe methods using spinning disk confocal microscopy to image single phagocytic events by mouse resident peritoneal macrophages. The protocols can be extended to other phagocytic cells.

Phagocytosis plays a key role in host defense, as well as in tissue development and maintenance, and involves rapid, receptor-mediated rearrangements of ...

In Vitro Differentiation of Mouse Granulocyte-macrophage-colony-stimulating Factor (GM-CSF)-producing T Helper (THGM) Cells

The granulocyte-macrophage-colony-stimulating factor (GM-CSF)-producing T helper (THGM) cell is a newly identified T helper cell subset that predominantly secretes GM-CSF without producing interferon (IFN)&#947; or interleukin (IL)-17 and is found to play an essential role in the autoimmune neuroinflammation. A method of isolation of naive CD4+ T cells from a single-cell suspension of splenocytes and THGM cell generation from naive CD4+ T cells would be a useful technique in the study of T cell-mediated immunity and autoimmune diseases. Here we describe a method that differentiates mouse naive CD4+ T cells into THGM cells promoted by IL-7. The outcome of the differentiation was assessed by the analysis of the cytokines expression using different techniques, including intracellular cytokine staining combined with flow cytometry, a quantitative real-time polymerase chain reaction (PCR), and enzyme-linked immunosorbent assays (ELISA). Using the THGM differentiation protocol as described here, about 55% of the cells expressed GM-CSF with a minimal expression of IFN&#945; or IL-17. The predominant expression of GM-CSF by THGM cells was further confirmed by the analysis of the expression of GM-CSF, IFN&#945;, and IL-17 at both mRNA and protein levels. Thus, this method can be used to differentiate naive CD4+ T cells to THGM cells in vitro, which will be useful in the study of THGM cell biology.

In Vitro Differentiation of Mouse Granulocyte-macrophage-colony-stimulating Factor GM-CSF-producing T Helper THGM Cells

Here, we present a protocol to differentiate murine granulocyte-macrophage-colony-stimulating-factor-producing T helper (THGM) cells from naive CD4+ T cells, including isolation of naive CD4+ T cells, differentiation of THGM, and analysis of differentiated THGM cells. This method can be applied to studies of the regulation and function of THGM cells.

The granulocyte-macrophage-colony-stimulating factor (GM-CSF)-producing T helper (THGM) cell is a newly identified T helper cell subset that predominantly ...