Name: high throughput fluorometric technique for assessment of macrophage phagocytosis and actin polymerization
Uploaded: 2014-11-27T15:00:00.000+00:00
Duration: 9 min 22 s
Description: Watch this Scientific Journal Video about High Throughput Fluorometric Technique for Assessment of Macrophage Phagocytosis and Actin Polymerization at JoVE.com

The authors would like to thank the following funding sources: RO1 DA 12104, RO1 DA 022935, RO1 DA031202, K05DA033881, P50 DA 011806, 1R01DA034582 (to S.R) and F31 DA026264-01A1, T32 DA07097 (to J.N.).

NOTE: This protocol can be used for multiple applications such as quantification of phagocytosis and actin polymerization as used in our previously published work12. The protocol lends itself to a variety of modification, and Table 1 lists cell types and particles successfully utilized in past studies. Standard use of this protocol for quantification of phagocytosis or actin polymerization is illustrated in Diagram 1.
<img alt="Diagram 1" fo:content-width="4in" src="/files/ftp_upload/52195/52195diagram1.jpg" width="600" />
Diagram 1: Diagram of fluorometric assay for quantification of phagocytosis and actin polymerization. After the cells are plated, treated, and allowed to adhere, particles labeled with green fluorescence (FITC) are added to the cells for phagocytosis. Reaction is stopped by trypan or antibiotic wash (to eliminate non-internalized particles) and fixation is performed with 4% paraformaldehyde. Cells are subsequently stained with red fluorescent actin label (rhodamine phalloidin) and blue fluorescent label (DAPI). Indexes of phagocytosis and actin polymerization are quantified as a ratio of relative fluorescence units of green/blue (FITC/DAPI), or red/blue (rhodamine/DAPI) fluorescence.
1. Plating and Treating Cells
NOTE: Before beginning, consider running the treatments with at least 4 technical replicates, and include the following controls in the plate layout: cells only (unstained, with DAPI alone or rhodamine alone) and particle only.
<ol>
	<li>Conduct all addition of reagents in steps 1-4 in a sterile cell culture hood in order to protect the samples from contamination.</li>
	<li>Plate macrophages (murine J774 cell line) in a 96-well plate at 1-5 x 104 cells in 50 &#956;l (per well) of supplemented preheated media. Ideal plates for this method are black 96-well plates with clear or black bottoms, depending on reader capabilities.
	<ol>
		<li>For supplemented media, use 10% heat inactivated FBS to avoid interference of complement cascade, and 1% penicillin/streptomycin (if not using live bacteria for phagocytosis). If using adherent cells allow 1-2 hr for adhesion, and for non-adherent cells, spin down the plate with cells for 10 min at 500 x g.</li>
	</ol>
	</li>
	<li>Add desired agonists/antagonists or vehicle control at 2x concentration (resuspended in PBS or more preferably, in supplemented media) in 50 &#956;l for a final volume of 100 &#956;l (1x) and incubate for a desired amount of time. Multichannel pipette and reagent reservoirs will facilitate fastest processing.</li>
</ol>
2. Opsonization of Fluorescent Particles
<img alt="Table 1" fo:content-width="5in" src="/files/ftp_upload/52195/52195table1.jpg" width="500" /> 
Table 1: Tested cell types and fluorescent particles in fluorometric analysis of phagocytosis and actin polymerization. Table 1 illustrates combinations of cell types (cell lines, and primary cells) that our group has used via the fluorometric method. Other than looking at phagocytosis and actin polymerization by wild type cells, we have also utilized some of these particles for examination of phagocytosis by transfected cells expressing a GFP (green fluorescent protein). Fluorescence of the cells transfected with plasmids containing fluorescent reporters can also be used as a cellular marker in addition to DAPI. Table legend: A - actin polymerization; P - phagocytosis; PT - phagocytosis by GFP labeled transfected cells; OPDex - opsonized dextran bead; HKOP - heat killed opsonized.
<ol>
	<li>Ensure that fluorescent particles and labels are protected from light at all times, during processing, washing, as well as during the incubation in order to prevent photobleaching
	<ol>
		<li>For dry particles, weigh out fluorescent heat killed opsonized (HKOP) bacteria (E2861: E. coli K-12 strain), using manufacturer&#8217;s specifications. Take care to make sure the mass accounts for 10:1 - 20:1 bacteria:cell ratio (or at least a 10:1 ratio - commonly used in quantification of phagocytosis)13,14. Number of particles can be provided by the manufacturer or determined via couting serial dilutions of 1 mg/ml stock solution.</li>
		<li>Resuspend in 50 &#956;l PBS and vortex for 1 minute at highest setting. Ensure proper vortexing (here and throughout the protocol) as the particles tend to aggregate which may affect efficacy of opsonization.</li>
		<li>For particles in suspension such as fluorescently labeled dextran beads (DEX beads), vortex the stock solution for 1 min, take 50 &#956;l of suspension or depending on the bead concentration in the volume determine mount of beads to facilitate 10:1 bead:cell ratio.</li>
	</ol>
	</li>
	<li>Add an equal volume of the opsonizing reagent to the particles and vigorously vortex for another minute.</li>
	<li>Incubate the particles with opsonizing reagent at 37 &#176;C for 1 hr.</li>
	<li>Following the incubation, centrifuge the particles for 5 min at 500 x g, and remove the supernant containing excess opsonizing reagent. Wash the particles by adding 100 &#956;l of PBS, vortex to resuspend the pellet, and centrifugate for 5 min at 500 x g.
	<ol>
		<li>Repeat these steps two times to assure proper opsonization and removal of unbound antibody.</li>
	</ol>
	</li>
	<li>Prepare the working solution of opsonized particles by resuspending them in 5 ml of media (sufficient for one 96-well plate) and store away from light until addition to the cells.</li>
</ol>
3. Phagocytosis
<ol>
	<li>Place the working solution of particles in a sterile reagent reservoir. From here, add 50 &#956;l of opsonized particles to the cells prepared in step 1 (for a final concentration of 50 ng/ml) and allow phagocytosis to occur at standard cell culture conditions. Illustrated below are two methods of phagocytosis which can be assessed using this method (Diagram 2).</li>
</ol>
<img alt="Diagram 2" fo:content-width="4in" src="/files/ftp_upload/52195/52195diagram2.jpg" width="600" /> 
Diagram 2: Comparison of continuous versus synchronize phagocytosis. Continuous phagocytosis indicates continuous internalization of particles over time as they slowly reach the cells on the bottom of the well. A contrasting synchronization step (centrifugation) forces the particles to sink to the bottom, enhancing the particle contact with the cell, and leading to immediate internalization by the cells. Synchronized phagocytosis is a quicker process which more rapidly internalizes particles due to the increased cell:particle contact.
<ol>
	<li>
	<ol>
		<li>For the continuous phagocytosis method (Diagram 2) that allow cells to continuously phagocytose opsonized bacteria as they slowly sink to the bottom and come in contact with cells, utilize longer time frames to allow for the bacteria to reach the bottom of the well and get in touch with the cells. If conducting a timecourse experiment, add bacteria at different time intervals and stop the entire plate of reactions at the same time to enhance the speed of processing.</li>
		<li>For the synchronized phagocytosis method (Diagram 2) that enhances particle: cell contact via centrifugation, spin down the entire plate containing cells and particles (5 min at 500 x g) as soon as the particles are added, to facilitate synchronization of all time points and to expedite the particle: cell interaction.
		<ol>
			<li>Measure the time course starting right after the completion of the centrifugation step. If conducting a time course experiment, stop each time point independently at different time intervals as described below.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
NOTE: The synchronized phagocytosis method is somewhat more laborious from the previous one, but provides higher phagocytic index in a shorter time.
4. Stopping Phagocytosis
<ol>
	<li>For adherent cells, remove the supernatant and add 50 &#956;l of diluted trypan blue with PBS (1:2 dilution), in order to extinguish fluorescence of non-internalized bacteria.</li>
	<li>For non-adherent cells, spin down the plate and remove supernatant. Add 50 &#956;l of diluted trypan blue. Ensure to centrifuge the plate at (5-10 min at 500 x g) between each wash in order to minimize the cell loss and enable adequate washing.</li>
	<li>Following trypan incubation, wash cells two times with PBS to remove any residual trypan (until the PBS removed from the well turns clear). Trypan has been shown to quench fluorescence15,16 of non-internalized fluorescently labeled, heat killed, bacterial particles, but not for live bacteria or OPDex beads. For live bacteria, include a wash with antibiotics such as penicillin, streptomycin, or gentamicin instead of the trypan wash to avoid confounding effects of fluorescence from non-internalized particles.</li>
	<li>Fix cells with 100 &#956;l of 4% paraformaldehyde, and incubate for 15 min at room temperature, then wash with PBS (at 100 &#956;l per well for each wash). At this step, save the cells at 4 &#176;C for up to a week, or proceed directly to staining and quantification. 
	NOTE: All the subsequent steps form this point foreword can be done outside of the cell culture hood.</li>
	<li>Remove all liquid and add 50 &#956;l of DAPI at 5 ng/ml reconstituted in PBS. Allow 5 min for staining, and then wash twice with PBS. Following the wash step, add 50 &#956;l of PBS to each well and record phagocytosis.</li>
</ol>
5. Actin Staining
NOTE: In addition to phagocytosis, fluorometric method will enable quantification of actin polymerization by measuring intensity of actin, which is reduced during inhibition of lamellipodia, pseudopodiae, and membrane ruffling as a result of treatment with an inhibitor (Figure 1A). This is an optional step if interested in quantification of actin polymerization in addition to phagocytosis. If actin polymerization is the final goal, using fluorescent particles for phagocytosis is optional.
<ol>
	<li>Following paraformaldehyde fixation (step 4.3), wash the plate twice with PBS at 100 &#956;l per well for each wash. Add 50 &#956;l of 0.1% Triton X-100 in PBS for permeabilization and incubate for&#160; -5 min at RT.</li>
	<li>Wash out 0.1% Triton X-100 2 times with PBS (as above), and block with 1% bovine serum albumin (BSA) for 20 min to avoid non-specific binding of the label.</li>
	<li>Following blocking, remove the BSA solution and immediately add rhodamine phalloidin. To prepare the working solution of rhodamine stain, use 100 &#956;l of methanoic solution (provided by the manufacturer) in 5 ml of PBS (sufficient for one plate). From the working solution add 50 &#956;l per well and incubate for 20 min at room temperature. 
	NOTE: Following the washing steps and DAPI staining as in step 4.4 cells will be ready for quantification.</li>
</ol>
6. Quantification
<ol>
	<li>To quantify the phagocytosis using a fluorescent plate reader, record green fluorescence using excitation filter at &#955; = 488 nm and emission filter at &#955; = 518 nm, and blue fluorescence using excitation filter at &#955; = 355 nm and emission filter at &#955; = 460 nm.</li>
	<li>To quantify the actin polymerization using a fluorescent plate reader, record blue fluorescence via &#955; = 355 nm and &#955; = 460 nm (as above), and red fluorescence via excitation filter at &#955; = 584 nm and emission filter &#955; = 620 nm. 
	NOTE: Orbital or central detection is acceptable for recording. Some cell types gather more to the edge of the well than the center, in which case orbital recording can be of advantage. In particular, average of three points around the orbit could also be used to obtain more accurate results.</li>
	<li>Divide the total RFU of green or red fluorescence by blue fluorescence readout (from DAPI), depending on the particle label or phagocytic vs. actin quantification (Diagram 1). Due to high plate-to-plate variance, use relative indexes (such as relative to t = 0 control), to best illustrate trends observed and to facilitate more reliable statistical analysis.</li>
</ol>

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The authors have no competing financial interests.

Major limitations of the fluorometric technique are experimental variance as well as cell loss associated with extensive washing and use of non-adherent cells.
Variance is observed largely due to the variation in fluorescent particles as weighing and pipetting during preparation of the stock solution is not an exact way of maintaining identical numbers of particles from experiment to experiment. To address the problem of variance between experiments, data can be expressed in relative terms, for example as % phagocytosis or fold change from control (control &#8211; t = 0; particles added and removed immediately). This form of analysis generates reproducible trends, and values of % phagocytosis from control is similar from experiment to experiment. This kind of data analysis will facilitate statistical significance with 3-6 experimental replicates. Fluorometric technique can be used to effectively examine and reproduce observed trends despite of its variability if utilizing adequate data analysis. With mastering of the technique it is possible for a single scientist to run approximately six 96-well plates, or 600 treatments in a day.
Cell loss has been largely observed in studies with non-adherent cell types. Adherent cell types are the best to use in methods such as this one because they can withstand of the numerous and extensive washing steps.
Major critical steps should be taken to keep the fluorescent components away from direct light to avoid photo bleaching. Particles, labels, and the plate containing each or all components should be kept in dark (or simply wrapped in aluminum foil). Another essential component is the importance of technical replicates within a plate. Due to high well to well variance it is essential to mix all the reagents very well before pipetting and to have 6-8 technical replicates per treatment (equivalent to one row), in order to most accurately observe trends in the data.
Additional considerations important for use of plate formats such as these, is that the plate wells contain a relatively small volumes of reagents and are often prone to evaporation. If the treatments are longer than 24 hr, it is advisable to use the peripheral wells of the plate (rows and columns next to the edge) containing PBS alone, in order to serve as humidifiers and to reduce evaporation of cell culture media and agonists.
This technique is fairly simple to master and the troubleshooting involved is minimal. It is important to choose the right phagocytic method for the study and to be careful while washing (especially when using non-adherent cells) in order to avoid cell loss.
Major significance of the fluorometric technique is that it provides a high-throughput method for screening of fluorescent particle internalization, or actin polymerization. Most techniques used to date such as microscopy or microbiology are more labor intensive and more time consuming in comparison. Flow cytometry utilizes very similar principles as fluorometry, but takes hours to quantify compared to minutes required by fluorometry. Therefore, the fluorometric method is a good alternative to current methods of phagocytic quantification largely due to its high-throughput capabilities.

Quantification of fluorescent signal has been widely used in a multitude of scientific methods ranging from PCR, flow cytometry, confocal microscopy, and FRET analysis to multiplex ELISA. Fluorescence imaging and quantification has a broad application and can be a great tool for quantitative analysis of various cellular processes. Use of fluorescent markers and their signal has been revolutionized in the last decade, and emergence of fluorescent plate readers has facilitated the high throughput quantification of fluorescence emitted during cellular processes.
Fluorometric analysis can serve as a great tool in quantification of phagocytosis. Phagocytosis has been studied since the discovery of phagocytes by Metchnikoff in 18001. Over the years, a variety of methods have been utilized to examine this important process essential for innate immune defense against invading bacterial, viral, fungal and parasitic pathogens2-5. Previous methods of quantification utilized microscopy and stereological techniques in order to visualize cells that are phagocytosing, which were then quantified by counting of internalized particles (manually or with use of software)6-8. Some disadvantages to using microscopy alone for quantitative analysis are that manual counting of bacteria is labor intensive and more prone to observer bias. Another method used in quantification of phagocytosis is the microbiological technique of plating of bacteria from cell lysates (following phagocytosis) on bacterial culture plates, but this method can fail to account for bactericidal mechanisms and presence of partially internalized bacteria. This method is even more labor intensive compared to microscopy and takes several days to analyze. Flow cytometry seems to be the quickest, most effective way to quantify phagocytosis and has been utilized by many groups9-11, but the high cost commonly associated with the instrument needed for the analysis makes it the most expensive method when compared to the previously mentioned assays.
The fluorometric method is a good alternative to flow cytometry for analysis of particle internalization since it offers unbiased quantification of fluorescence using equipment that is not as cost prohibitive. Other added benefits of fluorometry are high efficiency, and high throughput capabilities for quantifying fluorescence emitted by the labeled internalized particles.
Benefits of fluorometry can be extrapolated to quantification of processes other than phagocytosis. For example, fluorometric analysis can be applied to study any process leading to changes in expression of intracellular or membrane-bound receptors, changes in cell permeability/viability, transfection efficacy, and modulation in actin polymerization. One downside of fluorometric technique is that, depending on the labels used, there may be a high experiment to experiment variability which can usually be resolved by demonstrating data through relative quantification, such as fold change or percent increase from control.

<table><tbody><tr><td>1 &amp;mu;m yellow-green fluorescent Fluo-Spheres;</td><td>Molecular Probes</td><td>F8852</td><td>combine 50&amp;mu;l with 50&amp;mu;l opsonizing reagent for 30 min at 37&lt;sup&gt;o&lt;/sup&gt;C before use</td></tr><tr><td>Heat killed &lt;em&gt;E. coli&lt;/em&gt; BioParticles fluorescein conjugate</td><td>Molecular Probes</td><td>E2861</td><td>reconstitute (5 mg) in 50 &amp;mu;l of PBS and combine with 50 &amp;mu;l opsonizing reagent for 30 min at 37 &amp;deg;C before use</td></tr><tr><td>Opsonizing reagent</td><td>Molecular Probes</td><td>E2870</td><td></td></tr><tr><td>Rhodamine phalloidin</td><td>Molecular Probes</td><td>R415</td><td></td></tr><tr><td>DAPI</td><td>Sigma-Aldrich</td><td>D9542</td><td></td></tr><tr><td>Trypan blue</td><td>Gibco</td><td>15250-061</td><td></td></tr><tr><td colspan="4">The items below are available in many brands but the items we used in this study are from the following manufacturers:</td></tr><tr><td>RPMI Cell growth media</td><td>Gibco</td><td></td><td>Supplemented with 10% FBS and 1% pennicillin/streptomycin; warm in 37 &amp;deg;C before use</td></tr><tr><td>Fluorescent plate reader - Fluostar Omega</td><td>BMG Labtech</td><td></td><td></td></tr><tr><td>Paraformaldehyde (16%)</td><td>Fischer Scientific</td><td>AA433689M</td><td>dilute to 4% before use</td></tr><tr><td>96-well plates</td><td>Greiner</td><td>655097</td><td>clear or black or clear bottom - black plates</td></tr><tr><td>Multichannel pipette (8 - 12 channels)</td><td></td><td></td><td></td></tr><tr><td>Reagent reservoirs</td><td></td><td></td><td></td></tr><tr><td>1x PBS</td><td></td><td></td><td></td></tr><tr><td>Microfuge tubes (0.6 ml)</td><td></td><td></td><td></td></tr><tr><td>Conicles (10 ml)</td><td></td><td></td><td></td></tr></tbody></table>

high throughput fluorometric technique for assessment of macrophage phagocytosis and actin polymerization

The goal of fluorometric analysis is to serve as an efficient, cost effective, high throughput method of analyzing phagocytosis and other cellular processes. This technique can be used on a variety of cell types, both adherent and non-adherent, to examine a variety of cellular properties. When studying phagocytosis, fluorometric technique utilizes phagocytic cell types such as macrophages, and fluorescently labeled opsonized particles whose fluorescence can be extinguished in the presence of trypan blue. Following plating of adherent macrophages in 96-well plates, fluorescent particles (green or red) are administered and cells are allowed to phagocytose for varied amounts of time. Following internalization of fluorescent particles, cells are washed with trypan blue, which facilitates extinction of fluorescent signal from bacteria which are not internalized, or are merely adhering to the cell surface. Following the trypan wash, cells are washed with PBS, fixed, and stained with DAPI (nuclear blue fluorescent label), which serves to label nuclei of cells. By a simple fluorometric quantification through plate reading of nuclear (blue) or particle (red/green) fluorescence we can examine the ratio of relative fluorescence units of green:blue and determine a phagocytic index indicative of amount of fluorescent bacteria internalized per cell. The duration of assay using a 96-well method and multichannel pipettes for washing, from end of phagocytosis to end of data acquisition, is less than 45 min. Flow cytometry could be used in a similar manner but the advantage of fluorometry is its high throughput, rapid method of assessment with minimal manipulation of samples and quick quantification of fluorescent intensity per cell. Similar strategies can be applied to non adherent cells, live labeled bacteria, actin polymerization, and essentially any process utilizing fluorescence. Therefore, fluorometry is a promising method for its low cost, high throughput capabilities in the study of cellular processes.

Two major ways of quantifying phagocytosis and the subsequent actin polymerization through the use of this protocol is to observe continuous or synchronized phagocytosis.
Microscopic analysis (Figure 1B) illustrates fluorescent images comparable to what the fluorometer is recording (also see Diagram 1). In Figure 1B are red fluorescence of actin, green 268 fluorescence of FITC labels HKOP E. coli, nuclear DAPI stain, and the merged image of all three 269 fluorophores together. This image indicates effective phagocytosis, actin polymerization, and 270 effective trypan quenching of non-internalized particles.
Continuous phagocytosis of opsonized and unopsonized particles is quite comparable as seen in Figure 2A and C respectively, where J774 cells are internalizing dextran beads. Interestingly, actin polymerization following opsonophagocytosis increases, slowing down at later time points (Figure 2B), while the actin polymerization following internalization of unopsonized particles does not increase (Figure 2D). This is important to consider for future studies examining these processes since opsonophagocytosis leads to much greater actin polymerization than internalization of particles that are not opsonized. Neither of the phagocytic methods using dextran particles result in changes in cell viability as was confirmed via the MTT analysis (Figure 2E).
Unlike continuously increasing trends observed during continuous phagocytosis (Figure 2), synchronized phagocytosis of opsonized dextran beads has a bell shaped trend (Figure 3A), showing reduction in internalization following 30 min. This is to be expected considering processing time and nature of the protocol. This trend is also confirmed by the actin polymerization data (Figure 3B) of these cells where the highest polymerization time point is early on in the timecourse, while at 30 min polymerization returns to baseline. Although more labor intensive, this method has a value of examining phagocytosis right after the synchronization, therefore examining immediate phagocytic effects.
<img alt="Figure 1" src="/files/ftp_upload/52195/52195fig1highres.jpg" width="650" /> 
Figure 1: Confocal microscopy of phagocytosis and actin polymerization. Cells were allowed to internalize FITC conjugated OPDex bead (green) at 20:1 bead:cell ratio for 60 min, then cells were washed with trypan and PBS, fixed with paraforlmaldehyde, permeabilised with acetone, and stained for f-actin using rhodamine phalloidin (red), and DAPI (blue). Images illustrate confocal microscopic analysis at 60X with additional digital magnification; (A) white bar = 10 &#181;m (B) white bar = 50 &#181;m. (A) Illustrates changes in actin polymerization following addition of inhibitor. Arrowheads indicate pseudopodia formation, and arrows indicate changes in membrane ruffling. Figure 1A has been modified from Ninkovic and Roy12. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52195/52195fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52195/52195fig2highres.jpg" /> 
Figure 2: Fluorometric analysis following continuous phagocytosis. J774 murine macrophages were plated onto a 96-well plate and exposed to dextran beads at varying amounts of time, starting with 90, 60, 45, 30, and 15 min. Beads were added at different times and the phagocytic processes for all the reactions were stopped at the same time. Cells were washed with trypan, fixed, stained, and analyzed for phagocytosis of opsonized (A) or unopsonized (C) dextran particles and actin polymerization associated with each (B, D respectively). (E) MTT assay was conducted immediately following phagocytosis at different time points. This figure has been modified from Ninkovic and Roy12.
<img alt="Figure 3" src="/files/ftp_upload/52195/52195fig3highres.jpg" /> 
Figure 3: Fluorometric analysis following synchronized phagocytosis. FITC labeled OPDex beads were added to J774 murine macrophage cells plated in 96-well plates at a cell density of 104 cell/well. After a 30 min incubation the plate was centrifuged at 500 x g for 5 min. Phagocytosis was stopped one at a time, at time points stated in the x-axis by trypan wash, followed by PBS washes, paraformaldehyde fixation and actin staining. This figure has been modified from Ninkovic and Roy12.

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High Throughput Fluorometric Technique for Assessment of Macrophage Phagocytosis and Actin Polymerization

Here we present a protocol to quantify phagocytosis of fluorescent particles by adherent macrophage cell line using a fluorometric method. This method facilitates a high throughput quantification of particle internalization as well as the resulting actin polymerization.

The goal of fluorometric analysis is to serve as an efficient, cost effective, high throughput method of analyzing phagocytosis and other cellular ...

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Universidad San Sebastian

Research

JoVE Journal

Immunology and Infection

14.6K Views.  University of Minnesota. Here we present a protocol to quantify phagocytosis of fluorescent particles by adherent macrophage cell line using a fluorometric method. This method facilitates a high throughput quantification of particle internalization as well as the resulting actin polymerization.

Video: High Throughput Fluorometric Technique for Assessment of Macrophage Phagocytosis and Actin Polymerization

"Phagosome Closure Assay" to Visualize Phagosome Formation in Three Dimensions Using Total Internal Reflection Fluorescent Microscopy (TIRFM)

Phagocytosis is a mechanism used by specialized cells to internalize and eliminate microorganisms or cellular debris. It relies on profound rearrangements of the actin cytoskeleton that is the driving force for plasma membrane extension around the particle. In addition, efficient engulfment of large material relies on focal exocytosis of intracellular compartments. This process is highly dynamic and numerous molecular players have been described to have a role during phagocytic cup formation. The precise regulation in time and space of all of these molecules, however, remains elusive. In addition, the last step of phagosome closure has been very difficult to observe because inhibition by RNA interference or dominant negative mutants often results in stalled phagocytic cup formation. 
We have set up a dedicated experimental approach using total internal reflection fluorescence microscopy (TIRFM) combined with epifluorescence to monitor step by step the extension of pseudopods and their tips in a phagosome growing around a particle loosely bound to a coverslip. This method allows us to observe, with high resolution the very tips of the pseudopods and their fusion during closure of the phagosome in living cells for two different fluorescently tagged proteins at the same time.

"Phagosome Closure Assay" to Visualize Phagosome Formation in Three Dimensions Using Total Internal Reflection Fluorescent Microscopy TIRFM

We describe an experimental setup to visualize with unprecedented high resolution phagosome formation and closure in three dimensions in living macrophages, using total internal reflection fluorescence microscopy. It allows monitoring of the base of the phagocytic cup, the extending pseudopods, as well as the precise site of phagosome scission.

Phagocytosis is a mechanism used by specialized cells to internalize and eliminate microorganisms or cellular debris. It relies on profound rearrangements ...

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 ...

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

Using Enhanced Green Fluorescence Protein-expressing Escherichia Coli to Assess Mouse Peritoneal Macrophage Phagocytosis

This manuscript describes a simple and reproducible method to perform a phagocytosis assay. The first part of this method involves building a pET-SUMO-EGFP vector (SUMO = small ubiquitin-like modifier) and expressing enhanced green fluorescence protein (EGFP) in Escherichia coli (BL21DE). EGFP-expressing E. coli is coincubated with macrophages for 1 h at 37 &#176;C; the negative control group is incubated on ice for the same amount of time. Then, the macrophages are ready for assessment. The advantages of this technique include its simple and straightforward steps, and phagocytosis can be measured by both flow cytometer and fluorescence microscope. The EGFP-expressing E. coli are stable and display a strong fluorescence signal even after the macrophages are fixed with paraformaldehyde. This method is not only suitable for the assessment of macrophage cell lines or primary macrophages in vitro but also suitable for the evaluation of granulocyte and monocyte phagocytosis in peripheral blood mononuclear cells. The results show that the phagocytic capability of peritoneal macrophages from young (eight-week-old) mice is higher than that of macrophages from aged (16-month-old) mice. In summary, this method measures macrophage phagocytosis and is suitable for studying the innate immune system function.

Using Enhanced Green Fluorescence Protein-expressing Escherichia Coli to Assess Mouse Peritoneal Macrophage Phagocytosis

Here, we present a protocol to assess mouse peritoneal macrophage phagocytosis using enhanced green fluorescence protein-expressing Escherichia coli.

This manuscript describes a simple and reproducible method to perform a phagocytosis assay. The first part of this method involves building a ...

Mesenchymal Stem Cell Regulation of Macrophage Phagocytosis; Quantitation and Imaging

Mesenchymal stem cells (MSC) have traditionally been studied for their regenerative properties, but more recently, their immunoregulatory characteristics have been at the forefront. They interact with and regulate immune cell activity. The focus of this study is the MSC regulation of macrophage phagocytic activity. Macrophage (M&#934;) phagocytosis is an important part of the innate immune system response to infection, and the mechanisms through which MSC modulate this response are under active investigation. Presented here is a method to study M&#934; phagocytosis of non-opsonized zymosan particles conjugated to a pH-sensitive fluorescent molecule while in co-culture with MSC. As phagocytic activity increases and the labeled zymosan particles are enclosed within the acidic environment of the phagolysosome, the fluorescence intensity of the pH-sensitive molecule increases. With the appropriate excitation and emission wavelengths, phagocytic activity is measured using a fluorescent spectrophotometer and kinetic data is presented as changes in relative fluorescent units over a 70 min period. To support this quantitative data, the change in the phagocytic activity is visualized using dynamic imaging. Results using this method demonstrate that when in co-culture, MSC enhance M&#934; phagocytosis of non-opsonized zymosan of both naive and IFN-&#947; treated M&#934;. These data add to the current knowledge of MSC regulation of the innate immune system. This method can be applied in future investigations to fully delineate the underlying cellular and molecular mechanisms.

Presented here is a protocol to quantify and produce dynamic images of mesenchymal stem cell (MSC) mediated regulation of macrophage (M&#934;) phagocytosis of non-opsonized yeast (zymosan) particles that are conjugated to a pH-sensitive fluorescent molecule.

Mesenchymal stem cells (MSC) have traditionally been studied for their regenerative properties, but more recently, their immunoregulatory characteristics ...

Flow Cytometry Approach to Characterize Phagocytic Properties of Acutely-Isolated Adult Microglia and Brain Macrophages In Vitro

Microglia and central nervous system (CNS)-infiltrating macrophages, collectively called CNS mononuclear phagocytes (CNS-MPs), play central roles in neurological diseases including neurodegeneration and stroke. CNS-MPs are involved in phagocytic clearance of pathological proteins, debris and neuronal synapses, each with distinct underlying molecular pathways. Characterizing these phagocytic properties can provide a functional readout that compliments molecular profiling of microglia using traditional flow cytometry, transcriptomics and proteomics approaches. Phagocytic profiling of microglia has relied on microscopic visualization and in vitro cultures of mouse neonatal microglia. The former approach suffers from limited sampling while the latter approach is inherently poorly reflective of the true in vivo state of adult CNS-MPs. This paper describes optimized protocols to phenotype phagocytic properties of acutely-isolated mouse CNS-MPs by flow cytometry. CNS-MPs are acutely isolated from adult mouse brain using mechanical dissociation followed by density gradient centrifugation, incubated with fluorescent microspheres or fluorescent A&#946; fibrils, washed, and then labeled with panels of antibodies against surface markers (CD11b, CD45). Using this approach, it is possible to compare phagocytic properties of brain-resident microglia with CNS-infiltrating macrophages and then assess the effect of aging and disease pathology on these phagocytic phenotypes. This rapid method also holds potential to functionally phenotype acutely-isolated human CNS-MPs from post-mortem or surgical brain specimens. Additionally, specific mechanisms of phagocytosis by CNS-MP subsets can be investigated by inhibiting select phagocytic pathways.

Assessment of phagocytic properties of microglia and brain macrophages can provide a valuable functional dimension to molecular profiling studies. We describe a validated protocol that uses flow cytometry to rapidly and reliably quantify phagocytosis of fluorescent microspheres and amyloid beta fibrils by acutely-isolated microglia and brain macrophages from mouse models.

Microglia and central nervous system (CNS)-infiltrating macrophages, collectively called CNS mononuclear phagocytes (CNS-MPs), play central roles in ...

Neuroscience

In vitro Quantitative Imaging Assay for Phagocytosis of Dead Neuroblastoma Cells by iPSC-Macrophages

Microglia orchestrate neuroimmune responses in several neurodegenerative diseases, including Parkinson&#39;s disease and Alzheimer&#39;s disease. Microglia clear up dead and dying neurons through the process of efferocytosis, a specialized form of phagocytosis. The phagocytosis function can be disrupted by environmental or genetic risk factors that affect microglia. This paper presents a rapid and simple in vitro&#160;microscopy protocol for studying microglial efferocytosis in an induced pluripotent stem cell (iPSC) model of microglia, using a human neuroblastoma cell line (SH-SY5Y) labeled with a pH-sensitive dye for the phagocytic cargo. The procedure results in a high yield of dead neuroblastoma cells, which display surface phosphatidylserine, recognized as an &#34;eat-me&#34; signal by phagocytes. The 96-well plate assay is suitable for live-cell time-lapse imaging, or the plate can be successfully fixed prior to further processing and quantified by high-content microscopy. Fixed-cell high-content microscopy enables the assay to be scaled up for screening of small molecule inhibitors or assessing the phagocytic function of genetic variant iPSC lines. While this assay was developed to study phagocytosis of whole dead neuroblastoma cells by iPSC-macrophages, the assay can be easily adapted for other cargoes relevant to neurodegenerative diseases, such as synaptosomes and myelin, and other phagocytic cell types.

Neurodegenerative diseases are associated with dysregulated microglia functions. This article outlines an in vitro assay of phagocytosis of neuroblastoma cells by iPSC-macrophages. Quantitative microscopy readouts are described for both live-cell time-lapse imaging and fixed-cell high-content imaging.

Microglia orchestrate neuroimmune responses in several neurodegenerative diseases, including Parkinson&#39;s disease and Alzheimer&#39;s disease. ...

A Time-Efficient Fluorescence Spectroscopy-Based Assay for Evaluating Actin Polymerization Status in Rodent and Human Brain Tissues

Actin, the major component of cytoskeleton, plays a critical role in the maintenance of neuronal structure and function. Under physiological states, actin occurs in equilibrium in its two forms: monomeric globular (G-actin) and polymerized filamentous (F- actin). At the synaptic terminals, actin cytoskeleton forms the basis for critical pre- and post-synaptic functions. Moreover, dynamic changes in the actin polymerization status (interconversion between globular and filamentous forms of actin) are closely linked to plasticity-related alterations in synaptic structure and function. We report here a modified fluorescence-based methodology to assess polymerization status of actin in ex vivo conditions. The assay employs fluorescently labelled phalloidin, a phallotoxin that specifically binds to actin filaments (F-actin), providing a direct measure of polymerized filamentous actin. As a proof of principle, we provide evidence for the suitability of the assay both in rodent and post-mortem human brain tissue homogenates. Using latrunculin A (a drug that depolymerizes actin filaments), we confirm the utility of the assay in monitoring alterations in F-actin levels. Further, we extend the assay to biochemical fractions of isolated synaptic terminals wherein we confirm increased actin polymerization upon stimulation by depolarization with high extracellular K+.

We report a simple, time-efficient and high-throughput fluorescence spectroscopy-based assay for the quantification of actin filaments in ex vivo biological samples from brain tissues of rodents and human subjects.

Actin, the major component of cytoskeleton, plays a critical role in the maintenance of neuronal structure and function. Under physiological states, actin ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

Biology

Automated Imaging and Analysis for the Quantification of Fluorescently Labeled Macropinosomes

Macropinocytosis is a non-specific fluid-phase uptake pathway that allows cells to internalize large extracellular cargo, such as proteins, pathogens, and cell debris, through bulk endocytosis. This pathway plays an essential role in a variety of cellular processes, including the regulation of immune responses and cancer cell metabolism. Given this importance in biological function, examining cell culture conditions can provide valuable information by identifying regulators of this pathway and optimizing conditions to be employed in the discovery of novel therapeutic approaches. The study describes an automated imaging and analysis technique using standard laboratory equipment and a cell imaging multi-mode plate reader for the rapid quantification of the macropinocytic index in adherent cells. The automated method is based on the uptake of high molecular weight fluorescent dextran and can be applied to 96-well microplates to facilitate assessments of multiple conditions in one experiment or fixed samples mounted onto glass coverslips. This approach is aimed at maximizing reproducibility and reducing experimental variation while being both time-saving and cost-effective.

Automated assays using multi-well microplates are advantageous approaches for identifying pathway regulators by allowing the assessment of a multitude of conditions in a single experiment. Here, we have adapted the well-established macropinosome imaging and quantification protocol to a 96-well microplate format and provide a comprehensive outline for automation using a multi-mode plate reader.

Macropinocytosis is a non-specific fluid-phase uptake pathway that allows cells to internalize large extracellular cargo, such as proteins, pathogens, and ...

High Throughput Fluorometric Technique for Assessment of Macrophage Phagocytosis and Actin Polymerization

Summary

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Chapters in this video

"Phagosome Closure Assay" to Visualize Phagosome Formation in Three Dimensions Using Total Internal Reflection Fluorescent Microscopy (TIRFM)

Time-lapse 3D Imaging of Phagocytosis by Mouse Macrophages

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Using Enhanced Green Fluorescence Protein-expressing Escherichia Coli to Assess Mouse Peritoneal Macrophage Phagocytosis

Mesenchymal Stem Cell Regulation of Macrophage Phagocytosis; Quantitation and Imaging

Flow Cytometry Approach to Characterize Phagocytic Properties of Acutely-Isolated Adult Microglia and Brain Macrophages In Vitro

In vitro Quantitative Imaging Assay for Phagocytosis of Dead Neuroblastoma Cells by iPSC-Macrophages

A Time-Efficient Fluorescence Spectroscopy-Based Assay for Evaluating Actin Polymerization Status in Rodent and Human Brain Tissues

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Automated Imaging and Analysis for the Quantification of Fluorescently Labeled Macropinosomes