We thank Dr. Alexandre Benmerah (Institut Imagine Necker, Paris, France) for initial discussions on the experimental approach and Dr. Jamil Jubrail for reading the manuscript. Nad&#232;ge Kambou and Susanna Borreill are acknowledged for performing experiments with the method in our laboratory. This work was supported by grants from CNRS (ATIP Program), Ville de Paris and Agence Nationale de la Recherche (2011 BSV3 025 02), Fondation pour la Recherche M&#233;dicale (FRM INE20041102865, FRM DEQ20130326518 including a doctoral fellowship for FMA) to FN, and Agence Nationale de Recherche sur le SIDA et les h&#233;patites virales" (ANRS) including a doctoral fellowship for JM.

Note: The plasmid used Lifeact-mCherry is a kind gift of Dr. Guillaume Montagnac, Institut Curie, Paris, generated after 17.
1. Cells and Transfection
Note: RAW264.7 macrophages are grown to sub confluency in complete medium (RPMI (Roswell Park Memorial Institute) 1640 medium, 10 mM HEPES, 1 mM sodium pyruvate, 50 &#181;M &#946;-mercaptoethanol, 2 mM L-Glutamine and 10% FCS (Fetal Calf Serum)) in a 100 mm plate. They are transfected with plasmids encoding fluorescently tagged proteins by electroporation. Routinely approximately 5 - 6 x 106 cells are transfected with 20 &#181;g or 10 &#181;g of plasmid for each transfection or co-transfection respectively. Note that other means of transfection based on electroporation or lipofection can be used as alternative approaches.
<ol>
	<li>Scrape the cells with a cell lifter and resuspend them in 10 ml of culture medium by pipetting up and down several times.</li>
	<li>Centrifuge the cell suspension 300 x g, 5 min at RT in swinging angle rotor.</li>
	<li>Preheat 10 ml of complete medium supplemented with 10 &#181;g/ml of gentamicin at 37 &#176;C.</li>
	<li>Following centrifugation, discard the supernatant and resuspend the pellet in 3 ml of &#34;washing buffer A&#34; from the electroporation kit.</li>
	<li>Centrifuge the cell suspension 300 x g, 5 min at room temperature in swinging angle rotor.</li>
	<li>In the meantime prepare the DNA mix at room temperature: 120 &#181;l 2x Buffer B, 20 &#181;g of plasmid DNA coding for the protein of interest, q.s.p. 240 &#181;l H2O.</li>
	<li>Following centrifugation, discard the entire supernatant.</li>
	<li>Resuspend the cell pellet in the DNA mix and transfer into 4 mm electroporation cuvettes.</li>
	<li>Incubate at RT for 3 min.</li>
	<li>Electroporate at 250 V, 900 &#181;F.</li>
	<li>Immediately resuspend the cells in the pre-warmed (37 &#176;C) complete medium supplemented with gentamicin and plate them in a 100 mm dish.</li>
	<li>Incubate the transfected cells overnight at 37 &#176;C, 5% CO2.</li>
	<li>The next morning, replace the complete medium with 10 ml of serum-free microscopy medium (RPMI 1640 without phenol red, 10 mM HEPES, 1 mM sodium pyruvate, 50 &#181;M &#946;-mercaptoethanol, 2 mM L-Glutamine).</li>
</ol>
2. Opsonization of Red Blood Cells
Note: As a model of particle target for macrophages, sheep red blood cells (SRBCs) are used. Usually, around 7 x 106 SRBCs per 35 mm glass bottom dish is used.
<ol>
	<li>Wash the SRBCs with 100 &#181;l of 1x PBS (phosphate buffered saline)/0.1% BSA (bovine serum albumin) and centrifuge (600 x g, 4 min).</li>
	<li>Following centrifugation, discard the supernatant and resuspend the SRBCs in 100 &#181;l of 1x PBS/0.1% BSA and centrifuge (600 x g, 4 min).</li>
	<li>Following centrifugation, discard the supernatant and resuspend the SRBCs in 1x PBS/0.1% BSA with rabbit IgG anti-SRBCs at sub-agglutinating concentration. Use 500 &#181;l of solution containing antibody/5 &#181;l of SRBCs. 
	&#8203;Note: The sub-agglutinating concentration of antibodies corresponds to the lowest concentration that did not induce agglutination detected as formation of a network. In general, the sub-agglutinating concentration is 8.2 &#181;g/ml.
	<ol>
		<li>Determine the concentration of IgG anti-SRBCs required to opsonize the SRBCs by a hemagglutination test. Serially dilute the IgG anti-SRBCs (stock at 13.1 mg/ml) between 1/50 - 1/25,600 in a microplate. Add 2 x 106 SRBCs in each well. Incubate the plate in the dark at RT for several hr.</li>
	</ol>
	</li>
	<li>Incubate at RT for 30 mins with slow rotation.</li>
	<li>After two washes in 1x PBS/0.1% BSA as described above, resuspend the IgG-opsonized SRBCs (IgG-SRBCs) in pre-warmed serum-free microscopy medium (2 ml/dish).</li>
</ol>
3. Poly-lysine Coating of Coverslips
<ol>
	<li>In the meantime, treat 35 mm glass bottom dishes with 2 ml of 0.01% poly-L-lysine for 30 min at room temperature.</li>
	<li>Wash the dishes two times with 2 ml of 1x PBS.</li>
</ol>
4. Non-covalent Fixation of SRBCs on Glass Bottom Dishes
<ol>
	<li>Pour 2 ml of SRBCs suspension per 35 mm glass bottom dish.</li>
	<li>Centrifuge with a swinging rotor at 500 x g during 2 min onto 35 mm glass bottom dishes.</li>
	<li>Remove the supernatant and wash once with 2 ml of 1x PBS/10% BSA.</li>
	<li>Incubate the particles for 30 min with 2 ml of 1x PBS/10% BSA per dish.</li>
	<li>Wash the dishes three times with 2 ml of 1x PBS.</li>
	<li>Replace 1x PBS with 2 ml of pre-warmed (37 &#176;C) serum-free microscopy medium.</li>
</ol>
5. Phagocytosis Visualized by TIRFM
<ol>
	<li>Microscope 
	Note: TIRFM was performed using a microscope equipped with an oil-immersion objective (N 100X, NA1.49), a heating chamber with CO2, and two single photon detection cameras EMCCD (Electron Multiplying Charge Coupled Device) coupled with a 1.5X lens.
	<ol>
		<li>The day before the TIRF microscope session, turn on the heating chamber at 37 &#186;C&#8203; to allow a homogeneous heating of the microscope stage.</li>
	</ol>
	</li>
	<li>Critical Angle Determination 
	Note: ImageJ Color Profiler software is used to process TIRF image streams.
	<ol>
		<li>Place a 35 mm glass bottom dish containing opsonized SRBC with serum-free microscopy medium under the microscope.</li>
		<li>Scrape the cells and resuspend them in the medium before adding them (100 - 500 &#181;l) in the dish.</li>
		<li>Use the &#34;Live Acquisition&#34; software to control the microscope and perform excitation with a 491 nm and/or a 561 nm laser to identify cells expressing fluorescently tagged proteins.</li>
		<li>Identify a cell that expresses the fluorescently tagged proteins.</li>
		<li>Place it in the middle of the field and acquire 500 images at one excitation wavelength, with different angles starting from 0&#186; up to 5&#186;, with an increment of 0.01&#186; (Figure 2A). The angles are automatically changed by the microscope system.</li>
		<li>Determine the critical angle allowing the incident light to be totally reflected at the glass/ medium interface and generate the evanescent wave. Using ImageJ Color Profiler software, open the image sequence by clicking on &#34;File&#34;, &#34;Open&#34; and select the file.</li>
		<li>Select a region of interest (ROI), with the rectangular tool, in the cell with a uniform fluorescence (Figure 2B yellow 1).</li>
		<li>Plot the &#34;Z axis profile&#34; mean fluorescence intensity measured in the ROI with function of the angles on the x axis by clicking on &#34;Image&#34; tab, &#34;Stacks&#34; in the drop down menu and &#34;Plot Z-axis Profile&#34; (Figure 2B red 2). 
		Note: The critical angle is the angle leading to the maximum fluorescence before a sharp decrease in fluorescence.</li>
		<li>Use any value of angle on the x-axis superior to the critical angle during the microscopy session to obtain a TIRF signal as example 2.00 (Figure 2C).</li>
	</ol>
	</li>
	<li>Acquisitions
	<ol>
		<li>Using Live Acquisition software, set up the parameters of acquisition with the module &#8220;Protocol Editor&#8221; (Figure 3A).
		<ol>
			<li>Create a protocol with a &#8220;loop&#8221; comprising &#8220;Multi Channels&#8221; acquisition to acquire fluorescent signal from proteins of interest in the TIRF region. Enter the TIRF angle (for example 2.00), the exposure time (for example 50 msec) and the laser intensity (for example 50%) (Figure 3B 1).</li>
			<li>Introduce a &#8220;Z move&#8221; of the objective 3 &#181;m above the TIRF region to obtain signal acquisition in epifluorescence with the &#8220;Multi Channels &#8220;tool. Enter an angle below the critical angle (as example 1.00), the time of exposition (50 msec) and the laser intensity (50%) (Figure 3B, 2 and 3).</li>
			<li>Next add a &#34;z move&#34; of the objective 3 &#181;m below, to return in the TIRF region (Figure 3B 4). Add a &#34;snapshot&#34; of the cell in bright light LED (Light-Emitting Diode) (Figure 3B 5).</li>
		</ol>
		</li>
		<li>Find a cell of interest with a moderate level of fluorescently tagged protein expression that initiates phagocytosis of SRBC by extending plasma membrane around the particle.</li>
		<li>Place it in the middle of the field.</li>
		<li>Start streaming acquisition of 500 - 1,000 frames. In the &#8220;loop count&#8221; tab, enter &#8220;750 frames&#8221; (Figure 3B 1). 
		Note: ImageJ Color Profiler software is used to process TIRF image streams.</li>
		<li>Open the sequence images in Image J software by clicking &#8220;File&#8221;, &#8220;Open&#8221; and choosing the file.</li>
		<li>Separate the channels by clicking on the tab &#8220;Image &#8220;, &#8220;Hyperstacks&#8221; drop down menu and on &#8220;Stack to hyperstack&#8221; function. Complete the appeared window: Order: xyctz; Channel (c): number of channels (ex: &#8220;2&#8221; if you have two fluorochromes); Slices (z): number of slices in z axis (ex: &#8220;1&#8221; if there is no movement in the z- axis); Frames (t): number of images divided per number of channels; Display mode: Grayscale. 
		Note: Two separate image sequences are generated, corresponding to the two different channels.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The experimental protocol described here proposes an unprecedented method to follow in real time and in living cells, with high-resolution, the formation of a phagosome and in particular its closure. Several technical aspects have to be discussed. Firstly, the assay is very sensitive to temperature. It is very important to check that the heating chamber is at 37 &#176;C and that all media, devices or cells are kept within the chamber to avoid temperature changes that could impair the efficiency of phagocytosis. We notice...

Phagocytosis is a major cell function that starts with the recognition and binding of material to surface receptors, which then leads to the internalization and degradation of the ingested material. While single-celled eukaryotes such as the mold Dictyostelium discoideum and amoebae use phagocytosis for feeding on bacteria, higher organisms have evolved with professional cells. Macrophages or dendritic cells are the first line of defense against pathogens in various tissues and organs, and are crucial to activate the adaptive immune system through antigen presentation and cytokine production 1-4. Under certain circumstances phagocytosis can be performed by non-professional phagocytic cells, e.g., endothelial and epithelial cells. This process is important to maintain homeostasis during development and in adulthood for normal tissue turnover and remodeling. Finally, specialized phagocytes such as Sertoli cells in the testis or retinal pigment epithelial cells are extremely potent phagocytes 5.
The formation of a phagosome where degradation of microorganisms or cellular debris occurs starts with the clustering of phagocytic receptors on the surface of the phagocytic cell. Downstream signaling events following clustering of opsonic receptors such as Fc receptors (FcR) or complement receptors (CRs) have been well characterized. However, there are also numerous non-opsonic receptors including Toll-like receptors (TLRs), lectins, mannose receptors and scavenger receptors. These receptors recognize determinants on the particle surface such as mannose or fucose residues, phosphatidylserine, and lipopolysaccharides 1,6-9.
Pathogen or cell debris recognition involves binding to and clustering of several types of phagocytic receptor, which then lead to intense and transient actin remodeling. In parallel, focal exocytosis of intracellular compartments contributes to the release of membrane tension and is important for efficient phagocytosis of large particles. The signaling events leading to actin polymerization and membrane deformation were dissected in experimental models that triggered a single phagocytic receptor. During FcR-mediated phagocytosis, there is intense actin polymerization that is regulated by small GTPases (Rac, Cdc42). Among their downstream effectors, the Wiskott-Aldrich syndrome protein (WASP) leads to activation of the Actin-Related Protein 2/3 complex (Arp2/3) that nucleates actin filaments 1,2,4,10. Local production of phosphatidylinositol-4, 5-bisphosphate (PI(4,5)P2) is crucial for initial actin polymerization that drives pseudopod formation. Its conversion to PI(3,4,5)P3 is required for pseudopod extension and phagosome closure 11. Several pathways contribute to the disappearance of PI(4,5)P2. Firstly detachment of phosphatidylinositol phosphate kinases (PIPKIs) from the phagosome arrests PI(4,5)P2 synthesis. Secondly, it can be phosphorylated and consumed by class I PI3K kinases (PI3K) and converted in PI(3,4,5)P3 12. A role for phosphatases and phospholipases has also been implied in PI(4,5)P2 hydrolysis and F-actin removal during phagocytosis in mammalian cells and in Dictyostelium13,14. The phospholipase C (PLC)&#948; hydrolyzes PI(4,5)P2 into diacylglycerol and inositol-1,4,5-trisphosphate. The PI(4,5)P2 and PI(3,4,5)P3 phosphatase OCRL (oculocerebrorenal syndrome of Lowe) has also been implicated in phagosome formation. Precise local formation of F-actin and its depolymerization is tightly regulated in space and time and we have shown that recruitment of intracellular compartments is important to deliver locally the OCRL phosphatase, thus contributing to local actin depolymerization at the base of the phagocytic cup 13,15. For this, we used the experimental set up described here.
The mechanism and molecular players required for phagosome closure and membrane scission remain poorly defined because of the difficulties in visualizing and monitoring the site of phagosome closure. Until recently, phagocytosis was observed on fixed or living cells that internalize particles on their dorsal face or on their sides, making the timely visualization of the site of phagosome closure difficult. In addition, fixing methods could cause retraction of membranes and bias the results on pseudopodia extension and closure. By contrast, the assay that we have set up and describe here allows us to visualize pseudopod extension and the closure step of phagocytosis in living cells 13, based on total internal reflection microscopy (TIRFM) 16. This optical technique uses an evanescent wave to excite fluorophores in a thin area at the interface between a transparent solid (coverslip) and a liquid (cell culture medium). The thickness of the excitation depth is around 100 nm from the solid surface, allowing visualization of molecular events close to the plasma membrane. TIRFM allows a high signal-to-background ratio, and limits the out-of-focus fluorescence collected and the cytotoxicity due to illumination of cells.
Taking advantage of TIRFM, we developed the &#34;phagosome closure assay&#34; in which coverslips are activated with poly-lysine and then coated with IgG-opsonized red blood cells (IgG-SRBCs). Macrophages expressing transiently fluorescently tagged proteins of interest are then allowed to engulf the IgG-SRBCs. While cells detach the target particles that are non-covalently bound to the glass surface, the tips of the pseudopods can be observed and recorded in the TIRF mode. TIRF acquisitions are combined with acquisitions in the epifluorescence mode, after shifting the stage 3 &#181;m above, which allows the visualization of the base of the phagocytic cup. Addition of pharmacological drugs such as those inhibiting actin or dynamin during the process is also possible to further dissect the process at the molecular level.
The protocol described in detail here is for a RAW264.7 murine macrophage cell line and opsonized particles, but virtually, it can be adapted to any other phagocytic cell and with other targets such as beads. This method will allow better characterization of the regulation in time and space of the molecular players involved in pseudopod extension and phagosome closure during various phagocytic processes.

<table><tbody><tr><td>Anti-sheep red blood cells IgG</td><td>MP Biomedicals</td><td>55806</td><td></td></tr><tr><td>Bovine Serum Albumin heat shock fraction, pH 7, &amp;ge;98%&amp;nbsp;</td><td>Sigma</td><td>A7906</td><td></td></tr><tr><td>Cell lifter</td><td>Corning</td><td>3008</td><td></td></tr><tr><td>Cuvettes 4 mm</td><td>Cell project</td><td>EP104</td><td></td></tr><tr><td>DPBS, no calcium, no magnesium</td><td>Thermo Fischer Scientific</td><td>14190-094</td><td>Room temperature</td></tr><tr><td>Electrobuffer kit</td><td>Cell project</td><td>EB110</td><td></td></tr><tr><td>100 mm TC-Treated Cell Culture Dish</td><td>Corning</td><td>353003</td><td></td></tr><tr><td>Gene X-cell pulser</td><td>Biorad</td><td>165-2661</td><td></td></tr><tr><td>Gentamicin solution</td><td>Sigma</td><td>G1397</td><td></td></tr><tr><td>Glass Bottom Dishes 35 mm uncoated 1.5</td><td>MatTek corporation</td><td>P35G-1.5-14-C Case</td><td></td></tr><tr><td>iMIC</td><td>TILL Photonics</td><td></td><td>Oil-immersion objective (N 100X, NA1.49), heating chamber with CO&lt;sub&gt;2&lt;/sub&gt;, a camera single photon detection EMCCD (Electron Multiplying Charge Coupled Device) and a 1.5X lens</td></tr><tr><td>Poly-L-Lysine Solution 0.1%</td><td>Sigma</td><td>P8920-100ml</td><td>Dilution at 0.01% in water&amp;nbsp;</td></tr><tr><td>RPMI 1640 medium GLUTAMAX&amp;nbsp; Supplement</td><td>Life technologies</td><td>61870-010</td><td></td></tr><tr><td>RPMI 1640 medium, no phenol red (10 x 500 ml)</td><td>Life technologies</td><td>11835-105</td><td>Warm in 37 &amp;deg;C water bath before use</td></tr><tr><td>Sheep red blood cells (SRBCs)</td><td>Eurobio</td><td>DSGMTN00-0Q</td><td>Conserved in Alsever buffer at 4 &amp;deg;C before use</td></tr></tbody></table>

"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.

The experimental system described in this manuscript is schematically represented in Figure 1. Transfected RAW264.7 macrophages expressing the proteins of interest fused to a fluorescent tag are placed into contact with IgG-opsonized sheep red blood cells (SRBCs) that were non-covalently fixed on the coverslip. The macrophages can detach the SRBC from the coverslip to engulf it. The TIRF microscope used allows concomitant acquisition of signals from the TIRF area correspo...

Watch this Scientific Journal Video about "Phagosome Closure Assay" to Visualize Phagosome Formation in Three Dimensions Using Total Internal Reflection Fluorescent Microscopy TIRFM at JoVE.com

"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.

"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 ...

phagosome-closure-assay-to-visualize-phagosome-formation-three

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Immunology and Infection

6.4K Views.  Université Paris Descartes, Sorbonne Paris Cité. 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.

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

Study of Phagolysosome Biogenesis in Live Macrophages

Phagocytic cells play a major role in the innate immune system by removing and eliminating invading microorganisms in their phagosomes. Phagosome maturation is the complex and tightly regulated process during which a nascent phagosome undergoes drastic transformation through well-orchestrated interactions with various cellular organelles and compartments in the cytoplasm. This process, which is essential for the physiological function of phagocytic cells by endowing phagosomes with their lytic and bactericidal properties, culminates in fusion of phagosomes with lysosomes and biogenesis of phagolysosomes which is considered to be the last and critical stage of maturation for phagosomes. In this report, we describe a live cell imaging based method for qualitative and quantitative analysis of the dynamic process of lysosome to phagosome content delivery, which is a hallmark of phagolysosome biogenesis. This approach uses IgG-coated microbeads as a model for phagocytosis and fluorophore-conjugated dextran molecules as a luminal lysosomal cargo probe, in order to follow the dynamic delivery of lysosmal content to the phagosomes in real time in live macrophages using time-lapse imaging and confocal laser scanning microscopy. Here we describe in detail the background, the preparation steps and the step-by-step experimental setup to enable easy and precise deployment of this method in other labs. Our described method is simple, robust, and most importantly, can be easily adapted to study phagosomal interactions and maturation in different systems and under various experimental settings such as use of various phagocytic cells types, loss-of-function experiments, different probes, and phagocytic particles.

Phagocytic cells play a major role in the innate immune system by removing and eliminating invading microorganisms in their phagosomes. Phagosome ...

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.

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 ...

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

Biochemistry

Imaging the Neutrophil Phagosome and Cytoplasm Using a Ratiometric pH Indicator

Neutrophils are crucial to host innate defense and, consequently, constitute an important area of medical research. The phagosome, the intracellular compartment where the killing and digestion of engulfed particles take place, is the main arena for neutrophil pathogen killing that requires tight regulation. Phagosomal pH is one aspect that is carefully controlled, in turn regulating antimicrobial protease activity. Many fluorescent pH-sensitive dyes have been used to visualize the phagosomal environment. S-1 has several advantages over other pH-sensitive dyes, including its dual emission spectra, its resistance to photo-bleaching, and its high pKa. Using this method, we have demonstrated that the neutrophil phagosome is unusually alkaline in comparison to other phagocytes. By using different biochemical conjugations of the dye, the phagosome can be delineated from the cytoplasm so that changes in the size and shape of the phagosome can be assessed. This allows for further monitoring of ionic movement.

This manuscript describes a simple method to measure the phagosomal pH and area as well as the cytoplasmic pH of human and mouse neutrophils using the ratiometric indicator seminaphthorhodafluor (SNARF)-1, or S-1. This is achieved using live-cell confocal fluorescence microscopy and image analysis.

Neutrophils are crucial to host innate defense and, consequently, constitute an important area of medical research. The phagosome, the intracellular ...

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

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 ...

Quantitative Analysis of Autophagy using Advanced 3D Fluorescence Microscopy

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, traditional chemotherapeutic approaches have been largely unsuccessful. Hormone therapy is effective at early stage, but often fails with the eventual development of hormone-refractory tumors. We have been interested in developing therapeutics targeting specific metabolic deficiency of tumor cells. We recently showed that prostate tumor cells specifically lack an enzyme (argininosuccinate synthase, or ASS) involved in the synthesis of the amino acid arginine1. This condition causes the tumor cells to become dependent on exogenous arginine, and they undergo metabolic stress when free arginine is depleted by arginine deiminase (ADI)1,10. Indeed, we have shown that human prostate cancer cells CWR22Rv1 are effectively killed by ADI with caspase-independent apoptosis and aggressive autophagy (or macroautophagy)1,2,3. Autophagy is an evolutionarily-conserved process that allows cells to metabolize unwanted proteins by lysosomal breakdown during nutritional starvation4,5. Although the essential components of this pathway are well-characterized6,7,8,9, many aspects of the molecular mechanism are still unclear - in particular, what is the role of autophagy in the death-response of prostate cancer cells after ADI treatment? In order to address this question, we required an experimental method to measure the level and extent of autophagic response in cells - and since there are no known molecular markers that can accurately track this process, we chose to develop an imaging-based approach, using quantitative 3D fluorescence microscopy11,12.
Using CWR22Rv1 cells specifically-labeled with fluorescent probes for autophagosomes and lysosomes, we show that 3D image stacks acquired with either widefield deconvolution microscopy (and later, with super-resolution, structured-illumination microscopy) can clearly capture the early stages of autophagy induction. With commercially available digital image analysis applications, we can readily obtain statistical information about autophagosome and lysosome number, size, distribution, and degree of colocalization from any imaged cell. This information allows us to precisely track the progress of autophagy in living cells and enables our continued investigation into the role of autophagy in cancer chemotherapy.

Autophagy is a ubiquitous process that enables cells to degrade and recycle proteins and organelles. We apply advanced fluorescence microscopy to visualize and quantify the small, but essential, physical changes associated with the induction of autophagy, including the formation and distribution of autophagosomes and lysosomes, and their fusion into autolysosomes.

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, ...

Biology

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. ...

Neuroscience

Visualizing Clathrin-mediated Endocytosis of G Protein-coupled Receptors at Single-event Resolution via TIRF Microscopy

Many important signaling receptors are internalized through the well-studied process of clathrin-mediated endocytosis (CME). Traditional cell biological assays, measuring global changes in endocytosis, have identified over 30 known components participating in CME, and biochemical studies have generated an interaction map of many of these components. It is becoming increasingly clear, however, that CME is a highly dynamic process whose regulation is complex and delicate. In this manuscript, we describe the use of Total Internal Reflection Fluorescence (TIRF) microscopy to directly visualize the dynamics of components of the clathrin-mediated endocytic machinery, in real time in living cells, at the level of individual events that mediate this process. This approach is essential to elucidate the subtle changes that can alter endocytosis without globally blocking it, as is seen with physiological regulation. We will focus on using this technique to analyze an area of emerging interest, the role of cargo composition in modulating the dynamics of distinct clathrin-coated pits (CCPs). This protocol is compatible with a variety of widely available fluorescence probes, and may be applied to visualizing the dynamics of many cargo molecules that are internalized from the cell surface.

Clathrin-mediated endocytosis, a rapid and highly dynamic process internalizes many proteins, including signaling receptors. The protocol described here directly visualizes the kinetics of individual endocytic events. This is essential for understanding how core members of the endocytic machinery coordinate with each other, and how protein cargo influence this process.

Many important signaling receptors are internalized through the well-studied process of clathrin-mediated endocytosis (CME). Traditional cell biological ...

Live Fluorescence, Inverse Imaging of Cell Ruffling, and Macropinocytosis

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase nutrients and other material in cells. The dramatic extension of cell surface ruffles, their closure to form macropinosomes, and the maturation of internalized macropinosomes are key events in this pathway that can be difficult to capture using conventional confocal imaging based on tracking a bolus of fluorescent cargo. Fluorescent dextrans are commonly used experimentally as fluid phase markers for macropinosomes and for other endocytic pathways. A method the lab has adopted to optimize the imaging of dextran uptake involves using live imaging of cells bathed in high concentrations of fluorescent dextran in the medium, with the unlabeled cells appearing in relief (as black). The cell ruffles are highlighted to visualize ruffle closure, and internalized macropinosomes appear as fluorescent vacuoles in the cell interior. This method is optimal for visualizing macropinosome features and allows for easy segmentation and quantification. This paper describes dual-labeling of pathways with different sized dextrans and the co-expression of lipid probes and fluorescent membrane proteins to demark macropinosomes and other endosomes. The detection of internalized dextran at an ultrastructural level using correlative light and electron microscopy (CLEM) is also demonstrated. These cell processes can be imaged using multiple live imaging modalities, including in 3D. Taken together, these approaches optimize macropinosome imaging for many different settings and experimental systems.

This protocol demonstrates dextran imaging in live cells using continuous uptake and inverse images to optimize visualization of ruffling, macropinosome maturation, and analysis of dextran and other cell labelings.

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase ...