Name: quantification of efferocytosis by single-cell fluorescence microscopy
Uploaded: 2018-08-18T14:00:00.000+00:00
Duration: 6 min 15 s
Description: Watch this Scientific Journal Video about Quantification of Efferocytosis by Single-cell Fluorescence Microscopy at JoVE.com

This study was funded by Canadian Institutes of Health Research (CIHR) Operating Grant MOP-123419, Natural Sciences and Engineering Research Council of Canada Discovery Grant 418194, and an Ontario Ministry of Research and Innovation Early Research Award to BH. DGW contributed some of the images presented, to the optimization of the protocols and to the writing of the manuscript; he was funded by a pump-priming grant from the university of Liverpool. CY is funded by a Vanier Graduate Scholarship and CIHR MD/PhD Studentship. The funding agencies had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

The authors declare no conflict of interest.

The methods outlined in this protocol enable the imaging and quantification of the dynamic efferocytic process, using both fixed-cell and live-cell approaches. These approaches offer several advantages over commonly employed flow cytometry-based methods23,24. The use of inside-out staining with fixed samples provides a more robust and accurate quantification of the rate and extent of efferocytosis &#8212; indeed, many flow cytometry-based methods simply label apoptotic cells and macrophages with different fluorophores, and score efferocytosis as the fraction of macrophages co-staining with the apoptotic cell marker, thus lacking the capacity to differentiate between bound versus internalized apoptotic cell material. Alternative flow cytometry approaches include those using pHrodo labeled apoptotic cells24. pHrodo is a pH-sensitive fluorophore that increases in brightness at acidic pH. While this fluorophore does provide better resolution between non-internalized versus internalized materials, as the fluorescence increases specifically following internalization of the apoptotic cell and acidification of the efferosome, the results can be confounded by disease processes which impair efferosome acidification34,35, and this method will miss efferosomes in the early (pre-acidification) stages of efferocytosis36, located in acidification-poor regions of the cell27, or in cells which weakly acidify their lysosomes37. A second advantage of the method described in this protocol is the use of live-cell imaging to measure the dynamics of efferocytosis, as processes such as probing, the formation of the efferocytic synapse, and the intracellular trafficking of efferosomes cannot be detected using flow cytometry-based approaches.
While this method offers many advantages, experiments requiring the quantification of many hundreds of cells &#8212; e.g. experiments detecting rare events, or quantifying highly variable processes &#8212; can be difficult, with analysis of these large datasets taking an inordinate amount of time. High-throughput imaging approaches such as imaging flow cytometry38 may allow for higher throughput than conventional microscopy, although in our experience current automated image analysis programs are not always capable of accurately segmenting non-internalized versus internalized materials. Specifically, the tight efferocytic synapse which forms between the apoptotic cell and efferocyte can exclude streptavidin staining, and as such a bound apoptotic cell often appears as a solid mass in the cell tracking dye channel, that is partially enveloped by streptavidin staining (Figure 2). Thus, while efferosomes are readily identified algorithmically, the bound portion of the apoptotic cell is often misidentified as an efferosome due to the difficulty of accounting for partial streptavidin staining. While we have yet to find a program that can accurately identify and quantify the piecemeal uptake of apoptotic cells without human assistance, we have found that trainable semi-automated systems39 can greatly accelerate analysis, reducing fraction efferocytosed measurements from 2&#8211;3 min to &#60;60 s per cell. Alternatively, non-digestible apoptotic cell mimics or cell types which minimally fragment upon apoptosis, can be used instead. This simplifies detection to single, larger structures, which may be more amenable to automated approaches and may eliminate the need for z-stacking. Even with these advances, the acquisition and analysis speed of this method remains limiting, and as such flow cytometry remains the most viable method when analyses of large cell numbers is required23.
Although we have described this method using cell-line and primary human macrophages as efferocytes and apoptotic Jurkat cells (a T cell line) as targets, this method can be applied to any efferocytic cell type or apoptotic cell target. Indeed, similar approaches have been used to investigate efferocytosis in hepatocytes and epithelial cells9,40,41, and the efferocytosis of clinically relevant targets such as tumor cells42. It may be necessary to modify this protocol when using non-immune efferocytes, or when modeling specific efferocytic events. For example, Jurkat cells are non-adherent and therefore are unlikely to fully recapitulate the mechanical forces and spatial limitations efferocytes encounter when interacting with adherent apoptotic cells or apoptotic cells within solid tissues. Many cell types will maintain adhesion during apoptosis and therefore can be used as adherent targets; as one example, HeLa cells reproducibly undergo staurosporine-induced apoptosis, phosphatidylserine scrambling, and blebbing over a 4&#8211;6 h period while maintaining adhesion of the cell body43. Cells suspended in a collagen matrix or stem cell-derived organoids44 may be potential models for studying efferocytosis in solid tissues, although we are unaware of any studies which have used these approaches. For some models immune cells such as Jurkat cells and neutrophils should not be used as apoptotic targets, as these cells can release cytokine-based &#34;find-me&#34; signals such as CX3CL1 which may be a confounding factor in models where inflammatory or migratory processes are investigated45. Thus, while the versatility of this assay allows for it to be used to explore efferocytosis across a range of cell types and model systems, care must be taken to select appropriate efferocytes, target cells, and culture conditions to best model the physiological process under investigation.
The analysis methods described in this protocol are intended only as a starting point, with the imaging-based nature of these experiments enabling a broad range of analyses. For example, efferosome positioning and diameter measurements can be collected when performing fraction efferocytosed measurements, or measured within live-cell time-courses, in order to investigate processes such as the intracellular trafficking of efferosomes and the rate of apoptotic cell degradation13,15. Immunostaining (Figure 4) can be used to investigate the recruitment of proteins to efferosomes, while live-cell imaging combined with morphological analyses can be used to quantify processes such binding efficacy and rates of engulfment30,46. We frequently perform these assays in macrophages expressing fluorescent transgenes that report signaling molecule activation or the activity of cellular events during efferocytosis (Figure 6). These reporters enable the monitoring of signaling processes during efferocytosis; for example, we have used fluorescently-tagged Rab GTPases to explore the role of Rab5, Rab7 and Rab17 in mediating efferosome processing and the subsequent trafficking of apoptotic cell-derived antigens13,15. Similarly, the incorporation of reporters of cell death, efferosome pH, reactive oxygen species production, and other cellular processes into live-cell experiments can be used to investigate the processes mediating the degradation of efferosomes31,47. A common challenge in these experiments is identifying transgenes or reporters with compatible fluorophores; often, we will eliminate the cell tracking dye and/or streptavidin to free channels for imaging other fluorophores. For some experiments it is beneficial to replace the apoptotic cell with a non-fragmenting mimic. This allows for quantification of signaling dynamics and cellular processes without the complexity of tracking the multiple efferosomes derived from a single apoptotic cell. See Evans et al. for instructions on preparing apoptotic cell mimics28.
The most common difficulty when designing these experiments is finding a combination of fluorophores that allow for the desired processes to be imaged, while minimizing photobleaching and phototoxicity29. Fluorophore selection is largely dictated by the lasers/excitation filters, dichroics/cubes and emission filters of the microscope, and therefore the choice of fluorophores is often specific to individual microscopes. Generally, longer wavelength fluorophores (orange to far-red, e.g. emission maxima &#62;580 nm) are less prone to photobleaching and these wavelengths are less disruptive to cells. Green fluorophores (emission maxima ~525 nm) can be used, but care needs to be taken to limit phototoxicity to the cells. Fluorophores that excite at wavelengths less than 480 nm (violet and blue) should be avoided due to their high phototoxicity and propensity for these excitation wavelengths to bleach other fluorophores29. Where possible, high-brightness and stable fluorophores should be selected. Similarly, the image acquisition parameters should be adjusted to minimize photobleaching and phototoxicity &#8212; e.g. longer exposures at low excitation intensity are preferred over higher excitation intensities48. The addition of antioxidants such as rutin and removal of some media components can improve both photostability and reduce phototoxicity49. Even with careful selection of fluorophores and imaging conditions, the need to limit photobleaching often requires the capture of images with low signal-to-noise ratios (see Figure 5 for an example). If quantifying fluorescence intensity, great care needs to be taken to limit image-processing artefacts; ideally, raw images without any form of processing should be analyzed. If deconvolving images prior to quantification, a deconvolution algorithm that preserves fluorescent intensities must be used50,51.
Live cell acquisitions can be particularly challenging, with successful experiments requiring a careful balance between excitation intensity, exposure time, frame rate, and experiment duration. Excitation intensity and exposure time should be adjusted to minimize phototoxicity as described above, with the caveat that longer exposure times may result in motion artefacts due to cell movement or limit the frame rate of the acquisition. The frame rate can vary depending on the experimental requirements. Lower frame-rates (5&#8211;30 min between frames) allow for imaging over prolonged periods of time (12 h or more) but provide minimal data on phenomena such as membrane dynamics and post-efferocytosis trafficking of efferosomes. High frame-rates (as fast as 1 frame per second) provide excellent temporal resolution of efferocytic events and efferosome trafficking &#8212; but even with careful selection of fluorophores, excitation intensity and exposure times &#8212; photobleaching or phototoxicity will usually limit these experiments to less than an hour in duration. In our experience, acquiring images every 1&#8211;2 min, over experimental periods of 2&#8211;6 h, are an acceptable compromise that provides quantifiable images of a sufficient number of efferocytic events, with reasonable temporal resolution.
Altered and failed efferocytosis is known to be involved in the pathology of cancer, atherosclerosis and multiple autoimmune disorders, with efferocytosis-targeting therapies showing great clinical promise7,52,53,54,55. Further development in these fields will require identification and characterization of the cellular processes, receptors and signaling pathways which regulate efferocytosis. The assay presented in this protocol represents a powerful tool for these studies and can be modified to quantify many of the cellular and signaling processes regulating efferocytosis.

Apoptosis, or programmed cell death, is a highly-regulated physiological process that occurs in most multicellular organisms and is crucial for their development and homeostasis1. In addition to being involved in normal cell turnover and embryonic development, apoptosis enables the elimination of infected or damaged cells from tissues and can be triggered in response to infection, inflammation, cancer, and also by medical interventions such as radiotherapy or steroids1. Apoptotic cells expose &#34;eat-me&#34; signals on their cell surface which are recognized by receptors on a range of professional and non-professional phagocytes, collectively referred to as &#34;efferocytes&#34;. Engagement of these receptors induces the uptake and degradation of the apoptotic cell by the efferocyte through a process known as efferocytosis2,3. Phosphatidylserine is the best characterized eat-me signal driving efferocytosis. It is normally confined to the inner leaflet of the plasma membrane, with apoptosis activating a lipid scramblase which disrupts this membrane asymmetry, thus exposing phosphatidylserine on the cell surface4. Phosphatidylserine is found on the extracellular surface of some non-apoptotic cells, such as mature macrophages and activated platelets. However, these cells are not efferocytosed due to the presence of &#34;don&#39;t eat me&#34; signals, such as CD47, on their cell surface5,6,7. Exposed phosphatidylserine is recognized by an array of efferocytic receptors expressed by efferocytes. Binding of these receptors to phosphatidylserine, either directly or through the aid of opsonins, activates signaling pathways that promote the engulfment of the apoptotic cell into a membrane-bound vacuole termed the efferosome8,9,10,11,12. The efferosome fuses sequentially with endosomes and lysosomes, which deliver the molecular machinery necessary to acidify the efferosome and to degrade the apoptotic cell cargo13,14. Once degraded, the apoptotic cell-derived materials are trafficked to the recycling endosome &#8212; a process which limits immune responses to apoptotic cell-derived antigens, and which may allow for recovery of nutrients from the apoptotic cell13,15. A failure in efferocytosis results in impaired clearance of apoptotic cells; these uncleared cells eventually undergo secondary necrosis. Necrotic cells release pro-inflammatory cytosolic contents, pathogens, and autoantigens into the extracellular milieu, thus driving a range of infective, inflammatory and autoimmune diseases16,17. Together, apoptosis and efferocytosis facilitate the removal of dying and dead cells and allow for the maintenance of tissue homeostasis.
Investigating the molecular mechanisms underlying efferocytosis requires methods that provide a clear quantification of apoptotic cell uptake. This quantification is complicated by the fact that unlike other uptake mechanisms such as endocytosis and phagocytosis18,19, efferocytosis may not result in the engulfment of intact target cell, resulting in the piecemeal uptake of the apoptotic cell by the efferocyte20. The protocol described herein describes an in vitro efferocytosis assay that provides accurate delineation of the internalized versus non-internalized portions of individual apoptotic cells and can be combined with a variety of fixed-cell and live-cell microscopy approaches. Traditional phagocytosis assays add antibodies specific to the phagocytic target at the end of the experiment in order to label non-internalized targets, where as our method differs by labelling the apoptotic target with covalently-linked biotin21,22. While apoptotic cell specific antibodies can be used in this assay, the biotinylation approach allows for any protein-bearing target to be labeled and avoids potential issues with secondary antibody cross-reactivity if immunostaining is performed. Specifically, we outline the preparation of apoptotic Jurkat cells that have been dual-stained with both a cell tracking dye and biotin. The cell tracking dye allows for apoptotic cell-derived materials to be tracked during efferocytosis, whereas surface biotinylation allows for the discrimination of internalized from non-internalized portions of efferocytosed apoptotic cells. We also describe the culture and preparation of J774.2 and THP-1 cell lines for use as murine and human efferocytes, monocyte-derived M2 macrophages as an example of primary cell efferocytosis, and Jurkat cells for use as efferocytic targets. These methods can easily be applied to other cell lines or primary cells, to target cells undergoing any form of cell death (e.g. apoptosis, necrosis and necroptosis), and to micron-sized mimics which simulate apoptotic cells through lipid coatings or coating with ligands specific to an efferocytic receptor of interest.
The method outlined in this protocol has several advantages over the flow cytometry based methods commonly used in the field23,24. By directly imaging the phagocyte-apoptotic cell interaction, combined with clear labeling of both total and non-internalized apoptotic cell material, quantitative measures of efferocytosis can be made. Moreover, the use of pH-insensitive fluorophores limits confounding factors such as the suppression of FITC and GFP fluorescence at lysosomal pH that confounds some alternative methods25. Lastly, while not described in detail, these methods can be employed using efferocytes expressing fluorescently-labeled transgenes, or with post-fixation immunostaining, to allow for quantification of signaling molecule activity and monitoring of the cellular processes during efferocytosis.

<table><tbody><tr><td>RPMI 1640 Media</td><td>Wisent</td><td>3500-000-EL</td><td></td></tr><tr><td>DMEM Media</td><td>Wisent</td><td>319-005-CL</td><td></td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Wisent</td><td>080-150</td><td></td></tr><tr><td>PBS</td><td>Wisent</td><td>311-010-CL</td><td></td></tr><tr><td>18 mm circular glass coverslips #1.5 thickness</td><td>Electron Microscopy Sciences</td><td>72290-08</td><td>Size and shape of coverslip is not critical, but 18 mm fit into the wells of a standard 12-well plate which simplifies cell culture</td></tr><tr><td>Staurosporine</td><td>Cayman Chemical</td><td>81590</td><td>Dissolve in DMSO at 1 mM (1,000x stock solution)</td></tr><tr><td>Annexin V-Alexa 488</td><td>ThermoFisher</td><td>R37174</td><td></td></tr><tr><td>EZ-Link NHS-Biotin</td><td>ThermoFisher</td><td>20217</td><td>Store in a dessicator. Do not prepare a stock solution.</td></tr><tr><td>DMSO</td><td>Sigma-Aldrich</td><td>D2650</td><td></td></tr><tr><td>CellTrace FarRed</td><td>ThermoFisher</td><td>C34572</td><td></td></tr><tr><td>CellTrace Orange</td><td>ThermoFisher</td><td>C34851</td><td></td></tr><tr><td>Hoescht 33342</td><td>ThermoFisher</td><td>62249</td><td></td></tr><tr><td>FITC-Streptavadin</td><td>ThermoFisher</td><td>SA1001</td><td></td></tr><tr><td>Lympholyte-poly cell sepration medium</td><td>Cedarlane Labs</td><td>CL5071</td><td></td></tr><tr><td>Recombinant Human M-CSF</td><td>Peprotech</td><td>200-04</td><td></td></tr><tr><td>Recombinant Human IL-4</td><td>Peprotech</td><td>300-25</td><td></td></tr><tr><td>J774.2 Macrophage Cell Line</td><td>Sigma-Aldrich</td><td>85011428-1VL</td><td></td></tr><tr><td>THP-1 Human Monocyte Cell Line</td><td>ATCC</td><td>TIB-202</td><td></td></tr><tr><td>Jurkat T Cell Line</td><td>ATCC</td><td>TIB-152</td><td></td></tr></tbody></table>

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.

Overnight culture of Jurkat cells with 1 &#181;M staurosporine results in apoptosis of &#62;95% of cells, which can be confirmed with Annexin V staining (Figure 1). Other cell types can be used for these experiments, although the concentration of staurosporine and the duration of staurosporine treatment will need to be optimized for each cell line. For reliable detection and quantification of efferocytosis, &#62;80% of cells should be apoptotic prior to adding them to the efferocytes. Other inducers of apoptosis (e.g. heat-shock, etoposide and UV-light) can also be used, but in our experience, these produce a more heterogeneous induction of apoptosis and result in mixed cell populations containing apoptotic, secondary necrotic and non-apoptotic target cells.
For fixed-cell imaging with inside-out staining, closely apposed efferocyte-apoptotic cell interactions should be observed at all time points, with clearly delineated non-efferocytosed (streptavidin+/cell tracking dye+) and efferocytosed (streptavidin-/cell tracking+) materials visible (Figure 2). It is important to note that the synapse that forms between the efferocyte and the apoptotic cell is often tight enough to exclude streptavidin, and thus any cell tracking dye stained object bearing streptavidin at any point on its circumference should be considered a bound cell and not an efferosome. In most experiments the fraction of apoptotic cells that are efferocytosed will increase in a time-dependent fashion, either until no non-internalized apoptotic cell materials remain, or until the phagocyte reaches its maximum efferocytic capacity (Figure 3). Analysis is typically performed and recorded for individual cells (Figure 3A), however, data can be averaged across cells within individual experiments and the average values from multiple experimental repeats analyzed with conventional statistical approaches (Figure 3B). Modifications of this standardized protocol can be made to allow for use of primary cell types, alternative efferocytic targets, and for additional staining to be included. For example, Figure 4 shows a primary M2-polarized human macrophage that has efferocytosed apoptotic cell mimics comprised of 3 &#181;m diameter silica beads coated in a mixture of phosphatidylserine and phosphatidylcholine28, followed by subsequent fixation, permeabilization and immunostaining for the recycling endosome marker Rab17.
During live cell imaging efferocytic professional phagocytes will be observed to extend and retract small processes in a process termed &#34;probing&#34;30 (Figure 5, t = 0); when these processes encounter a target they firmly adhere to the target and draw it to the phagocyte. This probing activity may not be observed with non-professional phagocytes such as epithelial cells. The efferocyte will then form a tight &#34;efferocytic synapse&#34;32 between itself and the apoptotic cell, with this synapse often enveloping a large portion of the apoptotic cell (Figure 5, t = 10). The efferocyte will then draw pieces of the apoptotic cell from this synapse into its cytosol (Figure 5, 10&#8211;30 min). Soon after their formation, these nascent efferosomes are trafficked away from the synapse and towards the peri-nuclear area, a result of Rab7/RILP/dynein-dynactin mediated transport33. Over time the efferosomes will shrink and the resulting degraded materials redistributed throughout the cell, where they are likely absorbed or recycled13,15. The ability to detect these processes is highly dependent on the acquisition frame-rate and duration of the experiment. Rapid processes such as probing may not be observed at lower frame-rates, while slow events (efferosome trafficking and absorption) require longer imaging periods &#8212; in both cases, these large image acquisitions often require the capture of lower-intensity images to limit photobleaching and phototoxicity (Figure 5). As with the fixed-cell assay, the live-cell version of this assay can be modified to suit the needs of the investigator. Figure 6 shows a single frame of a live cell acquisition of J774.2 macrophages expressing transgenes which fluorescently demark the plasma membrane (PM-GFP) and which selectively binds the signaling lipid PI(3)P (FYVE-RFP). Generation of PI(3)P on the efferosome membrane can be detected as co-localization of FYVE-RFP with the PM-GFP+ efferosome membrane. By quantifying the intensity of FYVE-RFP on the efferosome, the dynamics of PI(3)P signaling on the efferosome can be quantified (Figure 6).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58149/58149fig1.jpg" /> 
Figure 1: Annexin V Staining of Apoptotic and Non-Apoptotic Jurkat Cells. Annexin V labels the phosphatidylserine &#34;eat-me&#34; signal exposed by apoptotic cells. (Top) Untreated (UT) Jurkat cells display a healthy (smooth and rounded) morphology (DIC) and do not stain with Annexin V. (Bottom) Jurkat cells cultured overnight with 1 &#181;M staurosporine (STS) take on an apoptotic morphology (irregular and blebbed) and stain with Annexin V. Scale bar = 10 &#181;m, image intensity is displayed as a color map. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58149/58149fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58149/58149fig2.jpg" /> 
Figure 2: Early and Late Engulfment of Apoptotic Jurkat Cells by J774.2 Macrophages. Images are of macrophages at an early (60 min) and late (120 min) stage of efferocytosis. White-light (DIC) image illustrates the tight interface between the macrophage (m&#966;) and apoptotic cell (AC). Cell tracking dye (CTD, red) reveals the location of all apoptotic cell-derived materials. FITC-Streptavidin staining (Str, green) identifies the portion of the apoptotic cell that has not yet been engulfed by the macrophage. Macrophage nuclei were pre-stained with DAPI (blue). Efferosomes (red) versus the non-internalized portion of the apoptotic cell (green/yellow) are readily identified in an overlay of the Streptavidin and cell tracking dye images. Note the absence of streptavidin staining at the macrophage-apoptotic cell interface at the early timepoint, created by the exclusion of streptavidin from the tightly formed efferocytic synapse (*). The macrophage nucleus is identified by Hoechst staining (blue). Scale bar = 10 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58149/58149fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58149/58149fig3.jpg" /> 
Figure 3: Time-Course of the Fraction of Apoptotic Cell Efferocytosed by J774.2 Macrophages. Efferocytosis assays were performed for the indicated times, and the fraction of engulfed apoptotic cell material determined at each time point using cell tracking dye/Streptavadin staining. (A) Fraction of efferocytosed materials within individual macrophages, data is presented from individual cells, horizontal bar indicates the mean. 12&#8211;17 cells per condition, data from 1 of 5 independent experiments. (B) Fraction of efferocytosed materials, averaged across 5 independent experiments, at least 10 cells/condition/repeat. * p &#60;0.05 compared to 30 min, Kruskal-Wallis test with Dunn correction. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58149/58149fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58149/58149fig4.jpg" /> 
Figure 4: Analysis of Rab17 Recruitment to Efferosomes in Primary Human Macrophages. A M2-polarized macrophage engulfed apoptotic cell mimics (arrows) and was immunostained for Rab17 (green) 60 min following engulfment. Scale bar = 5 &#181;m. Cell nucleus is stained with Hoeschst, DIC image shows cell morphology and is used to identify apoptotic cell mimics. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58149/58149fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58149/58149fig5.jpg" /> 
Figure 5: Live cell efferocytosis assay. A cell tracking dye labeledJ774.2 macrophage (m&#966;, green) was recorded as it engulfed an apoptotic Jurkat cell labeled with a different color cell tracking dye (AC, red). The apoptotic cell is broken into multiple fragments during the internalization process, resulting in piecemeal uptake and formation of multiple efferosomes (arrows). Scale bar = 10 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58149/58149fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58149/58149fig6.jpg" /> 
Figure 6: Using Fluorescent Transgenes to Investigate Efferocytic Signaling. A J774.2 macrophage was transfected with a GFP-tagged plasma membrane marker (PM, Green) and FYVE-RFP construct, which binds the signaling lipid PI(3)P (Red). Efferocytosis forms a nascent plasma membrane-derived efferosome that contains PI(3)P, as detected by FYVE-RFP recruitment (arrow). The dynamics of PI(3)P signaling was quantified as fold intensity relative to the FYVE-RFP signal present at the point of efferosome closure (t = 0, graph). This experiment used an apoptotic cell-mimicking bead (arrow, DIC) rather than apoptotic cells. Scale bar = 5 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58149/58149fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Quantification of Efferocytosis by Single-cell Fluorescence Microscopy at JoVE.com

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

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 ...

quantification-of-efferocytosis-by-single-cell-fluorescence-microscopy

Universidad San Sebastian

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JoVE Journal

Immunology and Infection

12.4K Views.  University of Western Ontario. 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.

Video: Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

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 ...

Preparation of Bead-supported Lipid Bilayers to Study the Particulate Output of T Cell Immune Synapses

Antigen-presenting cells (APCs) present three activating signals to T cells engaged in physical contact: 1) antigen, 2) costimulation/corepression, and 3) soluble cytokines. T cells release two kinds of effector particles in response to activation: trans-synaptic vesicles (tSVs) and supramolecular attack particles, which transfer intercellular messengers and mediate cytotoxicity, respectively. These entities are quickly internalized by APCs engaged in physical contact with T cells, making their characterization daunting. This paper presents the protocol to fabricate and use Bead-Supported Lipid Bilayers (BSLBs) as antigen-presenting cell (APC) mimetics to capture and analyze these trans-synaptic particles. Also described are the protocols for the absolute measurements of protein densities on cell surfaces, the reconstitution of BSLBs with such physiological levels, and the flow cytometry procedure for tracking synaptic particle release by T cells. This protocol can be adapted to study the effects of individual proteins, complex ligand mixtures, pathogen virulence determinants, and drugs on the effector output of T cells, including helper T cells, cytotoxic T lymphocytes, regulatory T cells, and chimeric antigen receptor-expressing T cells (CART).

Here we present the protocol for the stepwise reconstitution of synthetic antigen-presenting cells using Bead-Supported Lipid Bilayers and their use to interrogate the synaptic output from activated T cells.

Antigen-presenting cells (APCs) present three activating signals to T cells engaged in physical contact: 1) antigen, 2) costimulation/corepression, and 3) ...

Identification of a Murine Erythroblast Subpopulation  Enriched in Enucleating Events by Multi-spectral Imaging Flow Cytometry

Erythropoiesis in mammals concludes with the dramatic process of enucleation that results in reticulocyte formation. The mechanism of enucleation has not yet been fully elucidated. A common problem encountered when studying the localization of key proteins and structures within enucleating erythroblasts by microscopy is the difficulty to observe a sufficient number of cells undergoing enucleation.&#160;We have developed a novel analysis protocol using multiparameter high-speed cell imaging in flow (Multi-Spectral Imaging Flow Cytometry), a method that combines immunofluorescent microscopy with flow cytometry, in order to identify efficiently a significant number of enucleating events, that allows to obtain measurements and perform statistical analysis.
We first describe here two in vitro erythropoiesis culture methods used in order to synchronize murine erythroblasts and increase the probability of capturing enucleation at the time of evaluation.&#160;Then, we describe in detail the staining of erythroblasts after fixation and permeabilization in order to study the localization of intracellular proteins or lipid rafts during enucleation by multi-spectral imaging flow cytometry.&#160;Along with size and DNA/Ter119 staining which are used to identify the orthochromatic erythroblasts, we utilize the parameters &#8220;aspect ratio&#8221; of a cell in the bright-field channel that aids in the recognition of elongated cells and &#8220;delta centroid XY Ter119/Draq5&#8221; that allows the identification of cellular events in which the center of Ter119 staining (nascent reticulocyte) is far apart from the center of Draq5 staining (nucleus undergoing extrusion), thus indicating a cell about to enucleate.&#160;The subset of the orthochromatic erythroblast population with high delta centroid and low aspect ratio is highly enriched in enucleating cells.

The present protocol describes a novel method of identifying a population of enucleating orthochromatic erythroblasts by multi-spectral imaging flow cytometry, providing a visualization of the erythroblast enucleation process.

Erythropoiesis in mammals concludes with the dramatic process of enucleation that results in reticulocyte formation. The mechanism of enucleation has not ...

Biology

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal remodeling and vesicular trafficking in living cells. Quantitative methodologies using chimeric GFP fusions have been developed for many applications; however, GFP is somewhat resistant to proteolysis, thus its fluorescence persists in the lysosome/vacuole, which can impede quantification of cargo trafficking in the endocytic pathway. An alternative method for quantifying endocytosis and post-endocytic trafficking events makes use of superecliptic pHluorin, a pH-sensitive variant of GFP that is quenched in acidic environments. Chimeric fusion of pHluorin to the cytoplasmic tail of transmembrane cargo proteins results in a dampening of fluorescence upon incorporation of the cargo into multivesicular bodies (MVBs) and delivery to the lysosome/vacuole lumen. Thus, quenching of vacuolar fluorescence facilitates quantification of endocytosis and early events in the endocytic pathway. This paper describes methods using pHluorin-tagged cargos for quantification of endocytosis via fluorescence microscopy, as well as population-based assays using flow cytometry.

Accurate quantification of vesicular trafficking events often provides key insights into roles for specific proteins and the effects of mutations. This paper presents methods for using superecliptic pHluorin, a pH-sensitive GFP variant, as a tool for quantification of endocytic events in living cells using quantitative fluorescence microscopy and flow cytometry.

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal ...

Flow Cytometry-Based Assays for Microglial Engulfment of Synaptic Materials

Quantification of Microglial Engulfment of Synaptic Material Using Flow Cytometry

Microglia play a pivotal role in synaptic refinement in the brain. Analysis of microglial engulfment of synapses is essential for comprehending this process; however, currently available methods for identifying microglial engulfment of synapses, such as immunohistochemistry (IHC) and imaging, are laborious and time-intensive. To address this challenge, herein we present in vitro and in vivo* assays that allow fast and high-throughput quantification of microglial engulfment of synapses using flow cytometry.
In the in vivo* approach, we performed intracellular vGLUT1 staining following fresh cell isolation from adult mouse brains to quantify engulfment of vGLUT1+ synapses by microglia. In the in vitro synaptosome engulfment assay, we used freshly isolated cells from the adult mouse brain to quantify the engulfment of pHrodo&#160;Red-labeled synaptosomes by microglia. These protocols together provide a time-efficient approach to quantifying microglial engulfment of synapses and represent promising alternatives to labor-intensive image analysis-based methods. By streamlining the analysis, these assays can contribute to a better understanding of the role of microglia in synaptic refinement in different disease models.

Author Spotlight: Advancing Understanding Through Technological Innovations in Psychoneuroimmunology

Here we present two protocols to quantify microglial engulfment of vGLUT1-positive synapses and pHRodo&#160;Red-labeled crude synaptosomes using flow cytometry.

Microglia play a pivotal role in synaptic refinement in the brain. Analysis of microglial engulfment of synapses is essential for comprehending this ...

Neuroscience

Extracellular Vesicle Uptake Assay via Confocal Microscope Imaging Analysis

There is a need for practical assays to visualize and quantify the cells' extracellular vesicle (EV) uptake. EV uptake plays a role in intercellular communication in various research fields; cancer biology, neuroscience, and drug delivery. Many EV uptake assays have been reported in the literature; however, there is a lack of practical, detailed experimental methodology. EV uptake can be assessed by fluorescently labeling EVs to detect their location within cells. Distinguishing between internalized EVs in cells and the superficial EVs on cells is difficult, yet critical, to accurately determine the EV uptake. Therefore, an assay that efficiently quantifies EV uptake through three-dimensional (3D) fluorescence confocal microscopy is proposed in this work. Fluorescently labeled EVs were prepared using a nano-filtration-based microfluidic device, visualized by 3D confocal microscopy, and then analyzed through advanced image-processing software. The protocol provides a robust methodology for analyzing EVs on a cellular level and a practical approach for efficient analysis.

Extracellular Vesicle Uptake Assay via Confocal Microscope Imaging Analysis

Extracellular vesicles (EVs) contribute to cellular biology and intercellular communications. There is a need for practical assays to visualize and quantify EVs uptake by the cells. The current protocol proposes the EV uptake assay by utilizing three-dimensional fluorescence imaging via confocal microscopy, following EV isolation by a nano-filtration-based microfluidic device.

There is a need for practical assays to visualize and quantify the cells' extracellular vesicle (EV) uptake. EV uptake plays a role in intercellular ...

Single Extracellular Vesicle Transmembrane Protein Characterization by Nano-Flow Cytometry

Single particle characterization has become increasingly relevant for research into extracellular vesicles, progressing from bulk analysis techniques and first-generation particle analysis to comprehensive multi-parameter measurements such as nano-flow cytometry (nFCM). nFCM is a form of flow cytometry that utilizes instrumentation specifically designed for nano-particle analysis, allowing for thousands of EVs to be characterized per minute both with and without the use of staining techniques. High resolution side scatter (SS) detection allows for size and concentration to be determined for all biological particles larger than 45 nm, while simultaneous fluorescence (FL) detection identifies the presence of labeled markers and targets of interest. Labeled subpopulations can then be described in quantitative units of particles/mL or as a percentage of the total particles identified by side scatter.
Here, EVs derived from conditioned cell culture media (CCM) are labeled with both a lipid dye, to identify particles with a membrane, and antibodies specific for CD9, CD63, and CD81 as common EV markers. Measurements of comparison material, a concentration standard and a size standard of silica nanospheres, as well as labeled sample material are analyzed in a 1-minute analysis. The software is then used to measure the concentration and size distribution profile of all particles, independent of labeling, before determining the particles that are positive for each of the labels.
Simultaneous SS and FL detection can be utilized flexibly with many different EV sources and labeling targets, both external and internal, describing EV samples in a comprehensive and quantitative manner.

The latest generation of EV characterization tools are capable of single EV analysis across multiple parameters simultaneously. Nano-flow cytometry measures all biological particles larger than 45 nm without labeling and identifies specific characteristics of subpopulations by a variety of fluorescent labeling techniques.

Single particle characterization has become increasingly relevant for research into extracellular vesicles, progressing from bulk analysis techniques and ...

Fixed Cell Efferocytosis Assay: A Method to Study Efferocytic Uptake of Apoptotic Cells by Macrophages Using Fluorescence Microscopy

Source: Taruc, K., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/58149">Quantification of Efferocytosis by Single-cell Fluorescence Microscopy.</a> J. Vis. Exp. (2018).
In this video, we demonstrate the efferocytosis assay to study the clearance of apoptotic cells using confocal microscopy. Efferocytosis is a multi-step process where apoptotic cells are engulfed by phagocytes and degraded within vesicles called efferosomes. This protocol provides a method to visualize internalized and non-internalized portions of individual apoptotic cells and quantify efferocytosis.

Source: Taruc, K., et al. Quantification of Efferocytosis by Single-cell Fluorescence Microscopy. J. Vis. Exp. (2018).
In this video, we demonstrate the ...

JoVE Encyclopedia of Experiments

Biological Techniques

Assay Techniques

Cell-based assays

Utilizing Bead-Supported Lipid Bilayers to Investigate the Synaptic Output from T Cells

Source: C&#233;spedes, P. F., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/63130">Preparation of Bead-supported Lipid Bilayers to Study the Particulate Output of T Cell Immune Synapses.</a> J. Vis. Exp. (2022)
This video illustrates the reconstitution of synthetic antigen-presenting cells through Bead-Supported Lipid Bilayers and their application in assessing the synaptic output generated by activated T cells.

1. Protein density calibrations on BSLBs (Bead-supported lipid bilayers)

	Before taking the 5.00 &#177;0.05 &#181;m diameter non-functionalized silica ...

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Summary

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Chapters in this video

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

Preparation of Bead-supported Lipid Bilayers to Study the Particulate Output of T Cell Immune Synapses

Identification of a Murine Erythroblast Subpopulation Enriched in Enucleating Events by Multi-spectral Imaging Flow Cytometry

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast

Quantification of Microglial Engulfment of Synaptic Material Using Flow Cytometry

Extracellular Vesicle Uptake Assay via Confocal Microscope Imaging Analysis

Single Extracellular Vesicle Transmembrane Protein Characterization by Nano-Flow Cytometry

Fixed Cell Efferocytosis Assay: A Method to Study Efferocytic Uptake of Apoptotic Cells by Macrophages Using Fluorescence Microscopy

Utilizing Bead-Supported Lipid Bilayers to Investigate the Synaptic Output from T Cells