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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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

Nanyang Technological University

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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

Microwave Assisted Rapid Diagnosis of Plant Virus Diseases by Transmission Electron Microscopy

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample preparation for transmission electron microscopy (TEM) such investigations can take several days1,2 and are therefore not suited for a rapid diagnosis of plant virus diseases. Microwave fixation can be used to drastically reduce sample preparation time for TEM investigations with similar ultrastructural results as observed after conventionally sample preparation3-5. Many different custom made microwave devices are currently available which can be used for the successful fixation and embedding of biological samples for TEM investigations5-8. In this study we demonstrate on Tobacco Mosaic Virus (TMV) infected Nicotiana tabacum plants that it is possible to diagnose ultrastructural alterations in leaves in about half a day by using microwave assisted sample preparation for TEM. We have chosen to perform this study with a commercially available microwave device as it performs sample preparation almost fully automatically5 in contrast to the other available devices where many steps still have to be performed manually6-8 and are therefore more time and labor consuming. As sample preparation is performed fully automatically negative staining of viral particles in the sap of the remaining TMV-infected leaves and the following examination of ultrastructure and size can be performed during fixation and embedding.

This study describes a method that allows the rapid and clear diagnosis of plant virus diseases in about half a day by using a combination of microwave assisted plant sample preparation for transmission electron microscopy and negative staining methods.

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample ...

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly constituted by proteins and exopolysaccharides, among other factors 7-10. The microbial community encased within the biofilm often shows the differentiation of distinct subpopulation of specialized cells 11-17. These subpopulations coexist and often show spatial and temporal organization within the biofilm 18-21.
Biofilm formation in the model organism Bacillus subtilis requires the differentiation of distinct subpopulations of specialized cells. Among them, the subpopulation of matrix producers, responsible to produce and secrete the extracellular matrix of the biofilm is essential for biofilm formation 11,19. Hence, differentiation of matrix producers is a hallmark of biofilm formation in B. subtilis.
We have used fluorescent reporters to visualize and quantify the subpopulation of matrix producers in biofilms of B. subtilis 15,19,22-24. Concretely, we have observed that the subpopulation of matrix producers differentiates in response to the presence of self-produced extracellular signal surfactin 25. Interestingly, surfactin is produced by a subpopulation of specialized cells different from the subpopulation of matrix producers 15.
We have detailed in this report the technical approach necessary to visualize and quantify the subpopulation of matrix producers and surfactin producers within the biofilms of B. subtilis. To do this, fluorescent reporters of genes required for matrix production and surfactin production are inserted into the chromosome of B. subtilis. Reporters are expressed only in a subpopulation of specialized cells. Then, the subpopulations can be monitored using fluorescence microscopy and flow cytometry (See Fig 1).
The fact that different subpopulations of specialized cells coexist within multicellular communities of bacteria gives us a different perspective about the regulation of gene expression in prokaryotes. This protocol addresses this phenomenon experimentally and it can be easily adapted to any other working model, to elucidate the molecular mechanisms underlying phenotypic heterogeneity within a microbial community.

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Microbial biofilms are generally constituted by distinct subpopulations of specialized cells. Single-cell analysis of these subpopulations requires the use of fluorescent reporters. Here we describe a protocol to visualize and monitor several subpopulationswithin B. subtilis biofilms using fluorescence microscopy and flow cytometry.

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly ...

Visualization of Bacterial Toxin Induced Responses Using Live Cell Fluorescence Microscopy

Bacterial toxins bind to cholesterol in membranes, forming pores that allow for leakage of cellular contents and influx of materials from the external environment. The cell can either recover from this insult, which requires active membrane repair processes, or else die depending on the amount of toxin exposure and cell type1. In addition, these toxins induce strong inflammatory responses in infected hosts through activation of immune cells, including macrophages, which produce an array of pro-inflammatory cytokines2. Many Gram positive bacteria produce cholesterol binding toxins which have been shown to contribute to their virulence through largely uncharacterized mechanisms.
Morphologic changes in the plasma membrane of cells exposed to these toxins include their sequestration into cholesterol-enriched surface protrusions, which can be shed into the extracellular space, suggesting an intrinsic cellular defense mechanism3,4. This process occurs on all cells in the absence of metabolic activity, and can be visualized using EM after chemical fixation4. In immune cells such as macrophages that mediate inflammation in response to toxin exposure, induced membrane vesicles are suggested to contain cytokines of the IL-1 family and may be responsible both for shedding toxin and disseminating these pro-inflammatory cytokines5,6,7. A link between IL-1&beta; release and a specific type of cell death, termed pyroptosis has been suggested, as both are caspase-1 dependent processes8. To sort out the complexities of this macrophage response, which includes toxin binding, shedding of membrane vesicles, cytokine release, and potentially cell death, we have developed labeling techniques and fluorescence microscopy methods that allow for real time visualization of toxin-cell interactions, including measurements of dysfunction and death (Figure 1). Use of live cell imaging is necessary due to limitations in other techniques. Biochemical approaches cannot resolve effects occurring in individual cells, while flow cytometry does not offer high resolution, real-time visualization of individual cells. The methods described here can be applied to kinetic analysis of responses induced by other stimuli involving complex phenotypic changes in cells.

Methods for purifying the cholesterol binding toxin streptolysin O from recombinant E. coli and visualization of toxin binding to live eukaryotic cells are described. Localized delivery of toxin induces rapid and complex changes in targeted cells revealing novel aspects of toxin biology.

Bacterial toxins bind to cholesterol in membranes, forming pores that allow for leakage of cellular contents and influx of materials from the external ...

Single Cell Measurements of Vacuolar Rupture Caused by Intracellular Pathogens

Shigella flexneri are pathogenic bacteria that invade host cells entering into an endocytic vacuole. Subsequently, the rupture of this membrane-enclosed compartment allows bacteria to move within the cytosol, proliferate and further invade neighboring cells. Mycobacterium tuberculosis is phagocytosed by immune cells, and has recently been shown to rupture phagosomal membrane in macrophages. We developed a robust assay for tracking phagosomal membrane disruption after host cell entry of Shigella flexneri or Mycobacterium tuberculosis. The approach makes use of CCF4, a FRET reporter sensitive to &beta;-lactamase that equilibrates in the cytosol of host cells. Upon invasion of host cells by bacterial pathogens, the probe remains intact as long as the bacteria reside in membrane-enclosed compartments. After disruption of the vacuole, &beta;-lactamase activity on the surface of the intracellular pathogen cleaves CCF4 instantly leading to a loss of FRET signal and switching its emission spectrum. This robust ratiometric assay yields accurate information about the timing of vacuolar rupture induced by the invading bacteria, and it can be coupled to automated microscopy and image processing by specialized algorithms for the detection of the emission signals of the FRET donor and acceptor. Further, it allows investigating the dynamics of vacuolar disruption elicited by intracellular bacteria in real time in single cells. Finally, it is perfectly suited for high-throughput analysis with a spatio-temporal resolution exceeding previous methods. Here, we provide the experimental details of exemplary protocols for the CCF4 vacuolar rupture assay on HeLa cells and THP-1 macrophages for time-lapse experiments or end points experiments using Shigella flexneri as well as multiple mycobacterial strains such as Mycobacterium marinum, Mycobacterium bovis, and Mycobacterium tuberculosis.

We describe a method for tracking the endomembrane rupture elicited by the intracellular bacteria Shigella flexneri and Mycobacterium tuberculosis upon host cell invasion. Our assay makes use of CCF4, a host cytoplasmic FRET probe in live or fixed cells. This reporter is degraded by an enzyme activity present on the bacterial surface.

Shigella flexneri are pathogenic bacteria that invade host cells entering into an endocytic vacuole. Subsequently, the rupture of this membrane-enclosed ...

Fluorescence Microscopy Methods for Determining the Viability of Bacteria in Association with Mammalian Cells

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols to address these questions include colony count assays, gentamicin protection assays, and electron microscopy. Colony count and gentamicin protection assays only assess the viability of the entire bacterial population and are unable to determine individual bacterial viability. Electron microscopy can be used to determine the viability of individual bacteria and provide information regarding their localization in host cells. However, bacteria often display a range of electron densities, making assessment of viability difficult. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells. These assays were developed originally to assess survival of Neisseria gonorrhoeae in primary human neutrophils, but should be applicable to any bacterium-host cell interaction. These protocols combine membrane-permeable fluorescent dyes (SYTO9 and 4&#39;,6-diamidino-2-phenylindole [DAPI]), which stain all bacteria, with membrane-impermeable fluorescent dyes (propidium iodide and SYTOX Green), which are only accessible to nonviable bacteria. Prior to eukaryotic cell permeabilization, an antibody or fluorescent reagent is added to identify extracellular bacteria. Thus these assays discriminate the viability of bacteria adherent to and inside eukaryotic cells. A protocol is also provided for using the viability dyes in combination with fluorescent antibodies to eukaryotic cell markers, in order to determine the subcellular localization of individual bacteria. The bacterial viability dyes discussed in this article are a sensitive complement and/or alternative to traditional microbiology techniques to evaluate the viability of individual bacteria and provide information regarding where bacteria survive in host cells.

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells.

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols ...

Assessing Anti-fungal Activity of Isolated Alveolar Macrophages by Confocal Microscopy

The lung is an interface where host cells are routinely exposed to microbes and microbial products. Alveolar macrophages are the first-line phagocytic cells that encounter inhaled fungi and other microbes. Macrophages and other immune cells recognize Aspergillus motifs by pathogen recognition receptors and initiate downstream inflammatory responses. The phagocyte NADPH oxidase generates reactive oxygen intermediates (ROIs) and is critical for host defense. Although NADPH oxidase is critical for neutrophil-mediated host defense1-3, the importance of NADPH oxidase in macrophages is not well defined. The goal of this study was to delineate the specific role of NADPH oxidase in macrophages in mediating host defense against A. fumigatus. We found that NADPH oxidase in alveolar macrophages controls the growth of phagocytosed A. fumigatus spores4. Here, we describe a method for assessing the ability of mouse alveolar macrophages (AMs) to control the growth of phagocytosed Aspergillus spores (conidia). Alveolar macrophages are stained in vivo and ten days later isolated from mice by bronchoalveolar lavage (BAL). Macrophages are plated onto glass coverslips, then seeded with green fluorescent protein (GFP)-expressing A. fumigatus spores. At specified times, cells are fixed and the number of intact macrophages with phagocytosed spores is assessed by confocal microscopy.

A method to evaluate the ability of isolated mouse alveolar macrophages to control the growth of phagocytosed Aspergillus spores by confocal microscopy.

The lung is an interface where host cells are routinely exposed to microbes and microbial products. Alveolar macrophages are the first-line phagocytic ...

Quantification of Cytosolic vs. Vacuolar Salmonella in Primary Macrophages by Differential Permeabilization

Intracellular bacterial pathogens can replicate in the cytosol or in specialized pathogen-containing vacuoles (PCVs). To reach the cytosol, bacteria like Shigella flexneri and Francisella novicida need to induce the rupture of the phagosome. In contrast, Salmonella typhimurium replicates in a vacuolar compartment, known as Salmonella-containing vacuole (SCV). However certain mutants of Salmonella fail to maintain SCV integrity and are thus released into the cytosol. The percentage of cytosolic vs. vacuolar bacteria on the level of single bacteria can be measured by differential permeabilization, also known as phagosome-protection assay. The approach makes use of the property of detergent digitonin to selectively bind cholesterol. Since the plasma membrane contains more cholesterol than other cellular membranes, digitonin can be used to selectively permeabilize the plasma membrane while leaving intracellular membranes intact. In brief, following infection with the pathogen expressing a fluorescent marker protein (e.g. mCherry among others), the plasma membrane of host cells is permeabilized with a short incubation in digitonin containing buffer. Cells are then washed and incubated with a primary antibody (coupled to a fluorophore of choice) directed against the bacterium of choice (e.g. anti-Salmonella-FITC) and washed again. If unmarked bacteria are used, an additional step can be done, in which all membranes are permeabilized and all bacteria stained with a corresponding antibody. Following the staining, the percentage of vacuolar and cytosolic bacteria can be quantified by FACS or microscopy by counting single or double-positive events. Here we provide experimental details for use of this technique with the bacterium Salmonella typhimurium. The advantage of this assay is that, in contrast to other assay, it provides a quantification on the level of single bacteria, and if analyzed by microscopy provides the exact number of cytosolic and vacuolar bacteria in a given cell.

Quantification of Cytosolic vs. Vacuolar Salmonella in Primary Macrophages by Differential Permeabilization

We describe the quantification of cytosolic and vacuolar Salmonella typhimurium in bone-marrow derived macrophages using differential digitonin permeabilization.

Intracellular bacterial pathogens can replicate in the cytosol or in specialized pathogen-containing vacuoles (PCVs). To reach the cytosol, bacteria like ...

Measuring Phagosome pH by Ratiometric Fluorescence Microscopy

Phagocytosis is a fundamental process through which innate immune cells engulf bacteria, apoptotic cells or other foreign particles in order to kill or neutralize the ingested material, or to present it as antigens and initiate adaptive immune responses. The pH of phagosomes is a critical parameter regulating fission or fusion with endomembranes and activation of proteolytic enzymes, events that allow the phagocytic vacuole to mature into a degradative organelle. In addition, translocation of H+ is required for the production of high levels of reactive oxygen species (ROS), which are essential for efficient killing and signaling to other host tissues. Many intracellular pathogens subvert phagocytic killing by limiting phagosomal acidification, highlighting the importance of pH in phagosome biology. Here we describe a ratiometric method for measuring phagosomal pH in neutrophils using fluorescein isothiocyanate (FITC)-labeled zymosan as phagocytic targets, and live-cell imaging. The assay is based on the fluorescence properties of FITC, which is quenched by acidic pH when excited at 490 nm but not when excited at 440 nm, allowing quantification of a pH-dependent ratio, rather than absolute fluorescence, of a single dye. A detailed protocol for performing in situ dye calibration and conversion of ratio to real pH values is also provided. Single-dye ratiometric methods are generally considered superior to single wavelength or dual-dye pseudo-ratiometric protocols, as they are less sensitive to perturbations such as bleaching, focus changes, laser variations, and uneven labeling, which distort the measured signal. This method can be easily modified to measure pH in other phagocytic cell types, and zymosan can be replaced by any other amine-containing particle, from inert beads to living microorganisms. Finally, this method can be adapted to make use of other fluorescent probes sensitive to different pH ranges or other phagosomal activities, making it a generalized protocol for the functional imaging of phagosomes.

Phagosomal pH influences phagosome maturation, oxidant production, phagosomal killing as well as antigen presentation. Here we describe a ratiometric method for measuring time-course and endpoint pH changes in individual phagosomes in living phagocytes using fluorescence microscopy.

Phagocytosis is a fundamental process through which innate immune cells engulf bacteria, apoptotic cells or other foreign particles in order to kill or ...

A Simple Fluorescence Assay for Quantification of Canine Neutrophil Extracellular Trap Release

Neutrophil extracellular traps are networks of DNA, histones and neutrophil proteins released in response to infectious and inflammatory stimuli. Although a component of the innate immune response, NETs are implicated in a range of disease processes including autoimmunity and thrombosis. This protocol describes a simple method for canine neutrophil isolation and quantification of NETs using a microplate fluorescence assay. Blood is collected using conventional venipuncture techniques. Neutrophils are isolated using dextran sedimentation and a density gradient using conditions optimized for dog blood. After allowing time for attachment to the wells of a 96 well plate, neutrophils are treated with NET-inducing agonists such as phorbol-12-myristate-13-acetate or platelet activating factor. DNA release is measured by the fluorescence of a cell-impermeable nucleic acid dye. This assay is a simple, inexpensive method for quantifying NET release, but NET formation rather than other causes of cell death must be confirmed with alternative methods.

Neutrophil extracellular traps (NETs) are networks of DNA, histones and neutrophil proteins. Although a component of the innate immune response, NETs are implicated in autoimmunity and thrombosis. This protocol describes a simple method for canine neutrophil isolation and quantification of NETs using a microplate fluorescence assay.

Neutrophil extracellular traps are networks of DNA, histones and neutrophil proteins released in response to infectious and inflammatory stimuli. Although ...

Isolation, Characterization, and Purification of Macrophages from Tissues Affected by Obesity-related Inflammation

Obesity promotes a chronic inflammatory state that is largely mediated by tissue-resident macrophages as well as monocyte-derived macrophages. Diet-induced obesity (DIO) is a valuable model in studying the role of macrophage heterogeneity; however, adequate macrophage isolations are difficult to acquire from inflamed tissues. In this protocol, we outline the isolation steps and necessary troubleshooting guidelines derived from our studies for obtaining a suitable population of tissue-resident macrophages from mice following 18 weeks of high-fat (HFD) or high-fat/high-cholesterol (HFHCD) diet intervention. This protocol focuses on three hallmark tissues studied in obesity and atherosclerosis including the liver, white adipose tissues (WAT), and the aorta. We highlight how dualistic usage of flow cytometry can achieve a new dimension of isolation and characterization of tissue-resident macrophages. A fundamental section of this protocol addresses the intricacies underlying tissue-specific enzymatic digestions and macrophage isolation, and subsequent cell-surface antibody staining for flow cytometric analysis. This protocol addresses existing complexities underlying fluorescent-activated cell sorting (FACS) and presents clarifications to these complexities so as to obtain broad range characterization from adequately sorted cell populations. Alternate enrichment methods are included for sorting cells, such as the dense liver, allowing for flexibility and time management when working with FACS. In brief, this protocol aids the researcher to evaluate macrophage heterogeneity from a multitude of inflamed tissues in a given study and provides insightful troubleshooting tips that have been successful for favorable cellular isolation and characterization of immune cells in DIO-mediated inflammation.

This protocol allows a researcher to isolate and characterize tissue-resident macrophages in various hallmark inflamed tissues extracted from diet-induced models of metabolic disorders.