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Immunology and Infection

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

doi: 10.3791/58149 Published: August 18, 2018


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 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.


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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 "eat-me" signals on their cell surface which are recognized by receptors on a range of professional and non-professional phagocytes, collectively referred to as "efferocytes". 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 "don't eat me" 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 — 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.

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Collection of blood from healthy volunteers was approved by the Health Science Research Ethics Board of the University of Western Ontario. Venipuncture was performed in accordance with the guidelines of the Tri-Council Policy Statement on human research.

1. Culture and Preparation of the THP-1 Monocyte Cell Line

  1. Culture THP-1 monocytes as a suspension culture in T25 flasks at 37 °C + 5% CO2. Cells should be grown in 5 mL of Roswell Park Memorial Institute 1640 (RPMI 1640) + 10% Fetal Bovine Serum (FBS).
  2. Each day suspend cells evenly throughout the growth media by gently shaking the flask, then immediately count cells with a hemocytometer. Cells should be passaged once cell density reaches 1 x 106 cells/mL:
    1. Pre-warm RPMI 1640 + 10% FBS in a 37 °C water bath.
    2. Transfer 2 x 105 cells into a 1.5 mL microcentrifuge tube or a 15 mL conical tube, and pellet cells by centrifuging at 500 x g at room temperature for 5 min.
    3. Remove the supernatant without disturbing the cell pellet and resuspend the pellet in 1 mL (1.5 mL microcentrifuge tube) or 5 mL (15 mL conical tube) of phosphate-buffered saline (PBS).
    4. Centrifuge the tube at 500 x g at room temperature for 5 min. Remove the PBS without disturbing the cell pellet.
    5. Resuspend pellet in 1 mL of fresh RPMI 1640 + 10% FBS.
    6. Into a new T25 flask place 4 mL of warmed media, and to this add the resuspended cells from 1.2.5. Culture in a 37 °C + 5% CO2 incubator until the cells require passaging (typically 3 days), or until cells are required for an experiment.
  3. For an experiment with THP-1-derived macrophages, remove the required number of cells from the flask and plate prior to passaging:
    1. Place the required number of 18 mm circular glass coverslips (#1.5 thickness) into the wells of a 12-well plate — typically 1 coverslip per condition and/or timepoint. Into each well aliquot 5 x 104 THP-1 monocytes. The number of cells added to each well can be altered, if required.
    2. Bring up the total volume of each cell-containing well to 1 mL using RPMI + 10% FBS warmed to 37 °C.
    3. Add 100 nM phorbol 12-myristate 13-acetate (PMA) to each well and culture for 3 days to induce differentiation of THP-1 monocytes into macrophage-like cells.

2. Culture and Preparation of the J774.2 Macrophage Cell Line

  1. Culture J774.2 cells in T25 flasks at 37 °C + 5% CO2. Cells should be grown in 5 mL of Dulbecco’s Modified Eagle Medium (DMEM) + 10% FBS and passaged once the culture reaches 80-90% confluency. To passage cells:
    1. Remove all media from the flask and rise once with 5 mL of PBS.
    2. Remove PBS from the flask and replace with 5 mL of fresh DMEM + 10% FBS.
    3. Using a cell scraper, scrape the bottom of the flask to suspend the cells in the media. Vigorously pipette the cells several times to break up any cell aggregates.
    4. Dilute cells 1:5 by removing 4 mL of the cell suspension from the flask and replacing it with 4 mL of fresh media. The remaining cell suspension can be discarded, used to start a new cell culture in a fresh T25 flask, or used for an experiment.
  2. To set up for an efferocytosis assay using J774.2 cells:
    1. One day prior to the start of the experiment, suspend J774.2 cells into 5 mL of fresh media, as per steps 2.1.1–2.1.3. Count cells using a hemocytometer and prepare the necessary volume of cells at a concentration of 5 x 104 cells/mL.
    2. Place the necessary number of 18 mm circular glass coverslips (#1.5 thickness) into the wells of a 12-well plate. Into each well aliquot 1 mL of the 5 x 104 cells/mL cell suspension.
    3. Culture overnight to allow the cells to adhere to the coverslip and recover from passaging.

3. Culture of Primary Human M2 Macrophages

  1. Collect 10 mL of heparinized human blood for every 12-well plate of M2 macrophages required.
  2. In a 15 mL centrifuge tube, layer 5 mL of human blood on top of 5 mL of pre-warmed Lympholyte-poly cell separation medium. Prepare multiple tubes if processing >5 mL of blood rather than using larger volume tubes.
  3. Centrifuge at 300 x g for 35 min, using medium acceleration and no break.
  4. Carefully remove the upper mononuclear-cell rich band using a plastic pipettor and transfer to a 50 mL centrifuge tube. If multiple tubes were prepared in step 3.2, the bands can be pooled into a single 50 mL tube. Bring volume of tube up to 50 mL with PBS.
  5. Centrifuge at 300 x g for 8 min and remove the supernatant. During this step place autoclaved 18 mm diameter circular coverslips (#1.5 thickness) into each well of a 12-well plate.
  6. Resuspend the cell pellet in 300 µL of serum-free RPMI 1640 per desired number of wells; e.g., if 10 mL of blood was processed to prepare a full 12 well plate, suspend cell pellet in 3.6 mL of media.
  7. Add 300 µL of the cell suspension to each coverslip-containing well in the 12-well plate. Incubate for 1 h at 37 °C + 5% CO2.
  8. Gently wash coverslip 3x with 1 mL of warmed PBS to remove any non-adherent cells.
  9. Add 1 mL of RPMI 1640 + 10% FBS + 10 ng/mL recombinant human M-CSF + cell culture antibiotic/antimycotic. Incubate at 37 °C + 5% CO2 for 5 days.
  10. Replace media with RPMI 1640 + 10% FBS + 10 ng/mL recombinant human M-CSF + 10 ng/mL recombinant human IL-4 + cell culture antibiotic/antimycotic. Incubate at 37 °C + 5% CO2 for 2 days to complete M2 polarization.
  11. Polarized macrophages should be used within the next 3 days.

4. Preparation of Apoptotic Jurkat Cells

  1. Culture Jurkat cells in 5 mL of RPMI 1640 + 10% FBS at 37 °C + 5% CO2. Jurkats are a suspension cell line and can be maintained by passaging 1:5 into fresh, pre-warmed media every 3–5 days.
  2. To prepare apoptotic cells, allow the Jurkat culture to grow to high density (4–5 days after passaging). Aliquot 1.5 mL into a 1.5 mL microcentrifuge tube and pellet cells by centrifugation at 500 x g for 5 min.
  3. Discard supernatant and re-suspend cell pellet in 1 mL of serum-free RPMI 1640 medium containing 1 µM staurosporine.
  4. Incubate 16 h at 37 °C + 5% CO2 to render cells apoptotic.
  5. If desired, confirm induction of apoptosis by staining with Annexin V:
    1. Aliquot 100 µL of staurosporine-treated Jurkat cell culture into a 1.5 mL microcentrifuge tube. Pellet cells by centrifugation at 500 x g for 5 min, discard supernatant, and re-suspend cell pellet in 100 µL of serum-free RPMI 1640 medium.
    2. Add 1 µL of fluorescein isothiocyanate (FITC)-conjugated Annexin V and incubate for 10 min at room temperature in the dark.
    3. Add 900 µL of PBS and transfer the entire volume to a single well of a 12-well plate containing an 18 mm circular glass coverslip (#1.5 thickness). Spin down in a centrifuge equipped with a plate adaptor at 200 x g for 1 min to force cells to adhere to the coverslip. Alternatively, cells can be placed into a chambered slide for imaging.
    4. Image using a fluorescence microscope.

5. Quantifying Efferocytic Uptake and Dynamics Using a Fixed Cell Efferocytosis Assay and Inside-out Staining

  1. Prepare THP-1, J774.2 or M2 human macrophages as described in Sections 1-3, respectively.
  2. The evening prior to the start of the experiment prepare apoptotic Jurkat cells as described in Section 4.
  3. Immediately prior to the experiment, count apoptotic cells using a hemocytometer. Transfer sufficient numbers of apoptotic cells into a 1.5 mL microcentrifuge tube – we usually add 5 x 105 cells/well, providing a target:efferocyte ratio of 10:1.
  4. Pellet Jurkat cells by centrifugation at 500 x g for 5 min and resuspend in 500 μL of PBS.
  5. During the centrifugation, aliquot 10 µL of DMSO into a new 1.5 mL microcentrifuge tube. Dissolve into the DMSO a minimal amount of N-hydroxysuccinimidobiotin (NHS-Biotin). 5-10 crystals (~0.005 mg) is sufficient.
  6. Transfer the 500 μL apoptotic cell/PBS suspension to the DMSO/NHS-biotin containing tube. Then dilute a cell tracking dye to the manufacturer’s recommended concentration into the apoptotic cell suspension. Make certain to select a cell tracking dye that does not overlap spectrally with FITC-Streptavidin (e.g. red or far-red cell tracking dye).
  7. Incubate suspension for 20 min at room temperature in the dark. Add an equal volume of RPMI 1640 + 10% FBS and incubate for 5 min at room temperature in the dark to quench any unreacted dye.
  8. Pellet cells by centrifugation at 500 x g for 5 min, discard supernatant, and re-suspend the stained apoptotic cells in 100 µL of RPMI 1640 + 10% FBS per well of macrophages.
  9. Add 100 µL of stained apoptotic cell suspension dropwise to each well of macrophages. Centrifuge 200 x g for 1 min in a centrifuge equipped with a plate adaptor to force contact between macrophages and apoptotic cells.
  10. Incubate plate for the desired period of time at 37 °C + 5% CO2 in a tissue culture incubator. For macrophages, efferocytosed material is usually first detectable after 20–30 min, and is completed after 120–180 min.
  11. At the desired time point(s) remove cells from the incubator. Wash cells twice with 1 mL of room temperature PBS to stop efferocytosis and remove non-efferocytosed apoptotic cells.
  12. Add FITC-conjugated streptavidin at a 1:1,000 dilution to each well and incubate for 20 min in the dark. This will label the exposed biotin on any non-efferocytosed apoptotic cell material.
    NOTE: If desired, cell nuclei can be stained during this step by addition of 1:20,000 dilution of Hoechst 33342 or 4',6-Diamidino-2-Phenylindole, Dihydrochloride (DAPI).
  13. Wash cells 3 times with 1 mL PBS, gently shaking or rocking the samples for 5 min per rinse. Fix cells with 4% paraformaldehyde (PFA) in PBS for 20 min at room temperature. Rinse cells once with PBS to remove excess PFA.
  14. Mount coverslips on a slide for imaging and transfer to a fluorescence microscope. Capture z-stacks of a sufficient number of cells for accurate quantification — typically 10–30 cells per condition. Non-internalized apoptotic cell material will be apparent in the resulting image as cell tracking dye labeled material enveloped by FITC-streptavidin staining, while efferocytosed material forms discrete cell tracking dye puncta free of any streptavidin staining.
  15. Quantify efferocytosis in the resulting images via a variety of measures:
    1. Calculate the efferocytic index by determining the average number of discrete efferosomes (cell tracking dye+/streptavidin- puncta) per macrophage. Quantify only macrophages bound to an apoptotic cell or containing ≥1 visible efferosomes. Record macrophages bound to an apoptotic cell, but lacking discrete efferosomes, as having an efferocytic index of 0.
    2. Calculate efferocytic efficiency by measuring the fraction of macrophages that contain ≥1 efferosome.
    3. Calculate the rate of efferocytosis by imaging cells fixed and stained at multiple time-points. Only image macrophages bound to apoptotic cells, or containing visible efferosomes, with z-stacks captured of each cell. Once imaged, determine the rate of efferocytosis:
      1. Using the streptavidin staining as a guide, and the freehand or polygon selection tool in FIJI/ImageJ26 (or other image analysis software package), circle all of the cell tracking dye+/streptavidin- (e.g. internalized) material in a single plane of the z-stack. Measure the integrated intensity of the cell tracking dye this region. Individual measures for each efferosome (e.g. diameter, positioning relative to the cell border or nucleus, etc.)15,27 can easily be acquired simultaneously with these measurements, greatly increasing the data collected during analysis.
      2. On the same z-section, using the streptavidin staining as a guide and the freehand or polygon selection tool, circle all of the cell tracking dye+/streptavidin+ (e.g. non-internalized) material in a single plane of the z-stack. Only include staining from apoptotic cells in contact with the phagocyte. Measure the integrated intensity of this region.
      3. Repeat steps to for the remaining z-sections of the image. Sum the integrated intensity of the efferocytosed (Σeff) and non-efferocytosed (Σne) materials. The fraction of the apoptotic cell which has been efferocytosed can be calculated for the cell as:
      4. Quantify the fraction efferocytosed for all cells at all time points. Note that the fluorescent intensity of apoptotic cells/efferosomes can vary between images due to variations in the uptake of cell tracking dyes by individual apoptotic cells, and due to changes in image acquisition parameters such as exposure time. As such, only compare normalized values such as Fraction Efferocytosed, or intensity-independent values such as efferocytic index and efferocytic efficiency, between images, between experimental conditions, and between repeat experiments.
    4. To ensure the resulting dataset accurately reflects the variation in efferocytosis between cells, conduct these analyses on the maximum number of cells possible in each experiment.
      NOTE: Efferocytic efficiency and efferocytic index can be rapidly calculated (a few seconds/cell), and we typically aim to analyze at least 100 cells per experiment, repeating each experiment at least 3 times. Calculating the fraction of efferocytosed material, and efferosome-specific measures are more laborious, typically taking 2–3 min/cell for an experienced analyst. For these calculations we quantify a minimum of 15 cells per condition, per experiment.
    5. Record efferocytic index, fraction efferocytosed and individual efferosome data on a per-cell basis.
      NOTE: This allows for data analysis using single-cell approaches, thus allowing for inter-cell variations to be quantified. Population-level analyses can still be conducted on these datasets by averaging single-cell data acquired in individual experiments. Efferosome-specific measurements can be analyzed at the population level, single-cell level, and as ensembles (e.g. as populations of efferosomes independent of the cells containing them)15.

6. Live Cell Efferocytosis Assay Using Apoptotic Cells

  1. Prepare THP-1, J774.2 or M2 macrophages as described in steps 1-3, respectively. If required, cells should be transfected with transgenes or other genetic constructs at least 18 h prior to performing any experiments.
  2. The evening prior to the start of the experiment, prepare apoptotic Jurkat cells as described in Sections 4 and 5 with the following modifications:
    1. Dilute cells and induce apoptosis as per 4.1–4.3, and collect the required number of apoptotic cells as per 5.3–5.4.
    2. Add a cell tracking dye at the manufacturer’s recommended concentration to the apoptotic cell suspension. Incubate suspension for 20 min at room temperature in the dark. Add an equal volume of RPMI 1640 + 10% FBS and incubate for 5 min at room temperature in the dark to quench any unreacted dye. Do not add NHS-biotin for these experiments.
    3. Pellet cells by centrifugation at 500 x g for 5 min, discard supernatant, and re-suspend stained apoptotic cells in 100 µL of RPMI 1640 + 10% FBS per well of macrophages.
  3. If required, label the macrophages by removing culture media from each well and adding 500 µL of PBS containing the manufacturers recommended dilution of a cell tracking dye that does not overlap spectrally with the cell tracking dye added to the apoptotic cells or any fluorescent transgenes expressed by the macrophages. Incubate for 20 min at room temperature in the dark, then add an equal volume of DMEM + 10% FBS and incubate for 5 min at room temperature in the dark to quench any unreacted dye.
  4. Transfer the macrophage-containing coverslip to a Leiden chamber. Add 400 µL of DMEM + 10% FBS, followed by 100 µL of the labeled apoptotic cell suspension. Mix by gently pipetting media 2–3 times.
  5. Transfer the Leiden chamber to a heated and CO2-perfused chamber of a live cell fluorescent microscope.
    NOTE: For microscope setups that lack CO2 perfusion capabilities, use tissue culture medium buffered with 1 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) rather than sodium bicarbonate to maintain physiological pH while the culture is exposed to air.
  6. Capture a time-lapse series containing a white light (phase contrast or DIC) image, and images of all fluorescent labels:
    1. Use a 100X objective lens for experiments where resolving fine details is required (e.g. membrane dynamics of phagocyte-apoptotic cell interactions). Meanwhile, more general measures (rate of apoptotic cell uptake, interaction time between macrophages and apoptotic cells, etc.) are best imaged using a 60X or 63X objective.
    2. If available, use point-visiting to image multiple fields-of-view during a single acquisition, thus increasing the number of efferocytic events captured in a single experiment
    3. Because efferocytes often engulf small fragments of apoptotic cells, rather than intact cells, it is best to capture z-stacks. To minimize phototoxicity, capture z-sections separated by the focal depth of your microscopes objective (typically 0.5–1.0 µm), through the thickness of the cell (typically 8-10 µm for macrophages, e.g. 8–20 slices/cell). This ensures that all engulfment and trafficking events will be visible in at least one z-plane. Alternatively, use large (1–5 µm diameter) apoptotic cell mimics or apoptotic cells that undergo minimal fragmentation during efferocytosis (e.g. heat shocked neutrophils) instead without z-stacking. We have described the preparation of both mimics and apoptotic neutrophils previously28.
    4. To minimize photobleaching, use bright fluorophores and capture images using acquisition settings that minimize photobleaching — e.g. low-intensity excitation combined with high camera gain and short exposure times29. We strongly recommend using a microscope equipped with a high-sensitivity electromagnetic charge-coupled device (EM-CCD) camera or spinning-disk confocal, and a high-speed piezoelectric mechanical stage, for this form of imaging.
  7. The number of possible analysis methods that can be applied to these time-lapse videos is extensive and beyond our ability to review here. As some examples, use manual or automated tracking software to track the fusion, fission and movement of efferosomes within cells15, quantify efferosome fusion dynamics with endolysosomes by colocalization analysis in macrophages expressing compartment-specific fluorescent transgenes13, monitor uptake processes such as probing and cup formation30, determine the recruitment dynamics of signaling and trafficking proteins to the efferosome13, and/or quantify the degradative activity of efferosomes using apoptotic cells labeled with pH-sensitive, oxidant-sensitive or protease-activated fluorophores31.

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Representative Results

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Overnight culture of Jurkat cells with 1 µM staurosporine results in apoptosis of >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, >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 µm 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 "probing"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 "efferocytic synapse"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, 10–30 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 — 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).

Figure 1
Figure 1: Annexin V Staining of Apoptotic and Non-Apoptotic Jurkat Cells. Annexin V labels the phosphatidylserine "eat-me" 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 µM staurosporine (STS) take on an apoptotic morphology (irregular and blebbed) and stain with Annexin V. Scale bar = 10 µm, image intensity is displayed as a color map. Please click here to view a larger version of this figure.

Figure 2
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 (mφ) 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 µm. Please click here to view a larger version of this figure.

Figure 3
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. 12–17 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 <0.05 compared to 30 min, Kruskal-Wallis test with Dunn correction. Please click here to view a larger version of this figure.

Figure 4
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 µm. Cell nucleus is stained with Hoeschst, DIC image shows cell morphology and is used to identify apoptotic cell mimics. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Live cell efferocytosis assay. A cell tracking dye labeledJ774.2 macrophage (mφ, 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 µm. Please click here to view a larger version of this figure.

Figure 6
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 µm. Please click here to view a larger version of this figure.

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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 — 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 — e.g. experiments detecting rare events, or quantifying highly variable processes — 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 2–3 min to <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 4–6 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 "find-me" 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 >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 — 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 (5–30 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 — but even with careful selection of fluorophores, excitation intensity and exposure times — photobleaching or phototoxicity will usually limit these experiments to less than an hour in duration. In our experience, acquiring images every 1–2 min, over experimental periods of 2–6 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.

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The authors declare no conflict of interest.


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.


Name Company Catalog Number Comments
RPMI 1640 Media Wisent 3500-000-EL
DMEM Media Wisent 319-005-CL
Fetal Bovine Serum (FBS) Wisent 080-150
PBS Wisent 311-010-CL
18 mm circular glass coverslips #1.5 thickness Electron Microscopy Sciences 72290-08 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
Staurosporine Cayman Chemical 81590 Dissolve in DMSO at 1 mM (1,000x stock solution)
Annexin V-Alexa 488 ThermoFisher R37174
EZ-Link NHS-Biotin ThermoFisher 20217 Store in a dessicator. Do not prepare a stock solution.
DMSO Sigma-Aldrich D2650
CellTrace FarRed ThermoFisher C34572
CellTrace Orange ThermoFisher C34851
Hoescht 33342 ThermoFisher 62249
FITC-Streptavadin ThermoFisher SA1001
Lympholyte-poly cell sepration medium Cedarlane Labs CL5071
Recombinant Human M-CSF Peprotech 200-04
Recombinant Human IL-4 Peprotech 300-25
J774.2 Macrophage Cell Line Sigma-Aldrich 85011428-1VL
THP-1 Human Monocyte Cell Line ATCC TIB-202
Jurkat T Cell Line ATCC TIB-152



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Quantification of Efferocytosis by Single-cell Fluorescence Microscopy
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Taruc, K., Yin, C., Wootton, D. G., Heit, B. Quantification of Efferocytosis by Single-cell Fluorescence Microscopy. J. Vis. Exp. (138), e58149, doi:10.3791/58149 (2018).More

Taruc, K., Yin, C., Wootton, D. G., Heit, B. Quantification of Efferocytosis by Single-cell Fluorescence Microscopy. J. Vis. Exp. (138), e58149, doi:10.3791/58149 (2018).

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