A High-Throughput Real-Time Imaging Technique To Quantify NETosis and Distinguish Mechanisms of Cell Death in Human Neutrophils

Neutrophils are the most abundant type of WBCs in humans and play a critical role in the innate immune system, as well as in the regulation of the adaptive immune response. They are the first line of defense against microorganisms, and their microbicidal activity is modulated by various processes, such as the release of antimicrobial peptides present in neutrophil granules, including myeloperoxidase (MPO), neutrophil elastase, and matrix metalloproteinases through degranulation; phagocytosis and degradation of microbes inside phagolysosomes by production of reactive oxygen species (ROS); and the release of a meshwork of chromatin bound to granule peptides termed neutrophil extracellular traps (NETs) that can immobilize and kill microorganisms and activate other immune cells (reviewed in Ref. 1).

NETs are generated and extruded during a distinct form of programmed neutrophil cell death known as NETosis, which differs from apoptosis and necrosis (2). NETosis involves activation of multiple protein kinases (including Akt and protein kinase C) (3, 4) and the generation of ROS through NAD phosphate–oxidase complex or mitochondria (4, 5), followed by MPO-dependent translocation of elastase from granules to the nucleus (6). The nuclear translocation of these granules, along with the enzyme peptidyl-arginine deiminase type-4 (7), promote chromatin decondensation through histone degradation and histone citrullination. Eventually, chromatin and granule proteins mix in the cytoplasm, followed by cell membrane compromise and extracellular extrusion of a web of nuclear and granule material and neutrophil lysis. NETosis is a mechanism of death that appears to occur under conditions of inflammation (microbial or sterile). Under conditions of homeostasis, neutrophils die by apoptosis, a cell death process that can occur under normal physiological conditions or as a result of external stimuli. This caspase-dependent process is characterized by chromatin aggregation, nuclear and cytoplasmic condensation (8, 9), cell membrane blebbing, and formation of membrane-bound vesicles called apoptotic bodies that contain cytoplasmic and nuclear material (10, 11). Neutrophils can also die by necrosis, which is caused by physical disruption to the cell when exposed to extreme variations from physiological conditions. This results in influx of water and extracellular ions and leads to rounding of the cells and swelling of the organelles and cell membrane to the point of rupture and release of cytoplasmic contents, whereas the nucleus remains relatively intact (12). Apoptotic cells that are not properly cleared by phagocytes can also undergo secondary necrosis (reviewed in Ref. 13).

Neutrophils and NETosis play a key role in host defense against pathogens (2, 14–17) as well as in critical biological processes like modulation of innate and adaptive immunity (18) and endothelial damage (19). NETosis can be triggered by infectious and inflammatory “sterile” stimuli, including cytokines (20), autoantibodies (21), immune complexes (22), PMA (2), platelet products (e.g., platelet activating-factor) (23), bacterial toxins (e.g., nigericin) (24), and calcium ionophores (e.g., A23187) (4). NETs have been proposed to play crucial roles in the pathogenesis of systemic autoimmune diseases, including systemic lupus erythematosus (SLE) (25), rheumatoid arthritis (26), anti-phospholipid Ab syndrome (21), and systemic vasculitis (27), by contributing to the initiation and perpetuation of immune dysregulation, modification, and externalization of autoantigens and tissue damage (reviewed in Ref. 1). In addition, NETs have been implicated in a variety of pathological conditions beyond autoimmunity, including cancer (28), atherosclerosis (29), diabetes (30), thrombosis (31), and pancreatitis (32). As such, there is a critical need for better approaches to rapidly and accurately assess the role of various stimuli and inhibitors that modulate neutrophil physiology.

Commonly used methodologies to detect and quantify NETosis have been based on conventional fluorescence microscopy by identification of extruded granule proteins bound to DNA and/or histones. This method has become the gold standard procedure to visualize NETosis and assesses the endpoint of nuclear extrusion. However, this procedure is very laborious, lengthy, provides low throughput, and is usually performed at a single user-defined time point. It is not unusual that up to 2 d are required for quantification of NETosis by this method. Furthermore, it is susceptible to operator bias. Variations in plating techniques, cell adherence during staining steps, imaging, and counting practices can distort the quantification and limit the use of this method when multiple experimental conditions are desired. Other methods to quantify NETosis have included semiautomated plate assays that use membrane-impermeable dyes (e.g., SYTOX Green) that quantify extracellular DNA. These assays lack specificity, given the inability to visualize morphological changes, distinguish specific mechanisms of cell death, and accurately account for variations in cell count. We recently reported the use of high-speed multispectral imaging flow cytometry as a helpful technique in assessing NETosis in a rapid automated manner, also allowing for the ability to quantify the stages of NETosis preceding cell lysis (33). A limitation of this new technique is that the assessment also occurs only at a single time point and only images cells that are actively undergoing NETosis, with the potential to miss those cells that have already died or are delayed in their response. Thus, techniques to assess and quantitate NETosis in real time in an unbiased, reproducible, and efficient way are lacking.

In this article, we describe the optimization and validation of a novel method to automatically quantify the percentage of neutrophils undergoing NETosis using a two-color, live-content imaging platform IncuCyte ZOOM (Essen BioScience) system and a membrane integrity–dependent dual-dye system to stain DNA. In addition, using high-definition phase contrast imaging, in combination with the signal of fluorescent dyes, the IncuCyte ZOOM platform was able to distinguish among various types of neutrophil cell death induced by different stimuli based on morphological characterization of the cell and the nucleus. We show that the assay using this platform is rapid, unbiased, reproducible, and more sensitive than conventional fluorescence microscopy. The real-time kinetics data acquired on this platform can be used for studies using various stimuli and inhibitors of NETosis, apoptosis, or necrosis with potential use in the identification of prospective therapeutic targets.

All healthy controls and individuals with SLE were recruited at the Clinical Center, National Institutes of Health, and signed informed consent on National Institute of Arthritis and Musculoskeletal and Skin Diseases/National Institute of Diabetes and Digestive and Kidney Diseases Institutional Review Board–approved protocols. Peripheral blood was obtained by venipuncture and collected in heparinized tubes. Neutrophils were isolated at room temperature from freshly drawn peripheral blood using a Ficoll-density gradient (GE Healthcare) and dextran sedimentation, followed by RBC lysis with hypertonic solution, as described previously (34). Neutrophils were resuspended in RPMI 1640 without phenol (2 × 10⁶ neutrophils per milliliter). Lupus low-density granulocytes (LDGs) were purified as described previously by our group (35).

Neutrophils (2 × 10⁶ cells per milliliter) were incubated for 5 min in the dark at room temperature with the membrane-permeable NUCLEAR-ID Red DNA dye to stain nuclei (1 μl/1.5 ml of cell suspension; Enzo Life Sciences). Cells were washed by centrifugation at 2400 relative centrifugal force for 5 min, follow by supernatant removal and resuspension of the pellet in 1 ml of RPMI 1640 media. This washing process was repeated three times to remove excess dye. NUCLEAR-ID Red–stained neutrophils were plated at the optimized cell density of 20,000 neutrophils per 100 μl per well to facilitate segmentation and quantitation by the integrated image processing software in a 96-well flat clear-bottom polystyrene tissue culture–treated microplates (uncoated, Costar 3596; Corning). A final concentration of 0.2 μM membrane-impermeable dsDNA fluorescent SYTOX Green Nucleic Acid Stain (Life Technologies) was added to the plated cells concomitantly with the stimuli to assess cell death. For MPO staining, mouse anti-human MPO Ab FITC (1:100; Abcam) was added with the stimuli to the wells instead of SYTOX Green dye and imaged with phase contrast, red (1600-ms exposure), and green (800-ms exposure) channels.

Neutrophils were stimulated with bacterial toxin nigericin (0.5 μM), calcium ionophore A23187 (25 μM), PMA (0.5 μM), or platelet-activating factor (3 mM; all from Sigma-Aldrich) to induce NETosis. To induce apoptosis, cells were treated with staurosporine (1 μM; Sigma-Aldrich). Necrosis was induced by adding Triton X-100 (0.005%; AmericanBio). The stimulus and SYTOX Green dye were prepared at 2× in 100 μl of RPMI 1640 and added to the wells already containing 100 μl of neutrophils in RPMI 1640 media to make a total volume of 200 μl. For inhibition assays, neutrophils already stained with NUCLEAR-ID Red DNA dye were incubated with various concentrations of Akt inhibitor XI (Calbiochem) or diphenyleneiodonium chloride (Sigma-Aldrich) for 30 min before plating the cells and adding the stimuli for NETosis along with SYTOX Green dye.

Neutrophils were imaged within 10 min of plating using phase contrast, red (800-ms exposure), and green (400-ms exposure) channels in the IncuCyte ZOOM platform, which is housed inside a cell incubator at 37°C with 5% CO₂. Three image sets from distinct regions per well were taken every 5–15 min using a ×20 dry objective, and each condition was run in triplicate. To accomplish quantification using IncuCyte Basic Software, representative images of unstimulated and PMA-induced NETosis conditions at various time points were added to an image collection to describe to the software what objects to recognize as neutrophils versus neutrophils undergoing NETosis by defining specific parameters for fluorescence intensity, fluorescent area size, and radius. IncuCyte software’s processing definition was set to recognize neutrophils by the red-stained nucleus. Similarly, cells undergoing NETosis were identified by green staining following membrane damage. For the red and green channels, the TopHat method was selected for background correction. Similarly, the edge split tool was turned on in red and green channel analyses for accurate quantification of closely spaced objects. For the red channel, filters were applied to exclude objects with a radius < 10 μm, fluorescence threshold of 0.5 red corrected units, and area of 15 μm². For the green channel, edge sensitivity was set to −10, and hole fill was set to 100 μm². The filters applied excluded objects with a radius < 10 μm, fluorescence threshold of 1.00 green corrected units, and area of 100 μm². This minimum area threshold for green was used to recognize the decondensed chromatin of cells. The image collection was previewed using this processing definition and reviewed using the object masks to confirm that the red object mask captured all neutrophils’ nuclei stained with NUCLEAR-ID Red and that the green object mask captured the decondensed nucleus of neutrophils stained with SYTOX Green, which only binds to DNA of cells with compromised membranes (Fig. 1E).

Once this processing definition was optimized, it was stored and used for analyses of all future experiments. Graphics were generated with IncuCyte Basic Software graph/export functions. To determine the number of neutrophils per well, the average of red object counts (per image) for all three images per well was used. In the graph, the time point at which the red object counts plateau indicates that the cells have settled to the bottom of the well, and the object count at this time point is used for the total number of neutrophils. To quantify cells undergoing NETosis, the average of green object counts (per image) for all three images per well was graphed, and the time point of maximal NETosis was used to generate counts. To determine the percentage of cells undergoing NETosis, the green object count was divided by the red object count.

Graphics of the kinetics of cell death and NETosis fingerprint were generated in real time by plotting average green object counts per time point for each condition using the IncuCyte Basic Software or by exporting data and using GraphPad Prism software.

Healthy control neutrophils were stained with NUCLEAR-ID Red dye (red staining of the nucleus) and washed three times. They were then plated in the presence of stimuli and cell membrane–impermeable SYTOX Green dye in RPMI 1640 and imaged every 5 min using the IncuCyte ZOOM platform. Motion artifacts in jittery image stack were minimized using the Lucas–Kanade algorithm through the ImageStabilizer plugin in ImageJ software created by K. Li (http://www.cs.cmu.edu/∼kangli/code/Image_Stabilizer.html).

After 4 h of incubation with various stimuli and imaging with IncuCyte ZOOM, neutrophils plated in the 96-well plates were fixed by adding 10 μl of 4% paraformaldehyde and 10 μl of Hoechst 33342 (0.2 mg/ml). These wells were visualized using a Zeiss 780 confocal microscope with a ×10 Plan-Apochromat 0.45 NA dry objective in sequential mode. SYTOX Green was visualized with 488-nm laser light, and emission was collected at 449–552 nm; NUCLEAR-ID Red was visualized with 561-nm laser light, and emission was collected at 570–686 nm; and Hoechst was visualized with 405-nm laser light, and emission was collected at 410–472 nm.

Fluorescence microscopy quantification was done as previously described (34). Briefly, isolated neutrophils were incubated (37°C, 5% CO₂) in RPMI 1640, with or without stimulus conditions, on poly-l-lysine–coated glass coverslips for 3 h. The neutrophils were fixed in 4% paraformaldehyde and stored at 4°C overnight. NETs were stained by washing the fixed cells with PBS and incubating with rabbit anti-human MPO Ab (1:1000; Dako) for 60 min at 37°C, followed by incubation with secondary fluorochrome-conjugated Ab (1:200, A31572, Alexa Fluor 555 Donkey anti-Rabbit Ab; Life Technologies) for 30 min at 37°C. Nuclear DNA was detected by incubating cells with Hoechst 33342 (1:1000; Life Technologies) for 10 min at room temperature. After mounting (ProLong; Life Technologies), cells were visualized using fluorescence microscopy (Leica DMI4000B; Leica Microsystems). The images were loaded on Adobe Photoshop (Adobe Systems) for further analysis, in which NETs were manually quantified. The number of cells positive for neutrophil MPO and nuclear staining (Hoechst) were considered a NET and digitally marked to prevent multiple counts. The percentage of NETs was calculated as the average of three fields (original magnification ×20) normalized to the total number of cells.

Statistical analysis was performed using the paired t test, and data were analyzed using GraphPad Prism software version 6 (GraphPad). Results are presented as the mean ± SEM. The Spearman correlation coefficient was also used.

The IncuCyte ZOOM live content imaging system is configured to quantitatively measure in three imaging channels: phase contrast, green, and red fluorescence. Using the phase contrast images to assess cell morphology, the red dye to stain and count nuclei, and SYTOX Green to mark cells that were dying, we first sought to optimize the IncuCyte ZOOM platform to count neutrophils undergoing NETosis after PMA stimulation, a robust and reproducible inducer of NETs (2). To identify all of the nuclei of freshly isolated neutrophils without affecting the cells’ health, we used NUCLEAR-ID Red, an inert fluorescent membrane-permeable dsDNA dye. This dye freely crosses the cell membrane and fluoresces red when bound to nucleic acid (Fig. 1A). A concentration of 5 μM was found to have the optimal signal/noise ratio. The cells were stained for 5 min and then washed three times to remove excess dye. The cells were then seeded on a 96-well clear flat-bottom plate, and PMA with SYTOX Green dye in RPMI 1640 was added. After placing the plate in the IncuCyte ZOOM machine, images were acquired every 15 min. A processing definition was set up to define cells whose nuclei were stained red and cells that were stained green based on the fluorescence intensity, radius, and area. Total number of cells was calculated after 30 min to allow it to equilibrate and was based on red object counts (Fig. 1A, 1B). Cells that had undergone NETosis, based on nuclear decondensation, as assessed by a nuclear area size ≤ 100 μm² measured by fluorescence dsDNA SYTOX Green dye, were counted using the green mask (Fig. 1C, 1D). The percentage of cells undergoing NETosis was calculated by dividing NETotic cells by the total number of cells per well (Fig. 1E).

Once the initial processing definition was set up, automated real-time image processing was accomplished by applying the same processing definition at the onset of each experiment and analyzing images in real time as they were collected. This allowed us to simultaneously generate graphics of the data. Live imaging and real-time counts allowed for the assessment of the time to peak NETosis with various stimuli (Fig. 2A). With this unbiased and rapid technique, we saw distinct patterns in the kinetics of NETosis in neutrophils stimulated with various NET-inducing stimuli, the combination of which we call a NETosis fingerprint. PMA-induced NETosis occurred more rapidly compared with nigericin or A23187, both showing shifting of the NETosis curve to the right. These fingerprints were consistent across the various stimuli; however, they varied slightly between individuals (Supplemental Fig. 1A, 1B). Of note, NETosis induced by various stimuli was not different when comparing 96-well tissue culture–treated plates, optical bottom plates with cover glass base (uncoated, Nunc 164588; Thermo Scientific), or poly-l-lysine coating of the polystyrene or cover glass plates (Supplemental Fig. 1C–F).

IncuCyte ZOOM’s ability to directly visualize characteristic changes in cell and nuclear morphology, as well as the different kinetics of cell death of the entire population of cells, was used to distinguish distinct mechanisms of cell death induced by various stimuli (Tables I, II). Neutrophils stimulated with PMA (Figs. 2B, 3A), nigericin (Fig. 3B), A23187 (Supplemental Fig. 2A), and platelet-activating factor (Supplemental Fig. 2B) underwent NETosis; this was confirmed by visualization of the loss of nuclear lobulation, chromatin decondensation detected as an increase in nuclear diameter, and a decrease in the intensity of NUCLEAR-ID Red dye, followed by mixing of DNA content with cytoplasmic content and eventual cell membrane permeability and staining of nuclear content with SYTOX Green (Fig. 2B). NETosis was also confirmed by fixing the cells and staining with Hoechst in 96-well plates after 4 h and looking for NETs using a confocal microscope; extruded NETs, stained with red nuclear dye, Hoechst, and SYTOX, were visualized (Supplemental Fig. 3A–C). In addition, the findings were confirmed using IncuCyte ZOOM by staining neutrophils with mouse anti-human MPO-FITC that was added concomitantly with the stimuli instead of SYTOX Green. Characteristic of the NETosis process (34), MPO (stained green) colocalized with the DNA (stained red) once the membrane was permeabilized during NETosis (Supplemental Fig. 3D, 3E). In contrast, staurosporine, an apoptosis- specific stimulus, induced characteristic nuclear condensation with the appearance of cell membrane blebbing (36) (Figs. 2B, 3C), whereas necrosis induced by Triton X-100 was visualized as rapid disintegration of cell membrane resulting in staining of a relatively intact nucleus (Figs. 2B, 3D) (37). Time-lapse videos of neutrophils that were left unstimulated, as well as neutrophils undergoing PMA and nigericin-induced NETosis, staurosporine-induced apoptosis, and Triton X-100–induced necrosis are available (Supplemental Videos 1–5). Similarly, the kinetics of cell death by various mechanism was noted to be distinct and could also be used to distinguish among various types of cell death using this method (Fig. 2A). Plotting of the green signal in cells exposed to conditions that induced non-NETotic cell death allowed us to create additional fingerprints that were distinctly different from NETosis. Necrosis was associated with a rapid cell death, with time to peak reached within minutes, whereas apoptosis was characterized a delayed cell death, with an eventual peak reached at 7–9 h, enabling the use of this method to quantify and distinguish different forms of neutrophil cell death.

Having optimized the IncuCyte ZOOM for NETosis assessment, we proceeded to validate the quantitative imaging platform by comparing it with the established method of fluorescence microscopy (Fig. 4). The percentage of neutrophils undergoing NETosis correlated significantly with quantification done by fluorescence microscopy (R = +0.81, p < 0.0001) (Fig. 4G). The percentage of neutrophils undergoing NETosis at the 3-h time point was higher using the IncuCyte ZOOM method than by immunofluorescence microscopy, suggesting that IncuCyte ZOOM is a more sensitive method to assess NETosis because it allows for the identification of cells that have initiated this cell death process with cell membrane permeabilization but have not yet extruded the nuclear material to the extracellular space.

Activation of Akt, a serine/threonine-specific protein kinase, is critical in driving neutrophils to undergo NETosis. Inhibition of Akt significantly decreases PMA-induced NETosis (4, 38). Using the IncuCyte ZOOM and fluorescence microscopy for confirmation, we assessed the ability of different concentrations of Akt inhibitor XI to decrease the percentage of cells undergoing NETosis. This inhibitor decreased the percentage of PMA-induced NETosis in a dose-dependent manner, with peak inhibition of up to 75% achieved at a concentration of 30 μM (Fig. 5A–F). For NETosis induced by the microbial toxin nigericin, peak inhibition of up to 50% was achieved at a concentration of 20 μM (Fig. 5G–L). In addition to Akt inhibition, we tested the ROS inhibitor diphenyleneiodonium chloride, which also resulted in a dose-dependent inhibition of NETosis induced by PMA and nigericin (Supplemental Fig. 4) (17).

Overall, these results suggest that IncuCyte ZOOM accurately quantifies the effects of NET inhibitors.

A distinct subset of proinflammatory neutrophils has been identified in patients with SLE (35). These cells are isolated from the PBMC fraction and were named LDGs. These cells display significant enhancement in their capacity to spontaneously form NETs. Considering their putative important pathogenic role in this disease, inhibition of pathways of NET production may decrease the inflammatory capacity of these cells. Circulating LDGs demonstrated increased NETosis compared with normal-density neutrophils from lupus subjects or healthy controls, without addition of any stimuli (Fig. 6). This supports previous observations in which LDGs were shown to be primed in vivo to undergo NETosis and confirmed that IncuCyte ZOOM can quantify differences in NET formation among neutrophil subsets.

Neutrophils and NETosis have emerged as important putative players in the development of systemic autoimmune diseases, cancer, cardiovascular disease, and other acute and chronic diseases. Progress in our understanding of the role of NETs in health and disease has been hampered by the lack of a rapid, automated, accurate, and unbiased methodology to quantify NETosis in vitro and to distinguish this process from other forms of neutrophil cell death. In this article, we describe the optimization of a novel, reproducible, specific, and efficient imaging platform that acquires images in real time and in an operator-independent manner to quantify forms of neutrophil cell death (Table II). We have demonstrated the quantitative feature of this technology by quantifying neutrophil responses to various triggers of NETosis and NET inhibitors. In addition, we show that this platform is able to distinguish among distinct forms of neutrophil death through distinct morphology and by the generation of characteristic fingerprint curves.

The data analysis software is an innovative feature of IncuCyte ZOOM that helps to distinguish individual cells, analyze multiple conditions, and generate real-time kinetics data in a rapid and reproducible fashion. After setting up an initial processing definition, the operator can apply the same settings to analyze all images in the well with various stimuli, which are subsequently averaged for each well and replicates, thus eliminating potential selection bias. The processing definition can be optimized to select for appropriate cell size and fluorescence intensity to only account for cells undergoing NETosis. The edge split software function can reliably distinguish between closely located cells. Once the image-processing definition is set up and optimized, image analyses can be done automatically in real time, with results generated as the neutrophils are incubating with the stimulus. A time course of the green object count can be used to assess the number of dead cells. It is possible to generate kinetics data on the percentage of neutrophils dying by NETosis through this method at multiple user-specified time intervals.

IncuCyte ZOOM also offers a high-throughput platform, and the ability to perform experiments in 96-well plates considerably increases the capability to detect the effectiveness of various stimuli in parallel and to quantitate the efficiency of multiple concentrations of inhibitors to block NETosis. This methodology may prove to be a valuable tool in assessing the contribution of various proteins and regulatory complexes in neutrophil cell death and to identify and validate future putative therapeutic modalities. Indeed, compared with conventional fluorescence microscopy, the amount of time required to perform the analysis is decreased by nearly 6–8 h, depending on how many conditions are tested. Notably, the percentage of neutrophils undergoing NETosis, as quantified by IncuCyte ZOOM, correlates well with the gold standard of fluorescence microscopy. Indeed, the increased percentage of NETing neutrophils reported by IncuCyte ZOOM compared with immunofluorescent microscopy may reflect IncuCyte ZOOM’s ability to detect membrane compromise and mixing of granule with nuclear material intracellularly (an early event in NETosis) before release of NETs to the extracellular space. As such, IncuCyte ZOOM may provide higher sensitivity without decreasing specificity.

The IncuCyte ZOOM platform quantifies the percentage of neutrophils undergoing NETosis and does not quantify extracellular DNA. This is a considerable advantage over current plate-based assays, because it adjusts for the number of cells plated per well and increases sensitivity and specificity. We confirmed that cells quantified as undergoing NETosis by IncuCyte ZOOM were indeed extruding NETs when observed by confocal imaging. The ability to visualize actual NETs by confocal microscopy, but not by IncuCyte ZOOM, is likely due to the smaller numerical aperture and the number of pixels obtained per cell that affects the resolving power. It could also be due to limitations in the light source and detectors that can affect visualization of dimmer NETs. In addition, because NETs are narrower than the cell body, the autofocus of the IncuCyte ZOOM will preferentially image the contrasted cell body. Although an assay using confocal microscopy will provide great detail of individual cells undergoing NETosis, it will be unable to provide rapid real-time kinetic high-throughput results for multiple samples under various conditions. We show that this platform can generate results in multiple samples with different stimuli and inhibitors at multiple time points, with each run in triplicate, increasing the number of cells visualized per condition and considerably limiting bias.

Importantly, the IncuCyte ZOOM platform can distinguish among various forms of neutrophil cell death. In NETosis, the characteristic changes of loss of nuclear lobulation, nuclear decondensation, and mixing of nuclear content with cytoplasmic proteins can be monitored, in addition to the release of NETs to the extracellular space in the later stages of NETosis. In apoptosis, a more gradual condensation and fragmentation of the nuclei, with eventual membrane blebbing, become apparent. In contrast, necrosis induced by Triton X-100 was characterized by rapid cell swelling, immediate loss of cell membrane integrity, and a nucleus that remained lobulated. Due to dispersion of chromatin material outside the cell boundaries over time, necrosis should be further confirmed by direct visualization of the cells and the population kinetics of cell death, particularly with new stimuli. Other programmed cell death mechanisms, such as necroptosis (39), a receptor interacting protein-1 kinase–dependent process, and pyroptosis (40, 41), an inflammasome and caspase-1–dependent process, have similar morphological features as those seen in necrosis (12), and this methodology may not allow these forms of cell death to be distinguished from necrosis.

There are some other limitations of this methodology. The platform is only able to detect fluorescence in one green and one red channel, restricting the user from labeling other cellular components. Because of the limitation in the optical zoom focus, higher resolution of individual cells is not possible but may be complemented by assessment using fluorescent or confocal microscopy techniques, especially when testing new conditions. Impairment in plasma membrane integrity due to various nonspecific stimuli can give false positive results; however, direct visualization of the cells using a control without stimuli and stringent size and fluorescence intensity cutoff can help to mitigate these issues. The availability of the instrument may be a limiting factor; however, with the expanding role of this platform in studying the physiology of other cells (42–44), as well as its ability to accommodate multiple plates and flask sizes and image more than one plate at a time, the use of the IncuCyte ZOOM system may become more widespread.

In summary, we describe a novel, real-time, image-based platform using IncuCyte ZOOM that allows for a rapid unbiased quantification of NETosis and other types of neutrophil cell death, offering a significant advancement in the study of neutrophil biology. This imaging platform enables visualization and quantification of NETosis in real time. Our findings suggest that the specificity and sensitivity of this method may allow the accurate acquisition of kinetics data and visualization of various stages of cell death processes. This method supports a dynamic characterization not previously reported that can be used as a standard for comparison among samples and among different research groups. Furthermore, this technique can differentiate among distinct types of neutrophil cell death in response to various stimuli. This imaging platform represents an important improvement that can complement current methods to study neutrophil physiology in vitro and to evaluate novel therapeutic targets.

We thank Dr. Evelyn Ralston (National Institute of Arthritis and Musculoskeletal and Skin Diseases, National Institutes of Health) for support and feedback on this project.

The authors have no financial conflicts of interest.