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

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

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
<ol>
	<li>Culture THP-1 monocytes as a suspension culture in T25 flasks at 37 &#176;C + 5% CO2. Cells should be grown in 5 mL of Roswell Park Memorial Institute 1640 (RPMI...

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 apo...

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 addin...

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 ...

This method can be used to answer key questions in the efferocytosis field such as the role of signaling molecules in mediating the uptake of apoptotic cells. The main advantage of this technique is that it provides a clear quantification of apoptotic cell uptake including partial or piecemeal apoptotic cell uptake which isn't always apparent with other methods. Though this method can provide insight into efferocytosis by immune cells it can also be applied to other cell types such as epithelial or cancer cells.Generally, individuals new to this method will struggle because of the difficulty in optimizing microscope acquisition settings. After preparing human macrophages and apoptotic Jurkat cells, use a hemocytometer to count the apoptotic cells. Then transfer a sufficient quantity of apoptotic cells to a 1.5 milliliter microcentrifuge tube.Pellet the Jurkat cells via centrifugation at 500 times g for five minutes. Then resuspend the cells in 500 microliters of PBS. While the Jurkat cells are in the centrifuge, aliquot 10 microliters of DMSO into a new microcentrifuge tube.Next, dissolve a minimal amount of NHS-Biotin in the DMSO. After this, transfer 500 microliters of the apoptotic cell, and PBS suspension to the tube containing DMSO and NHS-Biotin. Then, dilute an appropriate cell-tracking dye according to the manufacturer's instructions.Incubate the cell suspension at room temperature for 20 minutes in the dark. Then, add an equal volume of RPMI 1640 with 10%FBS, and incubate the suspension at room temperature in the dark for an additional five minutes. After this, pellet the cells via centrifugation at 500 times g for five minutes.Then discard the supernatant, and resuspend the stained cells in 100 microliters of RPMI 1640 with 10%FBS. Add 100 microliters of stained cell suspension drop wise to each well of macrophages. Then centrifuge the plate at 200 times g for one minute.Incubate the plate at 37 degrees Celsius with 5%CO2 in a tissue culture incubator. After the appropriate incubation period, remove the cells from the incubator. Then wash the cells twice with one milliliter of PBS.After this, add diluted FITC-conjugated streptavidin to each well. And incubate the plate for 20 minutes in the dark. Wash the cells three times with one milliliter of PBS, and gently shake the samples for five minutes after each rinse.Then fix the cells with 4%paraformaldehyde in PBS for 20 minutes at room temperature. Rinse the cells once with PBS to remove any excess PFA. Mount coverslips on an imaging slide, and transfer the slide to a fluorescence microscope.Capture Z stacks of a sufficient number of cells for quantification. It is critical to set the distance between said sections such that all engulfed apoptotic material is detected. The maximum distance between said sections should be no greater than the focal depth of your objective lens, typically 0.5 to one micrometers.Load one of the Z stacks into the software using the Import feature. Then select Integrated density as a measurement option under the Set Measurements panel. Analysis is easiest to perform if both the streptavidin and cell tracking dye channels are displayed as an overlay.Any object with streptavidin staining anywhere along a circumference should be considered streptavidin positive, and therefore, should not be quantified as streptavidin negative. Once the image is loaded, using the streptavidin and the Freehand, or polygon selection tool, circle all of the cell tracking dye positives streptavidin negative material in a single plane of the Z stack. Then, measure the integrated intensity of the cell tracking dye in the region.In the same Z section use the streptavidin staining, and the Freehand or polygon selection tool to circle all of the cell tracking dye positive streptavidin positive material. Finally, measure the integrated intensity of this region. Using this procedure, more than 95%of Jurkat cells treated with Staurosporine went through apoptosis.In most experiments, the fraction of apoptotic cells that are efferocytosed increased in a time-dependent fashion. When optimizing this protocol for different efferocytic cell types, or different apoptotic cell targets, it is important to test the range of apoptotic cell to efferocytic cell ratios. Usually ratios of 10 to one or 30 to one work the best.This procedure can be easily modified to track signaling molecule activation, or intracellular trafficking by expressing fluorescent reporter genes in the efferocytic cells, thus allowing for these processes to be imaged during efferocytosis.

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Immunology and Infection

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting (FACS)

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known phenotype.  Other bulk methods of purification include panning, complement depletion and magnetic bead separation.  However, FACS has several advantages over other available methods.  FACS is the preferred method when very high purity of the desired population is required, when the target cell population expresses a very low level of the identifying marker or when cell populations require separation based on differential marker density.  In addition, FACS is the only available purification technique to isolate cells based on internal staining or intracellular protein expression, such as a genetically modified fluorescent protein marker.  FACS allows the purification of individual cells based on size, granularity and fluorescence.  In order to purify cells of interest, they are first stained with fluorescently-tagged monoclonal antibodies (mAb), which recognize specific surface markers on the desired cell population (1).  Negative selection of unstained cells is also possible.  FACS purification requires a flow cytometer with sorting capacity and the appropriate software.  For FACS, cells in suspension are passed as a stream in droplets with each containing a single cell in front of a laser.  The fluorescence detection system detects cells of interest based on predetermined fluorescent parameters of the cells.  The instrument applies a charge to the droplet containing a cell of interest and an electrostatic deflection system facilitates collection of the charged droplets into appropriate collection tubes (2).  The success of staining and thereby sorting depends largely on the selection of the identifying markers and the choice of mAb.  Sorting parameters can be adjusted depending on the requirement of purity and yield.  Although FACS requires specialized equipment and personnel training, it is the method of choice for isolation of highly purified cell populations.

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting FACS

For many scientific studies requiring a biological and chemical analysis of cell populations the cells must be in a high state of purity. Fluorescence activated cell sorting (FACS) is a superior method in which to obtain pure cell populations.

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known ...

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 ...

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.