This work was supported by a Perpetual IMPACT Grant (IPAP201601422) and Novo Nordisk Foundation Biomedical Project Grant (NNF17OC0028990). YZ also acknowledges the receipt of an Australian Postgraduate Award from the University of Sydney. We would like to thank Mr. Pat Pisansarakit and Ms. Morgan Jones for assistance with the monocyte isolation and tissue culture.

The HMDM were isolated from human buffy coat preparations supplied by the blood bank with ethics approval from the Sydney Local Health District.
1. HMDM Culture
<ol>
	<li>Isolate the monocytes from buffy coat preparations prepared from the peripheral blood of healthy human donors using a commercially available preparation to isolate lymphocytes, followed by countercurrent centrifugal elutriation19,20.</li>
	<li>Confirm the presence of monocytes by cytospinning and staining with modified Giemsa stain for monocyte characterization19.</li>
	<li>Under sterile conditions, adjust the density of monocytes to 1 x 106 cells/mL using RPMI-1640 media without serum. Add 1 mL of this cell suspension to each well of a 12 well tissue culture plate. Culture in a cell incubator at 37 &#176;C in the presence of 5% CO2 for 2 h to promote adherence to the tissue culture plate.</li>
	<li>Under sterile conditions, remove the cell media and replace with complete RPMI-1640 culture media containing 10% (v/v) pooled human serum and 20 mM L-glutamine.</li>
	<li>Culture the cells at 37 &#176;C in the presence of 5% CO2 in a cell incubator for 8 days, changing the media every 2 days.</li>
</ol>
2. Polarization of HMDM
<ol>
	<li>Under sterile conditions, prepare the M1 priming media by adding interferon &#947; (IFN&#947;; 20 ng/mL) and lipopolysaccharide (LPS; 1 &#956;g/mL) to the complete RPMI-1640 culture media. Prepare the M2 priming media by adding interleukin 4 (IL-4; 20 ng/mL) to the complete RPMI-1640 culture media.</li>
	<li>Under sterile conditions, aspirate media from the tissue culture plate wells that contain the HMDM, which have been seeded and cultured as described in section 1.</li>
	<li>Carefully wash the wells containing the cells 3x with sterile PBS (pre-warmed to 37 &#176;C), using 1 mL aliquots of PBS.</li>
	<li>Add 1 mL of either the M1 or M2 priming media to each well containing the HMDM (whichever is appropriate for the experiment).</li>
	<li>Incubate the cells for 48 h at 37 &#176;C in the presence of 5% CO2 in a cell incubator.</li>
</ol>
3. Stimulation of HMDM to induce MET Release
<ol>
	<li>Under sterile conditions, prepare the culture media containing different stimulators of MET release (whichever is appropriate for the experiment) to the complete RPMI-1640 media: PMA (25 nM), human recombinant TNF&#945; (25 ng/mL), or human recombinant IL-8 (50 ng/mL).</li>
	<li>For experiments with HOCl stimulation, prepare HOCl (200 &#956;M) in HBSS (pre-warmed to 37 &#176;C), immediately before the addition to the cells. Ensure that the HOCl is not prepared in complete cell media. 
	NOTE: The concentration of the stock solution of HOCl is quantified by measuring the UV absorbance of the solution at 292 nm and pH = 116 and using an extinction coefficient of 350 M-1cm-1 21.</li>
	<li>After the polarization treatment described in section 2, aspirate the cell media from each well and carefully wash the cells 3x with 1 mL aliquots of either: sterile PBS (for PMA, TNF&#945; and IL-8) or HBSS (for HOCl), which have been pre-warmed to 37 &#176;C.</li>
	<li>For experiments with PMA, TNF&#945;, or IL-8: add 1 mL of the complete media containing PMA, TNF&#945;, or IL-8 after removing the PBS in the final washing step.</li>
	<li>For experiments with TNF&#945;, incubate the cells for 6 h at 37 &#176;C in the presence of 5% CO2 in a cell incubator. For experiments with PMA and IL-8, incubate the cells for 24 h at 37 &#176;C in the presence of 5% CO2 in a cell incubator.</li>
	<li>For experiments with HOCl, add 1 mL of HOCl in HBSS after removing the HBSS in the final washing step. Then, incubate the cells for 15 min at 37 &#176;C in the presence of 5% CO2 in a cell incubator.
	<ol>
		<li>Carefully aspirate the cell supernatant and wash the cells 3x with 1 mL aliquots of HBSS as described in step 3.3.</li>
		<li>After removing the HBSS from the final wash step, add 1 mL of complete RPMI-1640 culture media. Then, incubate the cells for 24 h at 37 &#176;C in the presence of 5% CO2 in a cell incubator.</li>
	</ol>
	</li>
</ol>
4. Visualization of MET in Live Cell Culture
<ol>
	<li>Prepare SYTOX green dye in HBSS at a concentration of 40 &#956;M.</li>
	<li>At the end of treatments described in section 3 to induce MET release, directly add 25 &#956;L of 40 &#956;M of the dye to each well containing HMDM.</li>
	<li>Incubate cells at room temperature (RT) for 5 min in the dark.</li>
	<li>Place the HMDM in tissue culture wells on the microscope stage of an inverted fluorescent microscope for imaging.</li>
	<li>Microscope procedures
	<ol>
		<li>Turn on a broad-spectrum fluorescent light source, brightfield light source, and inverted microscope installed with a high-resolution color digital camera (see Table of Materials).</li>
		<li>Rotate the filter wheel to the &#8220;number 2&#8221; position for green fluorescence (excitation = 504 nm, emission = 523 nm) for imaging of the green stained samples contained within the tissue culture wells.</li>
		<li>Using the 5x objective, focus the image with the coarse focus, then the fine focus knobs on the microscope, until the image appears sharp, clear, and focused when viewed through the microscope eyepiece.</li>
		<li>Switch the microscope to the camera mode.</li>
		<li>Start the associated software.</li>
		<li>Select the Capture tab on the software.</li>
		<li>Click the Play button to preview the image and adjust the fine focus knob on the microscope until the image appears sharp, clear, and focused in the software preview window.</li>
		<li>Click the Capture button. 
		NOTE: The captured image will automatically be displayed in the accompanying software.</li>
		<li>Within the software, click File | Save as the required image file type.</li>
		<li>On the microscope, rotate the filter wheel to the &#8220;number 5&#8221; position for brightfield imaging and repeat steps 4.5.2&#8211;4.5.9 to obtain the corresponding brightfield image.</li>
		<li>Repeat the steps 4.5.2&#8211;4.5.10 as necessary for subsequent image acquisition.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The generation and visualization of MET formation using M1 differentiated HMDMs represents a new in vitro model that may be useful for investigating the potential pathological role of these macrophage structures, particularly under chronic inflammatory conditions. It provides a robust protocol for the stimulation of primary human macrophages to release METs, which can also be utilized in related studies with human monocyte or murine macrophage cell lines. The successful implementation of this protocol for the stimulation of MET formation by HMDM is dependent on careful cell culture and handling to maintain good cell viability. This will increase the extent and quality of METs observed. To provide better cell nutrition, the RPMI media used to maintain the HMDM should contain human serum, rather than fetal bovine serum, as well as L-glutamine. Each stock solution of this complete RPMI media should be used within 1 week. Correct storage of the media is also important. The media should be stored at 4 &#176;C in the absence of light.
In addition, it is important to regularly observe and monitor the shape and morphology of the cells by microscopy during the culturing of HMDM. Any abnormal or unexpected changes in cell shape observed prior to the addition of IFN&#947; and LPS or IL-4 (to polarize the macrophages) may indicate a reduction in cell survival rate. HMDM should be adherent and do not proliferate. It is important, therefore, to monitor the cells for any changes in morphology, cell density, or loss of adherence during the normal 8 day culture period. A dramatic decrease in cell density during first 2 days post-isolation suggests that the resulting HMDMs may not be sufficiently viable for subsequent experiments. A limitation of using primary human cells is the variation between donors, which can markedly influence the extent of MET release. In addition, the quality of the buffy coat preparations received for the isolation of monocytes can vary. It is important to repeat experiments with at least three individual cell donors to ensure that data are representative of larger populations of cells.
It is important to note that the protocol described for HMDM differentiation has been optimized for cells isolated from human buffy coat preparations by countercurrent elutriation, which results in a high purity monocyte preparation. Validation of the polarization treatments (to confirm phenotype) by flow cytometry and assessment of the expression of M1 marker CD86 and M2 marker CD206 have been performed previously using this preparation of HMDM6. It is appreciated that this isolation method may be not be widely accessible, and that alternative isolation methods (i.e., magnetic bead sorting) may be preferable. If an alternative source of monocytes or different isolation protocols is used, additional flow cytometry experiments to validate phenotype change are recommended, and some optimization of treatment conditions to stimulate MET release may be required.
For optimal fluorescence images, exposure time should be carefully adjusted and kept as short as possible to avoid background fluorescence and staining artifacts from the plastic culture plates, which have adherent coatings. It is important to take multiple images from each well containing cells. It is advantageous if the samples are blinded before fluorescence microscopy analysis. In addition to staining extracellular strands of DNA, there is also evidence for the cellular uptake of SYTOX green. This is believed to reflect a loss in membrane integrity, which may be because of MET release, and it can also reveal alternative pathways of cell death (see also below). This highlights the importance of performing additional experiments to quantify and further characterize MET release.
For a quantitative comparison of the extent of MET release under different treatment conditions or following different interventions to modulate MET release, further analysis is required. This can be achieved by performing qPCR analysis of nuclear and mitochondrial DNA present in the cell supernatants, as described previously6. This method is usually combined with lactate dehydrogenase release assays to control for cell lysis unrelated to MET release6. The SYTOX green staining can also be quantified by a fluorescence plate reader following DNase treatment to partially digest the METs and release them from culture plates. This methodology has been used extensively in previous work with neutrophil NETs1,2,23.
The future applications of this protocol relate to providing opportunities for further characterization of METs using immunohistochemistry. It will be possible to probe the protein composition of METs using this approach with targeted antibodies (i.e., against citrullinated histones or MPO) to gain additional insight into the roles of these structures in more complex biological samples. These experiments will also be important for shedding more light on the delineation of MET release following other forms of cell death, such as pyroptosis15. It is worth noting that in our experience, METs from macrophages do not adhere to the plates as strongly as NETs from neutrophils, which can make immunohistochemistry analysis more challenging. Nonetheless, the data from these experiments will be of interest, given the fact that macrophages often play a dominant role in chronic inflammatory diseases. Further characterization of these structures is critical to assess their roles in vivo, which has thus far only been performed to a limited extent16. It is believed that the described method using HMDM to achieve this purpose may be more clinically relevant in comparison to other immortalized cell line-derived macrophage sources; although, these cell lines may have utility in providing a more stable model for MET characterization with less potential for donor variation.

The release of ETs from neutrophils was first identified as an innate immune response triggered by bacterial infection1. They consist of a DNA backbone to which various granule proteins with anti-bacterial properties are bound, including neutrophil elastase and myeloperoxidase2. The primary role of neutrophil ETs (NETs) is to capture pathogens and facilitate their elimination3. However, in addition to the protective role of ETs in immune defense, an increasing number of studies have also discovered a role in disease pathogenesis, particularly during the development of inflammation-driven diseases (i.e., rheumatoid arthritis and atherosclerosis4). The release of ETs can be triggered by various pro-inflammatory cytokines including interleukin 8 (IL-8) and tumor necrosis factor alpha (TNF&#945;)5,6, and the localized accumulation of ETs can increase tissue damage and evoke a pro-inflammatory response7. For example, ETs have been implicated as playing a causal role in the development of atherosclerosis8, promoting thrombosis9, and predicting cardiovascular risk10.
It is now recognized that in addition to neutrophils, other immune cells (i.e., mast cells, eosinophils, and macrophages) can also release ETs on exposure to the microbial or pro- inflammatory stimulation11,12. This may be particularly significant in the case of macrophages, considering their key role in the development, regulation, and resolution of chronic inflammatory diseases. Therefore, it is important to gain a greater understanding of the potential relationship between ET release from macrophages and inflammation-related disease development. Recent studies have shown the presence of METs and NETs in intact human atherosclerotic plaques and organized thrombi13. Similarly, METs have been implicated in driving kidney injury through the regulation of inflammatory responses14. However, in contrast to neutrophils, there are limited data on the mechanisms of MET formation from macrophages. Recent studies using human in vitro models of MET formation show some differences in the pathways involved in each cell type (i.e., regarding the absence of histone citrullination with macrophages)6. However, some have shown that NET release can also occur in the absence of histone citrullination15.
The overall goal of this protocol is to provide a simple and direct method to assess MET release in a clinically relevant macrophage model. There are a number of different in vitro macrophage cell models that have been used to study METs (i.e., the THP-1 human monocyte cell line and various murine macrophage cell lines)16. There are some limitations associated with these models. For example, the differentiation of THP-1 monocytes to macrophages usually requires a priming step, such as the addition of phorbol myristate acetate (PMA), which itself activates protein kinase C (PKC)-dependent pathways. This process is known to trigger ET release4 and results in a low basal MET release from THP-1 cells. Other studies have highlighted some differences in bioactivity and inflammatory responses mounted by macrophages in vivo compared to PMA-treated THP-1 cells17.
Similarly, the behavior and inflammatory responses of different murine macrophage-like cell lines do not completely represent the response spectrum of primary human macrophages18. Therefore, for the purpose of investigating macrophage ET formation in the clinical setting, primary human monocyte-derived macrophages (HMDMs) are believed to be a more relevant model rather than monocytic or murine macrophage-like cell lines.
ET release from M1 polarized HMDMs has been demonstrated following exposure of these cells to a number of different inflammatory stimuli, including the myeloperoxidase-derived oxidant hypochlorous acid (HOCl), PMA, TNF&#945;, and IL-86. Described here is a protocol to polarize HMDMs to the M1 phenotype and visualize subsequent MET release upon exposure to these inflammatory stimuli. PMA is used as a stimulus of MET release to facilitate comparisons to previous studies that have used neutrophils. Importantly, HOCl, IL-8, and TNF&#945; are also used to stimulate MET release, which are believed to be better models of the inflammatory environment in vivo. The microscopic method for visualization of ET release involves staining the extracellular DNA in live cell cultures using SYTOX green, an impermeable fluorescent green nucleic acid stain that has been successfully applied in previous neutrophil studies. This method allows for rapid and qualitative assessment of ET release, but it is not appropriate as a stand-alone method for the quantification of ET release extent. Alternative methodology should be used if quantification is required to compare the extent of ET release resulting from different treatment conditions or interventions.

<table><tbody><tr><td>120Q broad spectrum fluorescent light source</td><td>EXFO Photonic Solutions, Toronto, Canada</td><td></td><td>x-cite series</td></tr><tr><td>Corning CellBIND Multiple Well Plate (12 wells)</td><td>Sigma-Aldrich</td><td>CLS3336</td><td>For cell culture</td></tr><tr><td>Differential Quik Stain Kit (Modified Giemsa)</td><td>Polysciences Inc.</td><td>24606</td><td>Characterisation of monocytes</td></tr><tr><td>Hanks balanced salt solution (HBSS)</td><td>Thermo-Fisher</td><td>14025050</td><td>For washing steps and HOCl treatment</td></tr><tr><td>Hypochlorous acid (HOCl)</td><td>Sigma-Aldrich</td><td>320331</td><td>For MET stimulation</td></tr><tr><td>Interferon gamma</td><td>Thermo-Fisher</td><td>PMC4031</td><td>For M1 priming</td></tr><tr><td>Interleukin 4</td><td>Integrated Sciences</td><td>rhil-4</td><td>For M2 priming</td></tr><tr><td>Interleukin 8</td><td>Miltenyl Biotec</td><td>130-093-943</td><td>For MET stimulation</td></tr><tr><td>L-Glutamine</td><td>Sigma-Aldrich</td><td>59202C</td><td>Added to culture media</td></tr><tr><td>Lipopolysaccharide</td><td>Integrated Sciences</td><td>tlrl-eblps</td><td>For M1 priming</td></tr><tr><td>Lymphoprep</td><td>Axis-Shield PoC AS</td><td>1114544</td><td>For isolation of monocytes</td></tr><tr><td>Olympus IX71 inverted microscope</td><td>Olympus, Tokyo, Japan</td><td></td><td></td></tr><tr><td>Phorbol 12- myristate 13-acetate (PMA)</td><td>Sigma-Aldrich</td><td>P8139</td><td>For MET stimulation</td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>Sigma-Aldrich</td><td>D5652</td><td>For washing steps</td></tr><tr><td>RPMI-1640 media</td><td>Sigma-Aldrich</td><td>R8758</td><td>For cell culture</td></tr><tr><td>SYTOX green</td><td>Life Technologies</td><td>S7020</td><td>For MET visulaization</td></tr><tr><td>TH4-200 brightfield light source</td><td>Olympus, Tokyo, Japan</td><td></td><td>x-cite series</td></tr><tr><td>Tumor necrosis factor alpha</td><td>Lonza</td><td>300-01A-50</td><td>For MET stimulation</td></tr></tbody></table>

in vitro stimulation and visualization of extracellular trap release in differentiated human monocyte-derived macrophages

The release of extracellular traps (ETs) by neutrophils has been identified as a contributing factor to the development of diseases related to chronic inflammation. Neutrophil ETs (NETs) consist of a mesh of DNA, histone proteins, and various granule proteins (i.e., myeloperoxidase, elastase, and cathepsin G). Other immune cells, including macrophages, can also produce ETs; however, to what extent this occurs in vivo and whether macrophage extracellular traps (METs) play a role in pathological mechanisms has not been examined in detail. To better understand the role of METs in inflammatory pathologies, a protocol was developed for visualizing MET release from primary human macrophages in vitro, which can also be exploited in immunofluorescence experiments. This allows further characterization of these structures and their comparison to ETs released from neutrophils. Human monocyte-derived macrophages (HMDM) produce METs upon exposure to different inflammatory stimuli following differentiation to the M1 pro-inflammatory phenotype. The release of METs can be visualized by microscopy using a green fluorescent nucleic acid stain that is impermeant to live cells (e.g., SYTOX green). Use of freshly isolated primary macrophages, such as HMDM, is advantageous in modeling in vivo inflammatory events that are relevant to potential clinical applications. This protocol can also be used to study MET release from human monocyte cell lines (e.g., THP-1) following differentiation into macrophages with phorbol myristate acetate or other macrophage cell lines (e.g., the murine macrophage-like J774A.1 cells).

Brightfield images showing the morphological changes of HMDM in response to stimuli for cell differentiation are shown in Figure 1. M1 polarized macrophages from experiments with HMDM exposed to IFN&#947; and LPS showed an elongated and spindle-like cell shape, as indicated by the black arrows in Figure 1 (middle panel). For comparison, the morphology of the M2 polarized macrophages after exposure of HMDM to IL-4 for 48 h were typically round and flat, as indicated by the black arrows in Figure 1 (far right panel).
The ability of differentiated HMDM phenotypes to release METs was visualized by live cell fluorescence imaging with SYTOX green, as presented in Figure 2. Figure 2A shows the control data obtained from each HMDM phenotype incubated for 24 h in the absence of any pro-inflammatory stimuli. In this case, there was very limited green staining, as was expected, given the cell impermeant nature of this stain. Figure 2B showed positive staining for METs, resulting from the exposure of M1 HMDMs to HOCl, PMA, IL-8, or TNF&#945;. The METs are indicated by the white arrows, shown as green streaks, resulting from the strands of extracellular DNA. With HOCl, in addition to staining extracellular DNA, there was some green staining apparent in the cells. This cellular staining was also observed to some extent with the other stimuli and is believed to reflect loss of membrane integrity resulting from ET-independent cell death as a result of the treatment conditions.
Figure 2B also shows the corresponding experiments performed with M2 HMDMs, which were exposed to IL-4. In this case, there was no release of DNA from the cells, as indicated by the absence of the strands/streaks of extracellular DNA; though, there was some cellular uptake of green fluorescent dye with the HOCl and TNF&#945;. For comparative purposes, Figure 3 shows representative data indicating MET release from THP-1 macrophages exposed to TNF&#945; (50 ng/mL) for 4 h. In this case, it should be noted that a non-polarized population of cells was used, and the THP-1 monocytes were differentiated to macrophages by pre-treatment with PMA (50 ng/mL for 72 h) as described previously22.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60541/60541fig01v2.jpg" /> 
Figure 1: Morphological changes of differentially polarized HMDMs.&#160;Representative brightfield images from non-differentiated and differentiated HMDMs (n &#8805;5). HMDMs were cultured with complete media containing human serum and glutamine for 8 days before priming to the M1 or M2 phenotype upon exposure to IFN&#947; and LPS or IL-4, respectively, for 48 h. Scale bar = 200 &#956;m. Arrows indicate examples of cells showing morphological characteristics of M1 or M2 HMDMs. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60541/60541fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60541/60541fig02v2.jpg" /> 
Figure 2: METs produced by M1 HMDMs following HOCl, PMA, IL-8, and TNF&#945; stimulation.&#160;Representative images of SYTOX green stained HMDMs from (A) Non-stimulated M1 and M2 HMDMs incubated for 24 h in the absence of any inflammatory stimuli were used as the control, demonstrating the absence of MET release. (B) M1 and M2 HMDMs were treated with 1) HOCl (200 &#181;M, 15 min), PMA (25 nM), or IL-8 (50 ng/mL) with incubation for 24 h or 2) TNF&#945; (25 ng/mL) with incubation for 6 h to induce MET release. METs were visualized by the addition of SYTOX green, as indicated by white arrows in the upper panel from M1 HMDMs. No METS were seen in the corresponding experiments with M2 HMDMs. Data are representative of replicate culture wells from n &#8805;3 individual donors. Scale bar = 500 &#956;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60541/60541fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60541/60541fig03v2.jpg" /> 
Figure 3: METs produced by non-polarized THP-1 macrophages following TNF&#945; stimulation.&#160;Representative images of SYTOX green-stained TNP-1 macrophages, which were differentiated by pre-treatment with PMA (50 ng/mL for 72 h) before further incubation for 4 h in the absence or presence of TNF&#945; (50 ng/mL) to induce MET release. METs were visualized by addition of SYTOX green. Data are representative of replicate culture wells from n &#8805; 3 experiments. Scale bar = 100 &#956;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60541/60541fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Vitro Stimulation and Visualization of Extracellular Trap Release in Differentiated Human Monocyte-derived Macrophages at JoVE.com

In Vitro Stimulation and Visualization of Extracellular Trap Release in Differentiated Human Monocyte-derived Macrophages

Presented here is a protocol to detect macrophage extracellular trap (MET) production in live cell culture using microscopy and fluorescence staining. This protocol can be further extended to examine specific MET protein markers by immunofluorescence staining.

The release of extracellular traps (ETs) by neutrophils has been identified as a contributing factor to the development of diseases related to chronic ...

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Universidad San Sebastian

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JoVE Journal

Immunology and Infection

9.5K Views.  Heart Research Institute. Presented here is a protocol to detect macrophage extracellular trap (MET) production in live cell culture using microscopy and fluorescence staining. This protocol can be further extended to examine specific MET protein markers by immunofluorescence staining.

Video: In Vitro Stimulation and Visualization of Extracellular Trap Release in Differentiated Human Monocyte-derived Macrophages

Whole-cell MALDI-TOF Mass Spectrometry is an Accurate and Rapid Method to Analyze Different Modes of Macrophage Activation

MALDI-TOF is an extensively used mass spectrometry technique in chemistry and biochemistry. It has been also applied in medicine to identify molecules and biomarkers. Recently, it has been used in microbiology for the routine identification of bacteria grown from clinical samples, without preparation or fractionation steps. We and others have applied this whole-cell MALDI-TOF mass spectrometry technique successfully to eukaryotic cells. Current applications range from cell type identification to quality control assessment of cell culture and diagnostic applications. Here, we describe its use to explore the various polarization phenotypes of macrophages in response to cytokines or heat-killed bacteria. It allowed the identification of macrophage-specific fingerprints that are representative of the diversity of proteomic responses of macrophages. This application illustrates the accuracy and simplicity of the method. The protocol we described here may be useful for studying the immune host response in pathological conditions or may be extended to wider diagnostic applications.

This protocol describes the use of whole-cell MALDI-TOF mass spectrometry on eukaryotic cells. Here, we illustrate the accuracy of this technique by analyzing the multiple activation states of macrophages in response to their microenvironment.

MALDI-TOF is an extensively used mass spectrometry technique in chemistry and biochemistry. It has been also applied in medicine to identify molecules and ...

Isolation of Human Monocytes by Double Gradient Centrifugation and Their Differentiation to Macrophages in Teflon-coated Cell Culture Bags

Human macrophages are involved in a plethora of pathologic processes ranging from infectious diseases to cancer. Thus they pose a valuable tool to understand the underlying mechanisms of these diseases. We therefore present a straightforward protocol for the isolation of human monocytes from buffy coats, followed by a differentiation procedure which results in high macrophage yields. The technique relies mostly on commonly available lab equipment and thus provides a cost and time effective way to obtain large quantities of human macrophages. Briefly, buffy coats from healthy blood donors are subjected to a double density gradient centrifugation to harvest monocytes from the peripheral blood. These monocytes are then cultured in fluorinated ethylene propylene (FEP) Teflon-coated cell culture bags in the presence of macrophage colony-stimulating factor (M-CSF). The differentiated macrophages can be easily harvested and used for subsequent studies and functional assays. Important methods for quality control and validation of the isolation and differentiation steps will be highlighted within the protocol. In summary, the protocol described here enables scientists to routinely and reproducibly isolate human macrophages without the need for cost intensive tools. Furthermore, disease models can be studied in a syngeneic human system circumventing the use of murine macrophages.

We present a simple and efficient protocol for the generation of human macrophages. Buffy coats are processed by double density gradient centrifugation and isolated monocytes are then differentiated to macrophages in Teflon-coated cell culture bags. This maximizes macrophage yields and facilitates cell harvesting for subsequent experiments.

Human macrophages are involved in a plethora of pathologic processes ranging from infectious diseases to cancer. Thus they pose a valuable tool to ...

Visualizing Macrophage Extracellular Traps Using Confocal Microscopy

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal microscopy methods to visualize macrophage extracellular traps (METs). These extracellular traps are defined by the presence of extracellular chromatin with co-expression of other components such as granule proteases, citrullinated histones, and peptidyl arginase deiminase (PAD). The expression of METs is generally measured after exposure to a stimulus and compared to un-stimulated samples. Samples are also included for background and isotype control. Cells are analyzed using well-defined image analysis software. Confocal microscopy may be used to define the presence of METs both in vitro and in vivo in lung tissue.

Macrophage extracellular traps are a newly described entity. This article will concentrate on confocal microscopy methods and how they are visualized in vitro and in vivo from lung samples.

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal ...

Quantification of Monocyte Chemotactic Activity In Vivo and Characterization of Blood Monocyte Derived Macrophages

Tissue homeostasis and repair are critically dependent on the recruitment of monocyte-derived macrophages. Both under- and over-recruitment of monocyte-derived macrophages can impair wound healing. We showed that high fat and high sugar diets promote monocyte priming and dysfunction, converting healthy blood monocytes into a hyper chemotactic phenotype poised to differentiate into macrophages with dysregulated activation profiles and impaired phenotypic plasticity. The over-recruitment of monocyte-derived macrophages and recruitment of macrophages with dysregulated activation profiles is believed to be a major contributor to the development of chronic inflammatory diseases associated with metabolic disorders, including atherosclerosis and obesity. The goal of this protocol is to quantify the chemotactic activity of blood monocytes as a biomarker for monocyte priming and dysfunction and to characterize the macrophage phenotype blood monocytes are poised to differentiate into in these mouse models. Using single cell Western blot analysis, we show that after 24 h 33%of cells recruited into MCP-1-loaded basement membrane-derived gel plugs injected into mice are monocytes and macrophages; 58% after day 3. However, on day 5, monocyte and macrophage numbers were significantly decreased. Finally, we show that this assays also allows for the isolation of live macrophages from the surgically retrieved basement membrane-derived gel plugs, which can then be subjected to subsequent characterization by single cell Western blot analysis.

Here we present a protocol to quantify the chemotactic activity of blood monocytes in mouse models, to assess the effects of nutritional, pharmacological and genetic interventions on blood monocyte and to characterize the blood monocytes derived macrophages in mouse models using monocyte-chemoattractant protein-1 (MCP-1)-loaded basement membrane-derived gel plugs.

Tissue homeostasis and repair are critically dependent on the recruitment of monocyte-derived macrophages. Both under- and over-recruitment of ...

Visualization of Inflammatory Caspases Induced Proximity in Human Monocyte-Derived Macrophages

Inflammatory caspases include caspase-1, -4, -5, -11, and -12 and belong to the subgroup of initiator caspases. Caspase-1 is required to ensure correct regulation of inflammatory signaling and is activated by proximity-induced dimerization following recruitment to inflammasomes. Caspase-1 is abundant in the monocytic cell lineage and induces maturation of the pro-inflammatory cytokines interleukin (IL)-1&#946; and IL-18 to active secreted molecules. The other inflammatory caspases, caspase-4 and -5 (and their murine homolog caspase-11) promote IL-1&#946; release by inducing pyroptosis. Caspase Bimolecular Fluorescence Complementation (BiFC) is a tool used to measure inflammatory caspase induced proximity as a readout of caspase activation. The caspase-1, -4, or -5 prodomain, which contains the region that binds to the inflammasome, is fused to non-fluorescent fragments of the yellow fluorescent protein Venus (Venus-N [VN] or Venus-C [VC]) that associate to reform the fluorescent Venus complex when the caspases undergo induced proximity. This protocol describes how to introduce these reporters into primary human monocyte-derived macrophages (MDM) using nucleofection, treat the cells to induce inflammatory caspase activation, and measure caspase activation using fluorescence and confocal microscopy. The advantage of this approach is that it can be used to identify the components, requirements, and localization of the inflammatory caspase activation complex in living cells. However, careful controls need to be considered to avoid compromising cell viability and behavior. This technique is a powerful tool for the analysis of dynamic caspase interactions at the inflammasome level as well as for the interrogation of the inflammatory signaling cascades in living MDM and monocytes derived from human blood samples.

This protocol describes the workflow to obtain monocytes-derived macrophages (MDM) from human blood samples, a simple method to efficiently introduce inflammatory caspase Bimolecular Fluorescence Complementation (BiFC) reporters into human MDM without compromising cell viability and behavior, and an imaging-based approach to measure inflammatory caspase activation in living cells.

Inflammatory caspases include caspase-1, -4, -5, -11, and -12 and belong to the subgroup of initiator caspases. Caspase-1 is required to ensure correct ...

Reprograming Model of Human Monocyte-derived Macrophages for In-vitro Assays

Cells of the monocyte-macrophage lineage are multifunctional and found in almost all body tissues. They coordinate innate and adaptive immunity&#180;s initiation and resolution phases, significantly affecting protective immunity and immune-mediated pathological injury. While tissue-resident macrophages are key players in maintaining homeostasis in a steady state, large amounts of monocytes are recruited from the peripheral blood into the tissue following damage or inflammatory insults. Monocyte-derived macrophages (M-DM) can differentiate into many dynamic subtypes, and their phenotypes and functions depend on the local tissue environment. 
To compare different stimuli or environmental conditions during M-DM differentiation and polarization, we standardized an in-vitro model of human nonpolarized M-DM M0 and some cardinal cytokine-polarized macrophages, IFN&#947;/LPS-derived M1, IL-4-derived M2a, and IL-10 or dexamethasone-derived M2c, to analyze skewing reprogramming of M-DM by flow cytometry and real-time PCR. We found that CD64, CD206, CD163, CD14, and MERTK can clearly discriminate unpolarized M0 and polarized M1, M2a, and M2c by flow cytometry. Moreover, we defined IRF1 and CXCL10 as specific genes for classical IFN&#947;/LPS-derived M1-, IRF4-, CCL22-, and TGM2-specific transcripts for IL-4-derived M2a, and the MERTK gene for dexamethasone-derived M2c. To summarize, our standardized M-DM protocol could give the cardinal in-vitro map to analyze the differentiation and polarization of human M-DM under diverse stimuli.

Monocytes and macrophages are very plastic and reprogrammable immune cells crucial for health and disease. They sense and respond to a broad range of stimuli by adopting specific differentiation programs and phenotypes. We have standardized an in-vitro model to study macrophage polarization and reprogramming, providing a valuable tool for research.

Cells of the monocyte-macrophage lineage are multifunctional and found in almost all body tissues. They coordinate innate and adaptive immunity&#180;s ...

In Vitro Culture and Differentiation of Primary Monocytes into Macrophages

Source: Plaisance-Bonstaff, K., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/59967">Isolation, Transfection, and Culture of Primary Human Monocytes.</a> J. Vis. Exp. (2019)
This video demonstrates the priming of primary monocytes into macrophages. Primary human monocytes can be isolated and differentiated to an M1-like pro-inflammatory phenotype using the granulocyte-macrophage colony-stimulating factor (GM-CSF) or to an M2-like anti-inflammatory phenotype using the macrophage colony-stimulating factor (M-CSF).

Source: Plaisance-Bonstaff, K., et al. Isolation, Transfection, and Culture of Primary Human Monocytes. J. Vis. Exp. (2019)
This video demonstrates the ...

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Immunology

Immune Systems and Components

Cell Differentiation Techniques

Visualization of Caspase Activation in Human Monocyte-Derived Macrophages

Source: Bol&#237;var, B. E., et. al., <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/63162">Visualization of Inflammatory Caspases Induced Proximity in Human Monocyte-Derived Macrophages.</a> J. Vis. Exp. (2022).
This video demonstrates the activation of caspase proteins in human macrophages using bimolecular fluorescence complementation reporter proteins. The pro-domains of caspase-1 are fused with non-fluorescent fragments of Venus proteins, which are recruited to inflammasome complexes. The proximity of inflammasomes induces the refolding and fluorescence of Venus fragments, confirming caspase activation in the macrophages.

Source: Bol&#237;var, B. E., et. al., Visualization of Inflammatory Caspases Induced Proximity in Human Monocyte-Derived Macrophages. J. Vis. Exp. 
...

Immune Response

Immune Modulation

In Vitro Stimulation and Visualization of Macrophage Extracellular Traps

Source: Zhang, Y., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/60541">In Vitro Stimulation and Visualization of Extracellular Trap Release in Differentiated Human Monocyte-derived Macrophages.</a> J. Vis. Exp. (2019).
This video demonstrates in vitro stimulation of macrophage extracellular traps (METs) in human monocyte-derive macrophages using different inflammatory stimuli such as TNF-alpha, PMA, and HOCL. The video also demonstrates the method of SYTOX dye staining of METs for visualization.

1. Polarization of HMDM

	Under sterile conditions, prepare the M1 priming media by adding interferon &#947; (IFN&#947;; 20 ng/mL) and lipopolysaccharide ...

Immunodiagnostics

Imaging and Visualization Techniques

Characterization of Macrophage Extracellular Traps in Mouse Lung Tissue Samples

Source: Sharma, R., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/56459">Visualizing Macrophage Extracellular Traps Using Confocal Microscopy.</a> J. Vis. Exp. (2017)
In this video, we describe a protocol for the immunostaining and confocal microscopy-based visualization of macrophage extracellular traps (METs) in murine lung tissue. Tissue-resident macrophages are characterized by the expression of the macrophage-specific F4/80 surface glycoprotein marker, while METs are visualized by labeling the MET markers matrix metalloproteinase 9, citrullinated histone H3, and extracellular chromatin.

Source: Sharma, R., et al. Visualizing Macrophage Extracellular Traps Using Confocal Microscopy. J. Vis. Exp. (2017)
In this video, we describe a protocol ...