We thank Mrs. Pia Stier for excellent technical assistance. M. von Lilienfeld-Toal is supported by the Center for Sepsis Control and Care (German Federal Ministry of Education and Health, BMBF, FKZ 01E01002) and InfectoGnostics Research Campus (BMBF, FKZ 13GW0096D). Thi Ngoc Mai Hoang is supported by the Jena School of Microbial Communication (Deutsche Forschungsgemeinschaft, FKZ 214/2)

This protocol includes the use of human buffy coats obtained from the Institute for Transfusion Medicine, Jena University Hospital and fresh venous blood drawn from patients, both after written informed consent of the donors in accordance to ethics committee approval 4357-03/15.
1. Preparation of Aspergillus fumigatus Conidia
<ol>
	<li>Grow A. fumigatus on 1.5% malt agar Petri dishes for 5 days at 37 &#176;C without CO2. 
	CAUTION: A. fumigatus is a biosafety level 2 microorganism and must be handled in an appropriate facility using a biosafety cabinet and wearing lab coat, gloves and a filter mask. 
	NOTE: The plate should be covered completely with a green-greyish layer of conidia. White cultures do not sporulate. Composition of malt agar: 4% (w/v) malt extract, yeast extract 0.4% (w/v), agar 1.5% (w/v), Aqua dest.</li>
	<li>Harvesting conidia
	<ol>
		<li>Place a paper towel wet with disinfectant into the biosafety cabinet and put the plate on top to prevent overdistribution of volatile conidia.</li>
		<li>Add 10 mL of phosphate buffered saline (PBS) + 0.01% detergent on top of the fungus, use a Drigalski spatula to spread the liquid over the plate and rub off the dark colored conidia. Be careful not to remove the white mycelium.</li>
		<li>Put the conidia suspension to a 50 mL tube using a 30 &#181;m cell strainer to remove any residual mycelium.</li>
		<li>Repeat steps 1.2.2 and 1.2.3 and collect in the same 50 mL tube.</li>
		<li>Spin for 5 min at 2,600 x g at room temperature.</li>
		<li>Remove the supernatant and resuspend in 20 mL of sterile Aqua dest.</li>
		<li>Determine conidia concentration with a Thoma counting chamber. 
		NOTE: The protocol can be paused here and the conidia suspension stored for up to 1 month at 4 &#176;C with the lid screwed tightly.</li>
	</ol>
	</li>
	<li>FITC labeling
	<ol>
		<li>Prepare a 0.1 mM solution of FITC powder in sterile 0.1 M Na2CO3 (dissolved in PBS). 
		CAUTION: FITC powder is hazardous and should be handled with gloves, goggles and a filter mask. Collect waste according to local regulations. 
		NOTE: Omit artificial light during this and the following steps involving conidia.</li>
		<li>Resuspend 1 x 108 (or less) conidia in 5 mL of FITC solution in a 15 mL tube. Incubate for 20 min at 37 &#176;C in a rotator.</li>
		<li>For the negative control, resuspend conidia in 0.1 M Na2CO3 (without FITC) in a 15 mL tube. Incubate for 20 min at 37 &#176;C in a rotator. 
		NOTE: When calculating the amount of conidia to be stained, take into account a loss of up to 70% during staining, swelling and all necessary washing steps. If no rotator is available, the suspension should be shaken three times during incubation.</li>
		<li>For washing, add 10 mL of PBS + 0.01% detergent to the suspension and spin for 5 min at 2,600 x g at room temperature.</li>
		<li>Remove supernatant and repeat washing twice with 10 mL of PBS + 0.01% detergent.</li>
	</ol>
	</li>
	<li>Swelling of conidia (may be omitted if not desired)
	<ol>
		<li>Resuspend FITC labeled conidia in 5 mL of Roswell Park Memorial Institute (RPMI) medium + 10% Fetal Calf Serum (FCS) and incubate in a rotator at 37 &#176;C for the desired time (e.g., 2 h, 4 h). 
		NOTE: If no rotator is available, the suspension should be shaken at least every 20 min during incubation.</li>
		<li>Add 10 mL of PBS + 0.01% detergent to the suspension and spin for 5 min at 2,600 x g at room temperature.</li>
		<li>Remove the supernatant and wash twice with 10 mL of PBS + 0.01% detergent.</li>
		<li>Filter through a 30 &#181;m cell strainer to remove large clumps of conidia.</li>
	</ol>
	</li>
	<li>Fixing of conidia (may be omitted if not desired)
	<ol>
		<li>Resuspend FITC labeled and swollen conidia in 1 mL of formaldehyde and incubate for 1 h at room temperature.</li>
		<li>Add 10 mL of PBS + 0.01% detergent to the suspension and spin for 5 min at 2,600 x g at room temperature.</li>
		<li>Remove the supernatant and wash twice with 10 mL of PBS + 0.01% detergent. 
		NOTE: The protocol can be paused here and the conidia stored in PBS in the dark (tube wrapped up in aluminum foil) for up to 1 week at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Preparation of Human Primary Leukocytes
<ol>
	<li>Put 5 mL of buffy coat in a 50 mL tube. Alternatively, use fresh human peripheral venous blood drawn into ethylenediaminetetraacetic acid (EDTA) monovettes (10 mL per 50 mL tube). 
	CAUTION: Human blood may transmit viruses such as Human Immunodeficiency Virus and Hepatitis B Virus. Handle only after being vaccinated against Hepatitis B. Handle in a biosafety cabinet wearing lab coat and gloves.</li>
	<li>Fill up the tube with Erythrocyte Lysis (EL) Buffer, invert three times and incubate for 5-8 min horizontally until the milky appearance of the mixture turns clear. 
	NOTE: Occasionally, the blood may take longer to lyse. Go by the appearance. It is not unusual that two tubes of the same blood take different times to lyse. Composition of EL buffer: 0.15 M NH4Cl, 10 mM KHCO3, 0.1 mM EDTA</li>
	<li>Spin for 10 min at 300 x g at room temperature. Discard the supernatant.</li>
	<li>Resuspend the pellet in 1 mL of EL Buffer by pipetting. Then add another 24 mL of EL buffer, and invert several times.</li>
	<li>Spin for 5 min at 300 x g at room temperature.</li>
	<li>Discard the supernatant and resuspend cells in 1 mL of RPMI + 10% FCS.</li>
	<li>Determine the cell concentration with a Neubauer counting chamber.</li>
</ol>
3. Phagocytosis Assay
<ol>
	<li>Incubate 2 x 106 leukocytes and 4 x 106 FITC-labeled conidia (multiplicity of infection = 2) in 1.5 mL of RPMI + 10% FCS in a 12-well cell culture plate. As controls, include cells only (no conidia) and cells + unlabeled conidia.</li>
	<li>Place in a humidified CO2 incubator at 37 &#176;C and incubate for the desired period of time (e.g., 0.5 h, 2 h or 4 h).</li>
	<li>After incubation, harvest cells with a cell scraper and put in a 15 mL tube.</li>
	<li>Spin for 5 min at 300 x g at room temperature.</li>
	<li>Collect supernatant for cytokine analysis or discard if not wanted. Resuspend each sample in 100 &#181;L of PBS + 2 mM EDTA.</li>
</ol>
4. Antibody Staining
<ol>
	<li>For each sample, prepare 100 &#181;L of antibody mix, including APC anti-FITC antibody, according to Table 1.</li>
	<li>Put a 100 &#181;L sample into one well of a 96-well V-bottom plate. Add 150 &#181;L of PBS + 2 mM EDTA for washing.</li>
	<li>For color compensation, place 1 x 106 cells for each color in further wells of the 96-well V bottom plate. Include a well of cells that is left unstained. Add 150 &#181;L of PBS + 2 mM EDTA for washing.</li>
	<li>Cover plate with an adhesive foil.</li>
	<li>Spin for 5 min at 300 x g at room temperature. Remove the foil.</li>
	<li>Discard the supernatant by quickly and forcefully inverting the plate only once over the sink or a disposable paper towel. 
	NOTE: Do not repeat or knock the plate on a paper until dry as this will result in a massive loss of cells from the plate.</li>
	<li>Resuspend cells in the 100 &#181;L antibody mix, and mix well by pipetting.</li>
	<li>For color compensation, resuspend the respective cells in 100 &#181;L of PBS + 2 mM EDTA and add a single antibody to each well at the same amount used in the antibody mix.</li>
	<li>Cover with an adhesive foil and incubate for 20 min at room temperature in the dark.</li>
	<li>Remove the foil. Add 150 &#181;L of PBS + 2 mM EDTA to each well for washing. Cover with an adhesive foil.</li>
	<li>Spin for 5 min at 300 x g at room temperature. Remove the foil. Discard the supernatant by quickly and forcefully inverting the plate over the sink or a disposable paper towel.</li>
	<li>Resuspend in 200 &#181;L of PBS + 2 mM EDTA and transfer cells from each well to a separate round bottom tube. Make sure there are no cell clusters in the suspension. Remove clusters otherwise. 
	NOTE: Every cluster that is large enough for the eye to see is large enough to potentially clog the cytometer.</li>
</ol>
5. Flow Cytometry
<ol>
	<li>Start the flow cytometer and let it warm up. Start the acquisition software.</li>
	<li>Create a new experiment and setup and label the samples.</li>
	<li>Set up parameters (FSC 250, SSC 250) and detectors for fluorophores FITC, APC, BUV395, V500, PerCP-Cy5.5.</li>
	<li>Compensation setup
	<ol>
		<li>Open compensation setup.</li>
		<li>Indicate individual colors.</li>
		<li>Using the control cells left unstained or with individual stainings, set the PMT detector voltages to include all events within the scale.</li>
		<li>Record at least 10,000 events of each control.</li>
		<li>Use the compensation setup to calculate the spillover of fluorophores and apply to the experiment&#39;s cytometer settings.</li>
	</ol>
	</li>
	<li>Recording sample data
	<ol>
		<li>Display FSC and SSC in the acquisition software and set gate around leukocytes.</li>
		<li>Based on the leukocyte gate, display dot plot SSC/CD45 and gate for CD45+ cells to separate from conidia.</li>
		<li>Display CD45+ cells in a dot plot CD14/CD66b and gate monocytes (CD14+) and neutrophils (CD66b+) separately.</li>
		<li>Display neutrophils in a dot plot anti-FITC/FITC.</li>
		<li>Using the sample with unlabeled conidia, set quadrants for anti-FITC and FITC signals, allowing a maximum of 1% of cells in the respective quadrants.</li>
		<li>Repeat steps 5.5.4 and 5.5.5 for monocyte gate.</li>
		<li>Record all samples with at least 20,000 events in the leukocyte gate.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Brown, G. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hidden+killers:+human+fungal+infections.">Hidden killers: human fungal infections.</a> Science Translational Medicine. 4 (165), 113 (2012).</li><li>Brakhage, A. A., Bruns, S., Thywissen, A., Zipfel, P. F., Behnsen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interaction+of+phagocytes+with+filamentous+fungi.">Interaction of phagocytes with filamentous fungi.</a> Current Opinion in Microbiology. 13 (4), 409-415 (2010).</li><li>Heinekamp, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interference+of+Aspergillus+fumigatus+with+the+immune+response.">Interference of Aspergillus fumigatus with the immune response.</a> Seminars in Immunopathology. 37 (2), 141-152 (2015).</li><li>Slesiona, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Persistence+versus+escape:+Aspergillus+terreus+and+Aspergillus+fumigatus+employ+different+strategies+during+interactions+with+macrophages.">Persistence versus escape: Aspergillus terreus and Aspergillus fumigatus employ different strategies during interactions with macrophages.</a> PLoS One. 7 (2), 31223 (2012).</li><li>Busetto, S., Trevisan, E., Patriarca, P., Menegazzi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+single-step,+sensitive+flow+cytofluorometric+assay+for+the+simultaneous+assessment+of+membrane-bound+and+ingested+Candida+albicans+in+phagocytosing+neutrophils.">A single-step, sensitive flow cytofluorometric assay for the simultaneous assessment of membrane-bound and ingested Candida albicans in phagocytosing neutrophils.</a> Cytometry A. 58 (2), 201-206 (2004).</li><li>Lowe, D. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+assay+of+antimycobacterial+activity+and+phagocytosis+by+human+neutrophils.">A novel assay of antimycobacterial activity and phagocytosis by human neutrophils.</a> Tuberculosis (Edinb). 93 (2), 167-178 (2013).</li><li>Nuutila, J., Lilius, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+cytometric+quantitative+determination+of+ingestion+by+phagocytes+needs+the+distinguishing+of+overlapping+populations+of+binding+and+ingesting+cells.">Flow cytometric quantitative determination of ingestion by phagocytes needs the distinguishing of overlapping populations of binding and ingesting cells.</a> Cytometry A. 65 (2), 93-102 (2005).</li><li>Saresella, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rapid+evaluation+of+phagocytosis+and+killing+of+Candida+albicans+by+CD13++leukocytes.">A rapid evaluation of phagocytosis and killing of Candida albicans by CD13+ leukocytes.</a> Journal of Immunological Methods. 210 (2), 227-234 (1997).</li><li>Fattorossi, A., Nisini, R., Pizzolo, J. G., D'Amelio, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New,+simple+flow+cytometry+technique+to+discriminate+between+internalized+and+membrane-bound+particles+in+phagocytosis.">New, simple flow cytometry technique to discriminate between internalized and membrane-bound particles in phagocytosis.</a> Cytometry. 10 (3), 320-325 (1989).</li><li>Heinzelmann, M., Gardner, S. A., Mercer-Jones, M., Roll, A. J., Polk, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+phagocytosis+in+human+neutrophils+by+flow+cytometry.">Quantification of phagocytosis in human neutrophils by flow cytometry.</a> Microbiology and Immunology. 43 (6), 505-512 (1999).</li><li>Perticarari, S., Presani, G., Mangiarotti, M. A., Banfi, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+flow+cytometric+method+to+measure+phagocytosis+and+oxidative+products+by+neutrophils.">Simultaneous flow cytometric method to measure phagocytosis and oxidative products by neutrophils.</a> Cytometry. 12 (7), 687-693 (1991).</li><li>Sveum, R. J., Chused, T. M., Frank, M. M., Brown, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+quantitative+fluorescent+method+for+measurement+of+bacterial+adherence+and+phagocytosis.">A quantitative fluorescent method for measurement of bacterial adherence and phagocytosis.</a> Journal of Immunolological Methods. 90 (2), 257-264 (1986).</li><li>de Boer, E. C., Bevers, R. F., Kurth, K. H., Schamhart, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Double+fluorescent+flow+cytometric+assessment+of+bacterial+internalization+and+binding+by+epithelial+cells.">Double fluorescent flow cytometric assessment of bacterial internalization and binding by epithelial cells.</a> Cytometry. 25 (4), 381-387 (1996).</li><li>Hartung, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+and+Quantitative+Evaluation+of+Human+Leukocyte+Interaction+with+Aspergillus+fumigatus+Conidia+by+Flow+Cytometry.">Fast and Quantitative Evaluation of Human Leukocyte Interaction with Aspergillus fumigatus Conidia by Flow Cytometry.</a> Cytometry A. 95 (3), 332-338 (2019).</li><li>Fei, C., Lillico, D. M. E., Hall, B., Rieger, A. M., Stafford, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Connected+component+masking+accurately+identifies+the+ratio+of+phagocytosed+and+cell-bound+particles+in+individual+cells+by+imaging+flow+cytometry.">Connected component masking accurately identifies the ratio of phagocytosed and cell-bound particles in individual cells by imaging flow cytometry.</a> Cytometry A. 91 (4), 372-381 (2017).</li></ol>

The authors have nothing to disclose.

This protocol presents a fast flow cytometric method to measure interaction of A. fumigatus conidia with a large number of primary human leukocytes that is not possible in common microscopic protocols. Imaging cells with a microscope and manually counting internalized conidia is cumbersome and can realistically be done for a few hundred cells only. Flow cytometry overcomes this problem by measuring thousands of cells within minutes. A hurdle common to both approaches is the distinction of phagocytosed and adherent conidia in or on cells, respectively. In microscopy, the dye calcofluor white is often used for staining adherent conidia but its usage is limited to microorganisms with a chitin-containing cell wall.
This protocol in contrast used distinct fluorophores for characterization of interaction events that allows the addition of further cell markers, such as the lineage markers CD45, CD14 and CD66b. Thus, it is also possible to discriminate phagocytosis of pathogens by monocytes and neutrophils in a single sample. While the choice of markers and fluorophores for cell identification can be adapted to the needs of the experiment and the capacities of the available cytometer, the usage of the specific APC anti-FITC antibody mentioned in this protocol is recommended as it was the most reliable antibody in our hands.
Since formaldehyde-fixed conidia are not internalized equally well, native spores are recommended for phagocytosis assays. However, native conidia will start or continue swelling during the co-incubation with human leukocytes and eventually germinate. Typically, A. fumigatus conidia germinate after about 8-9 h in glucose-containing media at 37 &deg;C. The combination of swelling time and co-incubation time with phagocytes should not exceed this time frame as germination causes the loss of FITC on the conidia surface. More important, germlings cannot be phagocytosed anymore by monocytes or neutrophils. Instead, these phagocytes accumulate around and stick to germlings that produce cell clusters that clog the cytometer, if not removed. Similarly, 4 h swollen conidia tend to generate clusters among themselves and with cells. Often these clusters cannot be separated mechanically anymore and the cells within are lost for flow cytometric analysis.
Although gating is straightforward and easy in the beginning, the more conidia are internalized by cells, the blurrier gating may become. Using MOIs &gt; 2 increases phagocytosis at initial time points but gating issues might arise earlier as well. Therefore, MOIs should be carefully determined with the specific cells and pathogens of interest.
A limitation of this protocol is in the duality of the cell population with adherent conidia (FITC+ APC+) that might also be harboring cells with adherent as well as internalized conidia. A possibility for further discrimination is the application of imaging flow cytometry15 that allows visual imaging of all cells measured in the flow cytometer.
Due to the unspecific FITC labeling of the conidial cell wall, this method is easily transferable to other fungi of interest such as clinically relevant molds of the genus Mucorales or the yeast Candida albicans. Moreover, also cell-wall bearing bacteria can be FITC-labeled. The universal counterstaining with the anti-FITC antibody allows fast and easy measurement of phagocytosis of all these pathogens alike by a large number of human leukocytes.

Invasive pulmonary aspergillosis is a great threat to immunocompromised patients as treatment options are limited and only successful upon early diagnosis, which leads to high mortality rates1. Infectious agents are conidia (spores) of the mold Aspergillus fumigatus that are ubiquitous to most habitats2. Conidia are inhaled, pass through the airways and can finally enter the lung alveoli. In immunocompetent humans, these conidia are cleared by innate immune cells such as monocytes or macrophages and neutrophil granulocytes, which take up (phagocytose) and digest the pathogens3. Phagocytosis is important for microbiologists and immunologists likewise when interested in host-pathogen interactions. Confrontation assays, such as co-incubation of leukocytes and conidia, often include labeling of the spores by fluorescein or its derivative fluorescein isothiocyanate (FITC). Using a microscope, it is straightforward to identify internalized fluorescent conidia and to determine attached/adherent conidia, although this approach is cumbersome and realistically restricted to a few hundreds of cells4. However, in flow cytometry which easily allows the analysis of hundreds of thousands of cells within minutes, differential staining of phagocytosed and adherent conidia is vital. Therefore, many protocols rely on Trypan Blue to quench FITC-fluorescence from adherent conidia5,6,7,8. Another approach is exploiting fluorescence resonance energy transfer of ethidium bromide and FITC to emit red instead of green fluorescence from adherent conidia9,10,11. If specific antibodies are available, as is the case for some bacteria, cell-bound particles can be directly stained12,13.
Here, we present a protocol to quickly and quantitatively assess phagocytosis of FITC-labeled A. fumigatus conidia by human leukocytes along with attachment of spores to cells and lack of interaction by employing an allophycocyanin (APC)-coupled anti-FITC antibody. The method also allows for the simultaneous flow cytometric analysis of further cell markers that can be employed for separate analysis of phagocytosis by monocytes and neutrophils from the same sample.
The protocol can be applied for characterization of fungal strains (e.g., several species of Aspergillus and other molds from the genus Mucorales presented here) and their mutants14 and immunological research on phagocytes, such as leukocytes from immunocompromised individuals.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Adhesive foil</td>
			<td>Brand</td>
			<td>701367</td>
			<td></td>
		</tr>
		<tr>
			<td>anti-CD14 V500</td>
			<td>BD Biosciences</td>
			<td>561391</td>
			<td>clone M5E2</td>
		</tr>
		<tr>
			<td>anti-CD45 BUV395</td>
			<td>BD Biosciences</td>
			<td>563792</td>
			<td>clone HI30</td>
		</tr>
		<tr>
			<td>anti-CD66b PerCP-Cy5.5</td>
			<td>BD Biosciences</td>
			<td>562254</td>
			<td>clone G10F5</td>
		</tr>
		<tr>
			<td>anti-FITC APC</td>
			<td>ThermoFisher Scientific</td>
			<td>17-7691-82</td>
			<td>clone NAWESLEE</td>
		</tr>
		<tr>
			<td>Cell culture plate, 12-well</td>
			<td>Greiner Bio-one</td>
			<td>665180</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell scraper</td>
			<td>Bioswisstech</td>
			<td>800020</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell strainer, 30 &#181;m</td>
			<td>Miltenyi Biotech</td>
			<td>130-098-458</td>
			<td>SmartStrainer</td>
		</tr>
		<tr>
			<td>Cytometer</td>
			<td>BD Biosciences</td>
			<td></td>
			<td>LSR Fortessa II, lasers: 488 nm (blue), 405 nm (violet), 355 nm (UV) and 640 nm (red)</td>
		</tr>
		<tr>
			<td>Detergent</td>
			<td>Sigma Aldrich</td>
			<td>P1379</td>
			<td>Tween 20, 0.01% in PBS</td>
		</tr>
		<tr>
			<td>Drigalski spatula</td>
			<td>Carl Roth</td>
			<td>PC59.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid (EDTA)</td>
			<td>Sigma Aldrich</td>
			<td>ED3SS-500g</td>
			<td>2 mM in PBS</td>
		</tr>
		<tr>
			<td>Erythrocyte lysis buffer</td>
			<td></td>
			<td></td>
			<td>0.15 M NH4Cl, 10 mM KHCO3, 0.1 mM EDTA</td>
		</tr>
		<tr>
			<td>Fetal Calf Serum (FCS)</td>
			<td>Biochrom AG</td>
			<td>S 0115</td>
			<td>10% in RPMI 1640</td>
		</tr>
		<tr>
			<td>Fluorescein isothiocyanate (FITC)</td>
			<td>Sigma Aldrich</td>
			<td>F3651-100MG</td>
			<td>0.1 mM in Na2CO3 /PBS solution</td>
		</tr>
		<tr>
			<td>Formaldehyd</td>
			<td>Carl Roth</td>
			<td>PO87.3</td>
			<td>Histofix</td>
		</tr>
		<tr>
			<td>Malt agar (1.5%)</td>
			<td></td>
			<td></td>
			<td>malt extract (40 g), yeast extract (4 g), agar (15 g), Aqua dest. (1 L), adjust pH to 5.7-6.0, sterilise at 121 &#176;C for 35 minutes</td>
		</tr>
		<tr>
			<td>Na2CO3</td>
			<td>Carl Roth</td>
			<td>8563.1</td>
			<td>0.1 M in PBS</td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Greiner Bio-one</td>
			<td>633180</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphat Buffered Saline (PBS)</td>
			<td>ThermoFisher Scientific</td>
			<td>189012-014</td>
			<td>without Calcium, without Magnesium</td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>ThermoFisher Scientific</td>
			<td>61870010</td>
			<td>RPMI 1640 Medium, GlutaMAX Supplement</td>
		</tr>
		<tr>
			<td>Rotator</td>
			<td>Miltenyi Biotech</td>
			<td>130-090-753</td>
			<td>MACSmix Tube Rotator</td>
		</tr>
		<tr>
			<td>Round-bottom tube, 7.5 mL</td>
			<td>Corning</td>
			<td>REF 352008</td>
			<td></td>
		</tr>
		<tr>
			<td>Software for data acquisition and analysis</td>
			<td>BD Biosciences</td>
			<td></td>
			<td>FACSDiva 8.0</td>
		</tr>
		<tr>
			<td>V-bottom plate, 96 well</td>
			<td>Brand</td>
			<td>781601</td>
			<td>untreated surface</td>
		</tr>
	</tbody>
</table>

measuring phagocytosis of aspergillus fumigatus conidia by human leukocytes using flow cytometry

Invasive pulmonary infection by the mold Aspergillus fumigatus poses a great threat to immunocompromised patients. Inhaled fungal conidia (spores) are cleared from the human lung alveoli by being phagocytosed by innate monocytes and/or neutrophils. This protocol offers a fast and reliable measurement of phagocytosis by flow cytometry using fluorescein isothiocyanate (FITC)-labeled conidia for co-incubation with human leukocytes and subsequent counterstaining with an anti-FITC antibody to allow discrimination of internalized and cell-adherent conidia. Major advantages of this protocol are its rapidness, the possibility to combine the assay with cytometric analysis of other cell markers of interest, the simultaneous analysis of monocytes and neutrophils from a single sample and its applicability to other cell wall-bearing fungi or bacteria. Determination of percentages of phagocytosing leukocytes provides a means to microbiologists for evaluating virulence of a pathogen or for comparing pathogen wildtypes and mutants as well as to immunologists for investigating human leukocyte capabilities to combat pathogens.

In measuring phagocytosis of A. fumigatus conidia by human phagocytic cells, discrimination between genuine internalization and mere attachment of conidia to the cells is an obstacle, especially when it comes to high-throughput methods such as flow cytometry. In order to overcome this hurdle, we present a fast and reliable protocol based on the staining of conidia with the fluorescent dye FITC prior to co-incubation of cells and conidia, followed by a counterstaining with an APC-labeled anti-FITC antibody after incubation (Figure 1A). As shown in Figure 1B, FITC-labeled conidia are phagocytosed by human monocytes and neutrophils that provide a green signal to the cells. These conidia are inaccessible for the anti-FITC antibody and, hence, cannot bind the antibody and the cells appear APC negative (FITC+, APC-). Non-interacting cells do not acquire a green signal from FITC-labeled conidia and remain FITC-, APC-. A few cells appear FITC-, APC+. Since the anti-FITC APC antibody should not be able to bind cells without FITC-labeled conidia, these events are considered staining artifacts. FITC-labeled conidia, which are attached to the cells but not internalized, render cells also positive for FITC but also provide a target for the anti-FITC antibody that makes these cells double positive for FITC and APC (FITC+, APC+). When analyzed microscopically, this population contained up to 20% of cells with attached conidia only in our experiments.
Using the antibodies described in this protocol and following the gating strategy in Figure 1D, a general gating of human leukocytes by FSC and SSC characteristics is followed by a separation of leukocytes and free conidia by the pan-leukocyte marker CD45. Especially when using swollen conidia and/or long incubation times, conidia can reach almost cell size at the time of flow cytometry and hence bias analysis. Since human primary monocytes and neutrophils take up conidia differently, this protocol allows to separately analyze these cell population based on staining with the well-established cell lineage markers CD14 for monocytes and CD66b for neutrophils. Gating for phagocytosing and adherent cell populations is done based on control samples with unlabeled conidia that carry neither a FITC nor an anti-FITC APC signal. When APC and FITC are plotted against each other, quadrants are set in such a manner that a maximum of 1% cells are allowed in the gates of interest.
The percentage of human primary phagocytes internalizing conidia can be highly variable among blood donors but also depends on experimental factors such as incubation time and swelling state of conidia. Spore internalization can be detected after 0.5 h of co-incubation already and increases with time (Figure 2A). Pre-swollen conidia are taken up easier than resting (not swollen) spores even at short incubation times (Figure 2B). If conidia are fixed with formaldehyde, phagocytosis is diminished compared to native conidia (Figure 2C).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>reagent</td>
			<td>&#181;l per sample</td>
		</tr>
		<tr>
			<td>CD45 BUV395</td>
			<td>1</td>
		</tr>
		<tr>
			<td>CD14 V500</td>
			<td>0.5</td>
		</tr>
		<tr>
			<td>CD66b PerCP-Cy5.5</td>
			<td>1.5</td>
		</tr>
		<tr>
			<td>anti-FITC APC</td>
			<td>0.5</td>
		</tr>
		<tr>
			<td>PBS + 2 mM EDTA</td>
			<td>97</td>
		</tr>
		<tr>
			<td>total</td>
			<td>100</td>
		</tr>
	</tbody>
</table>
Table 1: Antibody mix for flow cytometry. Amounts of each reagent are given as microliters per sample (1 x 106 cells) to be analyzed.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60397/60397fig1a.jpg" /> 
Figure 1: Setup and analysis of flow cytometric phagocytosis assay. (A) Scheme of the protocol including conidia and cell preparation and counterstaining. (B) Phagocytosis is analyzed by plotting of flow cytometric data of FITC-labeled conidia against anti-FITC APC counterstaining. Resulting populations indicate percentages of non-interacting leukocytes (FITC-, APC-), leukocytes with adherent conidia (FITC+, APC+) and phagocytosing leukocytes (FITC+, APC-) as well as staining artifacts (FITC-, APC+). (C) Double positive cell populations from 3 different experiments with 3 different donors (two of them performed in duplicates, one performed single) were microscopically counted for internalized and attached only conidia. (D) Representative gating strategy of flow cytometric data to detect leukocytes (CD45), identify monocytes (CD14) and neutrophils (CD66b) and determine interacting populations (FITC, anti-FITC). This figure has been modified from Hartung et al., Cytometry A, 95: 3, p. 332-338 (2019)14. <a href="https://www.jove.com/files/ftp_upload/60397/60397fig1alarge.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/60397/60397fig2a.jpg" /> 
Figure 2: Phagocytosis of conidia by human primary phagocytes depends on several conditions. (A) Percentage of cells internalizing resting conidia increased with co-incubation time. (B) Phagocytosis increased with conidial swelling time when co-incubated for 0.5 h. (C) Native conidia were better phagocytosed than fixed conidia. Data were obtained from 10 different donors (A,B) or 5 different donors (C) in 10 (A,B) or 5 (C) independent experiments. Error bars indicate SD. This figure has been modified from Hartung et al., Cytometry A, 95: 3, p. 332-338 (2019)14. <a href="https://www.jove.com/files/ftp_upload/60397/60397fig2alarge.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/60397/60397fig3a.jpg" /> 
Figure 3: Analysis of phagocytosis is a means to assess functionality of human primary phagocytes and characterize clinically relevant molds. (A) Exemplary comparison of monocytes from a healthy donor and an immunosuppressed patient (after hematopoietic stem cell transplantation) phagocytosing resting conidia for 0.5 h, 2 h and 4 h. (B) Phagocytosis of resting fungal conidia was determined for clinically relevant Aspergillus and Mucorales species after 2 h co-incubation. Data were obtained from 5 (Aspergillus) or 3 (Mucorales) different donors in 5 or 3 independent experiments each. Error bars indicate SD. Abbreviations: A. Aspergillus, L. Lichtheimia, M. Mucor, R. Rhizopus <a href="https://www.jove.com/files/ftp_upload/60397/60397fig3alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Measuring Phagocytosis of Aspergillus fumigatus Conidia by Human Leukocytes using Flow Cytometry at JoVE.com

Measuring Phagocytosis of Aspergillus fumigatus Conidia by Human Leukocytes using Flow Cytometry

This protocol provides a fast and reliable method to quantitatively measure phagocytosis of Aspergillus fumigatus conidia by human primary phagocytes using flow cytometry and to discriminate phagocytosis of conidia from mere adhesion to leukocytes.

Measuring Phagocytosis of Aspergillus fumigatus Conidia by Human Leukocytes using Flow Cytometry

Invasive pulmonary infection by the mold Aspergillus fumigatus poses a great threat to immunocompromised patients. Inhaled fungal conidia (spores) are ...

measuring-phagocytosis-aspergillus-fumigatus-conidia-human-leukocytes

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

6.7K Views.  Leibniz Institute for Infection Biology and Natural Product Research. This protocol provides a fast and reliable method to quantitatively measure phagocytosis of Aspergillus fumigatus conidia by human primary phagocytes using flow cytometry and to discriminate phagocytosis of conidia from mere adhesion to leukocytes.

Video: Measuring Phagocytosis of Aspergillus fumigatus Conidia by Human Leukocytes using Flow Cytometry

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression 1. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer's disease and multiple sclerosis 1. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. 2 In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.

This article will detail the protocol for measuring calpain activity in fixed and living cells using flow cytometry.

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, ...

Human In Vitro Suppression as Screening Tool for the Recognition of an Early State of Immune Imbalance

Regulatory T cells (Tregs) are critical mediators of immune tolerance to self-antigens. In addition, they are crucial regulators of the immune response following an infection. Despite efforts to identify unique surface marker on Tregs, the only unique feature is their ability to suppress the proliferation and function of effector T cells. While it is clear that only in vitro assays can be used in assessing human Treg function, this becomes problematic when assessing the results from cross-sectional studies where healthy cells and cells isolated from subjects with autoimmune diseases (like Type 1 Diabetes-T1D) need to be compared. There is a great variability among laboratories in the number and type of responder T cells, nature and strength of stimulation, Treg:responder ratios and the number and type of antigen-presenting cells (APC) used in human in vitro suppression assays. This variability makes comparison between studies measuring Treg function difficult. The Treg field needs a standardized suppression assay that will work well with both healthy subjects and those with autoimmune diseases. We have developed an in vitro suppression assay that shows very little intra-assay variability in the stimulation of T cells isolated from healthy volunteers compared to subjects with underlying autoimmune destruction of pancreatic &beta;-cells. The main goal of this piece is to describe an in vitro human suppression assay that allows comparison between different subject groups. Additionally, this assay has the potential to delineate a small loss in nTreg function and anticipate further loss in the future, thus identifying subjects who could benefit from preventive immunomodulatory therapy1. Below, we provide thorough description of the steps involved in this procedure. We hope to contribute to the standardization of the in vitro suppression assay used to measure Treg function. In addition, we offer this assay as a tool to recognize an early state of immune imbalance and a potential functional biomarker for T1D.

Human In Vitro Suppression as Screening Tool for the Recognition of an Early State of Immune Imbalance

Tregs are potent suppressors of the immune system. There is a lack of unique surface markers to define them, hence, definitions of Tregs are primarily functional. Here we describe an optimized in vitro assay capable of identifying immune imbalance in subjects at risk to develop T1D.

Regulatory T cells (Tregs) are critical mediators of immune tolerance to self-antigens. In addition, they are crucial regulators of the immune response ...

Examination of Thymic Positive and Negative Selection by Flow Cytometry

A healthy immune system requires that T cells respond to foreign antigens while remaining tolerant to self-antigens. Random rearrangement of the T cell receptor (TCR) &alpha; and &beta; loci generates a T cell repertoire with vast diversity in antigen specificity, both to self and foreign. Selection of the repertoire during development in the thymus is critical for generating safe and useful T cells. Defects in thymic selection contribute to the development of autoimmune and immunodeficiency disorders1-4. 
T cell progenitors enter the thymus as double negative (DN) thymocytes that do not express CD4 or CD8 co-receptors. Expression of the &alpha;&beta;TCR and both co-receptors occurs at the double positive (DP) stage. Interaction of the &alpha;&beta;TCR with self-peptide-MHC (pMHC) presented by thymic cells determines the fate of the DP thymocyte. High affinity interactions lead to negative selection and elimination of self-reactive thymocytes. Low affinity interactions result in positive selection and development of CD4 or CD8 single positive (SP) T cells capable of recognizing foreign antigens presented by self-MHC5.
Positive selection can be studied in mice with a polyclonal (wildtype) TCR repertoire by observing the generation of mature T cells. However, they are not ideal for the study of negative selection, which involves deletion of small antigen-specific populations. Many model systems have been used to study negative selection but vary in their ability to recapitulate physiological events6. For example, in vitro stimulation of thymocytes lacks the thymic environment that is intimately involved in selection, while administration of exogenous antigen can lead to non-specific deletion of thymocytes7-9. Currently, the best tools for studying in vivo negative selection are mice that express a transgenic TCR specific for endogenous self-antigen. However, many classical TCR transgenic models are characterized by premature expression of the transgenic TCR&alpha; chain at the DN stage, resulting in premature negative selection. Our lab has developed the HYcd4 model, in which the transgenic HY TCR&alpha; is conditionally expressed at the DP stage, allowing negative selection to occur during the DP to SP transition as occurs in wildtype mice10.
Here, we describe a flow cytometry-based protocol to examine thymic positive and negative selection in the HYcd4 mouse model. While negative selection in HYcd4 mice is highly physiological, these methods can also be applied to other TCR transgenic models. We will also present general strategies for analyzing positive selection in a polyclonal repertoire applicable to any genetically manipulated mice.

We present a flow cytometry-based method to examine T cell development in vivo using genetically manipulated mice on a wildtype or T cell receptor transgenic background.

A healthy immune system requires that T cells respond to foreign  antigens while remaining tolerant to self-antigens. Random rearrangement of the  T cell ...

Measuring Attachment and Internalization of Influenza A Virus in A549 Cells by Flow Cytometry

Attachment to target cells followed by internalization are the very first steps of the life cycle of influenza A virus (IAV). We provide here a detailed protocol for measuring relative changes in the amount of viral particles that attach to A549 cells, a human lung epithelial cell line, as well as in the amount of particles that are internalized into the cell. We use biotinylated virus which can be easily detected following staining with Cy3-labeled streptavidin (STV-Cy3). We describe the growth, purification and biotinylation of A/WSN/33, a widely used IAV laboratory strain. Cold-bound biotinylated IAV particles on A549 cells are stained with STV-Cy3 and measured using flow cytometry. To investigate uptake of viral particles, cold-bound virus is allowed to internalize at 37 &#176;C. In order to differentiate between external and internalized viral particles, a blocking step is applied: Free binding spots on the biotin of attached virus on the cell surface are bound by unlabeled streptavidin (STV). Subsequent cell permeabilization and staining with STV-Cy3 then enables detection of internalized viral particles. We present a calculation to determine the relative amount of internalized virus. This assay is suitable to measure effects of drug-treatments or other manipulations on attachment or internalization of IAV.

We present a protocol describing a semi-quantitative method for measuring both, the attachment of influenza A virus to A549 cells, as well as the internalization of virus particles into the target cells by flow cytometry.

Attachment to target cells followed by internalization are the very first steps of the life cycle of influenza A virus (IAV). We provide here a detailed ...

Measuring Granulocyte and Monocyte Phagocytosis and Oxidative Burst Activity in Human Blood

The granulocyte and monocyte phagocytosis and oxidative burst (OB) activity assay can be used to study the innate immune system. This manuscript provides the necessary methodology to add this assay to an exercise immunology arsenal. The first step in this assay is to prepare two aliquots ("H" and "F") of whole blood (heparin). Then, dihydroethidium is added to the H aliquot, and both aliquots are incubated in a warm water bath followed by a cold water bath. Next, Staphylococcus aureus (S. aureus) is added to the H aliquot and fluorescein isothiocyanate-labeled S. aureus is added to the F aliquot (bacteria:phagocyte = 8:1), and both aliquots are incubated in a warm water bath followed by a cold water bath. Then, trypan blue is added to each aliquot to quench extracellular fluorescence, and the cells are washed with phosphate-buffered saline. Next, the red blood cells are lysed, and the white blood cells are fixed. Finally, a flow cytometer and appropriate analysis software are used to measure granulocyte and monocyte phagocytosis and OB activity. This assay has been used for over 20 years. After heavy and prolonged exertion, athletes experience a significant but transient increase in phagocytosis and an extended decrease in OB activity. The post-exercise increase in phagocytosis is correlated with inflammation. In contrast to normal weight individuals, granulocyte and monocyte phagocytosis is chronically elevated in overweight and obese participants, and is modestly correlated with C-reactive protein. In summary, this flow cytometry-based assay measures the phagocytosis and OB activity of phagocytes and can be used as an additional measure of exercise- and obesity-induced inflammation.

The purpose of this manuscript is to present a method for measuring monocyte and granulocyte phagocytosis and oxidative burst activity in human blood samples.

The granulocyte and monocyte phagocytosis and oxidative burst (OB) activity assay can be used to study the innate immune system. This manuscript provides ...

Phenotypic Characterization of Macrophages from Rat Kidney by Flow Cytometry

There is increasing evidence suggesting the important role of inflammation and, subsequently, macrophages in the development and progression of renal disease. Macrophages are heterogeneous cells that have been implicated in kidney injury. Macrophages may be classified into two different phenotypes: classically activated macrophages (M1 macrophages), that release pro-inflammatory cytokines and promote fibrosis; and alternatively activated macrophages (M2 macrophages) that are associated with immunoregulatory and tissue-remodeling functions. These macrophage phenotypes need to be discriminated and analyzed to determine their contribution to renal injury. However, there are scarce studies reporting consistent phenotypic and functional information about macrophage subtypes in inflammatory renal disease models, especially in rats. This fact may be related to the limited macrophage markers used in rats, contrary to mice. Therefore, novel strategies are necessary to quantify and characterize the renal content of these infiltrating cells in a reliable way. This manuscript details a protocol for kidney digestion and further phenotypic and quantitative analysis of macrophages from rat kidneys by flow cytometry. Briefly, kidneys were incubated with collagenase and total macrophages were identified according to the dual presence of CD45 (leukocytes common antigen) and CD68 (PAN macrophage marker) in live cells.This was followed by surface staining of CD86 (M1 marker) and CD163 (M2 marker). Rat peritoneal macrophages were used as positive control for macrophage marker detection by flow cytometry. Our protocol resulted in low cellular mortality and allowed characterization of different intracellular and surface protein markers, thus limiting the loss of cellular integrity observed in other protocols. Moreover, this procedure allows the use of macrophages for further techniques, including cell sorting and mRNA or protein expression studies, among others.

This manuscript describes a detailed protocol for phenotypic and quantitative analysis of resident macrophages from rat kidneys by flow cytometry. The resulting stained cells can be also used for other applications, including cell sorting, gene expression analysis or functional studies, thus increasing the information obtained in the experimental model.

There is increasing evidence suggesting the important role of inflammation and, subsequently, macrophages in the development and progression of renal ...

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and their important role in lung development and function, as well as their lung-localized responses to infection and inflammation. To date, no unified method for identification, isolation, and handling of alveolar macrophages from humans and mice exists. Such a method is needed for studies on these important innate immune cells in various experimental settings. The method described here, which can be easily adopted by any laboratory, is a simplified approach to harvesting alveolar macrophages from bronchoalveolar lavage fluid or from lung tissue and maintaining them in vitro. Because alveolar macrophages primarily occur as adherent cells in the alveoli, the focus of this method is on dislodging them prior to harvest and identification. The lung is a highly vascularized organ, and various cell types of myeloid and lymphoid origin inhabit, interact, and are influenced by the lung microenvironment. By using the set of surface markers described here, researchers can easily and unambiguously distinguish alveolar macrophages from other leukocytes, and purify them for downstream applications. The culture method developed herein supports both human and mouse alveolar macrophages for in vitro growth, and is compatible with cellular and molecular studies.

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

This communication describes methodologies for isolation and culture of alveolar macrophages from humans and murine models for experimental purposes.

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and ...

Characterization of Human Monocyte Subsets by Whole Blood Flow Cytometry Analysis

Monocytes are key contributors in various inflammatory disorders and alterations to these cells, including their subset proportions and functions, can have pathological significance. An ideal method for examining alterations to monocytes is whole blood flow cytometry as the minimal handling of samples by this method limits artifactual cell activation. However, many different approaches are taken to gate the monocyte subsets leading to inconsistent identification of the subsets between studies. Here we demonstrate a method using whole blood flow cytometry to identify and characterize human monocyte subsets (classical, intermediate, and non-classical). We outline how to prepare the blood samples for flow cytometry, gate the subsets (ensure contaminating cells have been removed), and determine monocyte subset expression of surface markers &#8212; in this example M1 and M2 markers. This protocol can be extended to other studies that require a standard gating method for assessing monocyte subset proportions and monocyte subset expression of other functional markers.

Here we present a protocol for characterizing monocyte subsets by whole blood flow cytometry. This includes outlining how to gate the subsets and assess their expression of surface markers and giving an example of the assessment of the expression of M1 (inflammatory) and M2 markers (anti-inflammatory).

Monocytes are key contributors in various inflammatory disorders and alterations to these cells, including their subset proportions and functions, can ...

Preparation of Whole Bone Marrow for Mass Cytometry Analysis of Neutrophil-lineage Cells

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a myeloid-biased 39-antibody CyTOF panel to evaluate the hematopoietic system with a focus on the neutrophil-lineage cells by using this protocol. The CyTOF result was analyzed with an open-resource dimensional reduction algorithm, viSNE, and the data was presented to demonstrate the outcome of this protocol. We have discovered new neutrophil-lineage cell populations based on this protocol. This protocol of fresh whole BM preparation may be used for 1), CyTOF analysis to discover unidentified cell populations from whole BM, 2), investigating whole BM defects for patients with blood disorders such as leukemia, 3), assisting optimization of fluorescence-activated flow cytometry protocols that utilize fresh whole BM.

Here, we present a protocol to process fresh bone marrow (BM) isolated from mouse or human for high-dimensional mass cytometry (Cytometry by Time-Of-Flight, CyTOF) analysis of neutrophil-lineage cells.

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a ...

Measuring Erythrocyte Complement Receptor 1 Using Flow Cytometry

CR1 (CD35, Complement Receptor type 1 for C3b/C4b) is a high molecular weight membrane glycoprotein of about 200 kDa that controls complement activation, transports immune complexes, and participates in humoral and cellular immune responses. CR1 is present on the surface of many cell types, including erythrocytes, and exhibits polymorphisms in length, structure (Knops, or KN, blood group), and density. The average density of CR1 per erythrocyte (CR1/E) is 500 molecules per erythrocyte. This density varies from one individual to another (100&#8211;1,200 CR1/E) and from one erythrocyte to another in the same individual. We present here a robust flow cytometry method to measure the density of CR1/E, including in subjects expressing a low density, with the help of an amplifying immunostaining system. This method has enabled us to show the lowering of CR1 erythrocyte expression in diseases such as Alzheimer&#39;s disease (AD), systemic lupus erythematosus (SLE), AIDS, or malaria.

The aim of this method is to determine the CR1 density in the erythrocytes of any subject by comparing with three subjects whose erythrocyte CR1 density is known. The method uses flow cytometry after immunostaining of the subjects&#39; erythrocytes by an anti-CR1 monoclonal antibody coupled to an amplified system using phycoerythrin (PE).

CR1 (CD35, Complement Receptor type 1 for C3b/C4b) is a high molecular weight membrane glycoprotein of about 200 kDa that controls complement activation, ...