We are grateful for the technical assistance of Dr. Suchitra Swaminathan who runs the Northwestern Feinberg School of Medicine Flow Cytometry core. Funding was provided by (NIH) DK101675.

All animal work was approved by the Northwestern IACUC. All experiments were completed in accordance and compliance with all relevant regulatory and institutional guidelines. For human subjects donating blood, an informed consent was presented and signed; in addition, all human subjects in this study were treated in accordance with the institutional and federal guidelines for human welfare.
NOTE: The protocol steps are listed under the following subsections: (i) Mouse Bone Marrow Cell Isolation; (ii) PMN Isolation from Murine Bone Marrow; (iii) PMN Isolation from Human Blood; (iv) MP Isolation from PMN Supernatants (by Ultracentrifugation); (v) Characterization of MPs by Western Blotting; (vi) Characterization of MPs by Flow Cytometry; (viii) Application of isolated MPs to Study Wound Healing
1. Mouse Bone Marrow Cell Isolation
<ol style="margin-left: 40px;">
	<li>Dissection of the femur and tibia from the mouse hind legs
	<ol>
		<li>Prepare a euthanasia box (e.g., an empty 1 mL tip box) for inhalation of anesthesia. Add a few drops of 100% isoflurane onto a sterile cloth taped to the inside of a plastic box lid. As per new IACUC guidelines, animals should not come in direct contact with isoflurane, thus place a tip holder between the mouse and the gauze containing the isoflurane. 
		NOTE: Isoflurane is a powerful inhalational anesthetic and should be handled with extreme caution inside a fume hood.</li>
		<li>Euthanize mice as per IACUC recommendation. Place a mouse inside the euthanasia box described in step 1.1.1 for 1&#8211;5 min until it has ceased breathing. Ensure the animal&#39;s death by performing a cervical dislocation.</li>
		<li>Lift the abdominal skin using stainless steel, 12 cm curved, 0.17 mm X 0.1 mm tweezers and cut the skin half way between the upper and lower extremities. Peel the skin from here to the most distal section of the mouse body including the lower extremities. With 10 cm long, straight dissecting scissors, remove the muscles from the hind legs and dislocate the hip joint leaving the femur head intact.</li>
		<li>Use scissors to dissect the muscles from the femur and the tibia and separate the femur from the tibia. Make sure the ends of the bones remain intact as they contain significant amounts of bone marrow.</li>
		<li>Remove all remaining flesh by rolling the bones inside a paper towel. Place the clean bones in a Petri dish containing Serum-Free Medium (SFM, i.e., Dulbecco&#39;s Modified Eagle&#39;s medium (DMEM) without serum or antibiotics).</li>
		<li>Rinse the bones in 70% ethanol and then in SFM.</li>
		<li>Prepare 50 mL of SFM in a 50 mL conical tube. Load 10 mL of SFM into a 10 mL syringe and attach it to a 25 and 5/8 inch gauge needle.</li>
		<li>Trim the end of the bones with scissors until an opening to the bone marrow (reddish color) is visible.</li>
		<li>Flush the bone marrow cells into a new 50 mL tube. Use a 10 mL syringe with a 26 and 5/8 inch gauge needle with approximately 3 mL/bone of SFM. When finished flushing, resuspend the cells in the tube by pipetting (use disposable 1 mL plastic tips) to break cell aggregates.</li>
	</ol>
	</li>
</ol>
2. PMN Isolation from Murine Bone Marrow
<ol>
	<li>Red cell lysis
	<ol>
		<li>Pellet the collected bone marrow by centrifuging at 350 x g for 8 min at 4 &#176;C.</li>
		<li>Resuspend the cell pellet in 20 mL of 0.2% NaCl for approximately 20-30 s to lyse the erythrocyte fraction. Do not exceed 30 s as this will result in PMN lysis. Add 20 mL of 1.6% NaCl to restore osmolality. 
		NOTE: This step may lead to very little PMN activation.</li>
		<li>Wash the cells with 2 mL of PBS and centrifuge as in step 2.1.1 above.Remove the supernatant.</li>
		<li>Proceed to isolating PMNs by density gradient centrifugation.</li>
	</ol>
	</li>
	<li>Isolation of neutrophils by density gradient centrifugation
	<ol>
		<li>Warm polysucrose and sodium diatrizoate solutions, with densities of 1.119 g/mL and 1.077 g/mL (see Table of Materials), to room temperature (RT) before use.</li>
		<li>Prepare polysucrose and sodium diatrizoate gradients fresh, immediately prior to bone marrow cell application by slowly layering 3 mL of the 1.077 g/mL density solution over 3 mL of the 1.119 g/mL density solution in 15 mL conical tubes. 
		NOTE: Preparation of the gradient solution too far in advance will result in mixing of the layers and poor neutrophil purity and recovery.</li>
		<li>Resuspend the bone marrow cell pellet in 1 mL of ice-cold PBS and overlay the resulting cell suspension on top of the polysucrose and sodium diatrizoate gradient (slowly, to avoid mixing of the layers).</li>
		<li>Centrifuge for 30 min at 850 x g at 25 &#176;C at RT, without brake. Keep the reagents at RT for effective separation of the neutrophils from other cells present in the bone marrow. Set the centrifuge at slow deceleration to allow for the gradient to be efficiently formed and maintained after the centrifugation step.</li>
		<li>Visually locate a mononuclear cell layer at the interface of PBS and 1.077 g/mL density solution (upper layer). Remove this layer and the rest of the density solution without disturbing the PMN band.</li>
		<li>Collect an entire PMN layer and some of the underlying polysucrose and sodium diatrizoate 1.119 g/mL solution into 15 mL tubes using a transfer pipette. 
		NOTE: The purity of the isolated PMNs will depend on a complete removal of the upper layer.</li>
		<li>Top the tubes with filtered PBS (filtered through a 0.1 &#181;m pore size filter) and centrifuge at 300 x g for 5 min, at 4 &#176;C.</li>
		<li>Wash the pelleted PMNs with 2 mL of PBS using centrifugation conditions as in step 2.2.7.</li>
		<li>Resuspend in 1 mL of filtered PBS and count the isolated PMNs by hemocytometer. 
		NOTE: For this isolation method, the resulting PMN purity amounts to ~ 85-90%, as was assessed by flow cytometry. The remaining contaminating fractions consists primarily of mononuclear lymphocytes.</li>
	</ol>
	</li>
	<li>PMN stimulation to induce MP release 
	NOTE: PMNs can be stimulated to release MPs using various activating conditions, including N-formyl-l-methionyl-leucyl-l-phenylalanine (fMLF), phorbol-12-myristate-13-acetate (PMA), or Interferon gamma (IFN&#947;). Importantly, prior to simulation, PMNs should be kept on ice at all times.
	<ol>
		<li>Pellet PMNs (300 x g, 5 min, 4 &#176;C) and resuspend in 0.1 &#181;m filtered Hank&#39;s Balanced Salt solution (HBSS) solution, containing the stimulating agent of choice. For optimal stimulation, use 100 &#181;L stimulation solution per 10 million PMNs for 20-30 min at 37 &#176;C in 15 mL tubes. 
		NOTE: The following concentrations of activators were successfully used in our previous work: fMLF(0.5-1 &#181;M for human and 2-5 &#181;M for mouse PMNs), PMA (200 nM), and IFN&#947; (100 ng/mL). If the release of pre-labeled MPs is desired, incubate unstimulated PMNs (1-5 x 106) with N-(2-aminoethyl) maleimide-FITC (see Table of Materials) (1 &#181;M in 100 &#181;L HBSS, 15 min, 37 &#176;C). Proceed with steps 2.3.1-2.3.3.</li>
		<li>Spin down at 850 x g for 5 min at 4 &#176;C, to remove PMNs and transfer cell-free supernatants to new 1.5 mL microcentrifuge tubes.</li>
		<li>For MP isolation from cell-free supernatants, continue to step 4.2.</li>
	</ol>
	</li>
</ol>
3. PMN Isolation from Human Blood
<ol>
	<li>Recruit healthy volunteers for blood extraction according to Institutional IRB guidelines.
	<ol>
		<li>Collect blood from the forearm of a healthy volunteer into commercially available 5 mL sodium citrate-containing tubes (see Table of Materials).</li>
		<li>Pipette 5 mL of dextran and sodium diatrizoate solution (density of 1.113 g/mL, see Table of Materials) into sterile 15 mL tubes and slowly layer 5 mL of freshly isolated human peripheral blood on top of it.</li>
		<li>Centrifuge the tubes for 50 min at 400 x g at RT (25 &#176;C) with low acceleration and no brake.</li>
		<li>At the end of the centrifugation, remove the plasma and mononuclear cells (upper layer) using a sterile plastic suction pipette.</li>
		<li>Transfer the bottom layer (PMNs) into a new sterile 50 mL conical tube.</li>
		<li>Add an equal volume of 0.45% NaCl to PMNs (mix well and gently by rotating the tube).</li>
		<li>Centrifuge for 10 min at 400 x g at 25 &#176;C.</li>
	</ol>
	</li>
	<li>Red cell lysis
	<ol>
		<li>Add 10 mL of ice-cold, sterile water to pelleted cells for 45 s followed by 10 mL of ice-cold sterile 1.8% NaCl to restore osmolality. 
		NOTE: All following steps for PMN isolation should be performed on ice to avoid PMN activation and premature MP release. The lysis step itself results in minor but not significant PMN activation, however, it is essential for isolating a pure PMN population for functional assays.</li>
		<li>Centrifuge the cells for 10 min at 250 x g at 4 &#176;C with soft deceleration.</li>
		<li>Repeat steps 3.2.1 and 3.2.2.</li>
		<li>Resuspend PMNs in 5 mL of ice-cold HBSS without calcium or magnesium. Count total live cells (use a hemocytometer and viability staining at 1:2 dilution, see Table of Materials) before stimulation. 
		NOTE: Isolated PMNs should be used for stimulation or other experiments within 2 h of isolation to avoid functional and cellular changes and cell death.</li>
	</ol>
	</li>
</ol>
4. MP Isolation from Activated PMN Supernatants
NOTE: A similar protocol is used for the isolation of MPs from murine and human PMNs.
<ol>
	<li>Pellet PMNs (300 x g, 5 min, 4 &#176;C) and resuspend in 0.1 &#181;m filtered HBSS solution, containing the stimulating agent of choice (for example 1 &#181;M fMLF, see Note: 2.3.1). For optimal stimulation use 100 &#181;L stimulation solution per 10 million PMNs for 20-30 min at 37 &#176;C in 15 mL tubes.</li>
	<li>Following stimulation, spin down activated PMNs at 600 x g for 5 min at 4 &#176;C and transfer cell-free supernatants to new 1.5-mL microcentrifuge tubes.</li>
	<li>Centrifuge the PMN-cleared supernatant at 13,000 x g, 10 min, 4 &#176;C to remove cell debris and transfer the cleared supernatant into a new ultracentrifuge tube.</li>
	<li>Centrifuge for 1 h at 100,000 x g, at 4 &#176;C. Remove supernatants before MP storage. As needed, store pelleted MPs in paraffin sealed ultracentrifuge tubes at -80 &#176;C until use in experiments.</li>
</ol>
5. Examination of PMN-MPs by Western Blotting
<ol>
	<li>Add 1% SDS buffer (50-200 &#181;L with 100 mM Tris pH: 7.4) to MPs pellet (from step 4.4), transfer to new 1.5 mL microcentrifuge tubes and boil the lysed MPs for 5 min. 
	NOTE: The number of MPs and the lysis buffer volume must be adjusted and optimized for each protein of interest.</li>
	<li>Load equal amounts of PMN-MPs per lane in loading buffer containing 10% &#946;-mercapto-ethanol. 
	&#8203;NOTE: With our method we load MPs that were isolated from the same number of stimulated PMNs.
	<ol>
		<li>Separate the lysates by SDS-PAGE on 10% polyacrylamide gel (use 50-100 V for 1-1.5 h) and transfer total proteins onto nitrocellulose membranes as described9.</li>
	</ol>
	</li>
	<li>Block membranes for 1 h with 5% non-fat milk in 0.05% Tween-20 Tris-buffered saline, and incubate it with an appropriate primary (overnight at 4 &#176;C), followed by secondary HRP-conjugated antibodies.</li>
	<li>Visualize the bands by adding chemoluminescence solution to the membrane and exposure to an X-ray film inside a light-proof cassette (1-24 h).</li>
</ol>
6. Examination of PMN-MPs by Flow Cytometry
<ol>
	<li>For antibody staining, dilute antibodies appropriately in FACS buffer (PBS with 0.1 mM EDTA, 0.1% sodium azide). Resuspend pelleted PMN-MPs (derived from 1-3 x 106 PMNs/condition) in 100 &#181;L antibody solution.
	<ol>
		<li>If staining for Annexin V, resuspend pelleted PMN-MPs in Annexin V buffer containing 5 &#181;L of FITC-conjugated Annexin V. If co-staining with antibodies, add the desired antibodies (for an example see Table of Materials) directly to the Annexin V solution. Add fluorescent lipid dye, N-(2-aminoethyl) maleimide (see Table of Materials), at 1 &#181;M concentration in 100 &#181;L of FACS buffer to label and identify MPs.</li>
	</ol>
	</li>
	<li>Incubate the MPs in staining solution for at least 20 min at 4 &#176;C.</li>
	<li>Wash the MPs by ultracentrifugation (1 h at 100,000 x g, at 4 &#176;C); discard the supernatant.</li>
	<li>Resuspend in FACS buffer and run the samples on a designated flow cytometer equipped with an upgraded electronic unit for increased sensitivity (see Table of Materials).</li>
</ol>
7. Applications for Isolated PMN-MPs to Study PMN Function in Wound Healing
<ol>
	<li>In vivo administration of PMN-MPs to colonic wounds
	<ol>
		<li>Anesthetize mice by an intraperitoneal injection of a mixture of ketamine (100 mg/kg) and xylazine (5 mg/kg). Confirm anesthesia by pedal reflex (firm toe pinch) and adjust as necessary.</li>
		<li>Place the mouse on its stomach and using 28-cm biopsy forceps and an endoscope equipped with a high-resolution camera (such as a veterinary endoscope for small animal use, see Table of Materials), generate 3-5 superficial wounds along the dorsal side of the colon. Upon completion of wounding, return mice to a cage placed on a temperature-controlled (37 &#176;C) mat until recovered.</li>
		<li>Anesthetize mice (as in step 7.1.1). Use an endoscope to acquire images of inflicted wounds 24 h (day 1) post-wounding. Let mice recover as in step 7.1.2.</li>
		<li>At the same time (24 h post-wounding), administer murine PMN-MPs derived from 2 x 106 PMNs in 100 &#181;L of HBSS+ directly into each wound site using a colonoscopy-based microinjection system (use a 29 G needle)9. Three days later (day 4 post-wounding) anesthetize mice (as in step 7.1.1) and use an endoscope to reacquire images of healing wounds. Let mice recover as in step 7.1.2.</li>
		<li>Using preferred image analysis software (see Table of Materials), outline visible wound regions in acquired images, measure the area of the same wound at days 1 and 4 post-wounding, and calculate the percentage of wound closure.</li>
	</ol>
	</li>
	<li>Human epithelial cells in vitro wound healing
	<ol>
		<li>Count Caco-2 BBe or T84, human intestinal epithelial cells by hemocytometer and seed 5 x 105 cells/well in 24-well culture plates. Incubate the cells in a standard incubation chamber at 37 &#176;C and 5% CO2. Caco-2 BBe or T84 reach confluency within 48 h9. To culture epithelial cells, use DMEM and DMEM-F12 (50:50) with supplements as previously described9.</li>
		<li>Generate mechanical scratch wounds in the monolayer using a pipette tip and low suction as previously described4,9. Acquire images of wound areas immediately after scratching using an inverted phase-contrast differential interference microscope (use 5X or 10X objectives), to be used as a time = 0 reference point.</li>
		<li>Add MPs (derived from 2-4 X 106 PMNs) to scratched-wounded epithelial cell monolayers, and incubate for an additional 24 h (48 h post-wounding, at 37 &#176;C with 5% CO2). At this time re-acquire images of the scratch-wounds.</li>
		<li>Use preferred image analysis software (see Table of Materials) to measure wound areas at t = 0 and t = 48 h. Calculate the percentage of wound closure by determining the difference in wound area at the selected time points.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Hickey, M. J., Kubes, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intravascular+immunity:+the+host-pathogen+encounter+in+blood+vessels.">Intravascular immunity: the host-pathogen encounter in blood vessels.</a> Nat Rev Immunol. 9 (5), 364-375 (2009).</li><li>Nathan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neutrophils+and+immunity:+challenges+and+opportunities.">Neutrophils and immunity: challenges and opportunities.</a> Nat Rev Immunol. 6 (3), 173-182 (2006).</li><li>Kolaczkowska, E., Kubes, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neutrophil+recruitment+and+function+in+health+and+inflammation.">Neutrophil recruitment and function in health and inflammation.</a> Nat Rev Immunol. 13 (3), 159-175 (2013).</li><li>Butin-Israeli, V., 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=Deposition+of+microparticles+by+neutrophils+onto+inflamed+epithelium:+a+new+mechanism+to+disrupt+epithelial+intercellular+adhesions+and+promote+transepithelial+migration.">Deposition of microparticles by neutrophils onto inflamed epithelium: a new mechanism to disrupt epithelial intercellular adhesions and promote transepithelial migration.</a> FASEB J. , (2016).</li><li>Gasser, O., Schifferli, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activated+polymorphonuclear+neutrophils+disseminate+anti-inflammatory+microparticles+by+ectocytosis.">Activated polymorphonuclear neutrophils disseminate anti-inflammatory microparticles by ectocytosis.</a> Blood. 104 (8), 2543-2548 (2004).</li><li>Swamydas, M., Lionakis, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation,+purification+and+labeling+of+mouse+bone+marrow+neutrophils+for+functional+studies+and+adoptive+transfer+experiments.">Isolation, purification and labeling of mouse bone marrow neutrophils for functional studies and adoptive transfer experiments.</a> J Vis Exp. (77), e50586 (2013).</li><li>Shet, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizing+blood+microparticles:+technical+aspects+and+challenges.">Characterizing blood microparticles: technical aspects and challenges.</a> Vasc Health Risk Manag. 4 (4), 769-774 (2008).</li><li>Dey-Hazra, E., 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=Detection+of+circulating+microparticles+by+flow+cytometry:+influence+of+centrifugation,+filtration+of+buffer,+and+freezing.">Detection of circulating microparticles by flow cytometry: influence of centrifugation, filtration of buffer, and freezing.</a> Vasc Health Risk Manag. 6, 1125-1133 (2010).</li><li>Slater, T. W., 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=Neutrophil+Microparticles+Deliver+Active+Myeloperoxidase+to+Injured+Mucosa+To+Inhibit+Epithelial+Wound+Healing.">Neutrophil Microparticles Deliver Active Myeloperoxidase to Injured Mucosa To Inhibit Epithelial Wound Healing.</a> J Immunol. 198 (7), 2886-2897 (2017).</li><li>Enjeti, A. K., Lincz, L., Seldon, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bio-maleimide+as+a+generic+stain+for+detection+and+quantitation+of+microparticles.">Bio-maleimide as a generic stain for detection and quantitation of microparticles.</a> Int J Lab Hematol. 30 (3), 196-199 (2008).</li><li>Headland, S. E., 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=Neutrophil-derived+microvesicles+enter+cartilage+and+protect+the+joint+in+inflammatory+arthritis.">Neutrophil-derived microvesicles enter cartilage and protect the joint in inflammatory arthritis.</a> Sci Transl Med. 7 (315), (2015).</li><li>Andersson, T., Dahlgren, C., Lew, P. D., Stendahl, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+surface+expression+of+fMet-Leu-Phe+receptors+on+human+neutrophils.+Correlation+to+changes+in+the+cytosolic+free+Ca2++level+and+action+of+phorbol+myristate+acetate.">Cell surface expression of fMet-Leu-Phe receptors on human neutrophils. Correlation to changes in the cytosolic free Ca2+ level and action of phorbol myristate acetate.</a> J Clin Invest. 79 (4), 1226-1233 (1987).</li></ol>

The authors have no conflict of interest of any kind related to this communication

Protocols for the isolation and characterization of PMN-derived MPs are described in this communication. Several key critical points must be taken into account for the success of the procedure. First, PMNs must be isolated fresh and used in experiments within 2 h of isolation to prevent spontaneous activation and degranulation. All handling of PMNs during isolation and up to the point of stimulation must be performed on ice to prevent activation and premature MP release12. Second, ultracentrifugation for 1 h or more maximizes the pelleting of MPs. Third, for analysis by flow cytometry, the instrument must be properly calibrated with a mix of instrument-specified fluorescent beads (the beads type is adjusted to a particular instrument) to correctly define MP sizes. For example, while one manufacturer of flow cytometers recommends using FSC beads as a size-related parameter, others have been optimized for SSC beads. In addition, the flow cytometer to be used for MP analysis must be equipped with upgraded electronics to best resolve MPs from the background noise.
To correctly identify MPs in addition to size parameters, the use of fluorescence staining as described in the procedures above is highly recommended. Furthermore, all solutions during the preparation and isolation should be filtered through a 0.1 &#181;m filter system to minimize inclusion of dust particles or other precipitates that will increase the background noise during acquisition by flow cytometry. Importantly, storage conditions of MPs should be considered. Little evidence8, and our own unpublished observations, suggest that MP freezing leads to membrane rupture and MP breakage. Finally, varying PMN activation conditions and timing may change and improve MP yield.
There are some limitations to this method. Isolation of mouse PMNs from bone marrow is typically not pure (~85&#8211;90%) and is contaminated by other immune cells (primarily mononuclear lymphocytes) as was determined by flow cytometric analysis. Thus, the resulting isolates may include small amounts of MPs released by other immune cells. Alternatives are available and include isolation of PMNs by commercially available magnetic beads, however, this would be a longer and a significantly costlier procedure. Finally, without fluorescence labeling, in addition to size parameters, definitive differentiation of MPs from dust or other soluble or airborne particles of similar sizes that can contaminate the sample is challenging and error-prone.
In the future, sorting of PMN-MPs labeled with specific markers will help elucidate MP composition under specific stimulatory conditions or models of disease, and hopefully allow for the use of MPs as diagnostic markers and potential therapeutic targets to treat inflammatory diseases.

Recently, &#34;microparticles/microvesicles&#34; originating from the cell cytosol or plasma membrane have become of great scientific interest, as emerging data suggest that these structures, ranging from 50-1,000 nm in diameter, can carry biological information and serve as a non-canonical method of cellular communication. Immune cell-derived MPs and particularly those produced by polymorphonuclear neutrophils (PMNs) are of great interest given the important role of PMNs in host defense1,2, inflammatory responses3, and wound-healing4. Intriguingly, thus far, numerous reports have shown both pro-inflammatory and anti-inflammatory functions of PMN-MPs5, suggesting a potential context-, disease-, species-, and organ-specific role of MPs.
Described protocols in this communication provide a cost-effective, innovative, and adaptable method to study the function of MPs in health and disease. They are applicable to many model organisms, organs, and stimulation conditions. They allow for the identification of several types of MPs and can be used in the future to address their pro-inflammatory and anti-inflammatory functions. As an example, described here is how to study the function of PMN-MPs in epithelial wound healing in vitro and in vivo. The presented protocol for isolation of mouse bone marrow-derived PMNs was adapted with some modifications from a previously described method6.
Furthermore, protocols described in this study allow for the detection and characterization of specific markers that can be found on PMN-MPs by two complementary methods: Western blot and flow cytometry. We find that immunoblotting of MPs using standard protocols5 is easy and reliable, however, recent advances in sensitivity of flow cytometry instruments, and improved noise-to-signal ratio now allow for further analysis of MPs using this method. The described protocols in this study incorporate recent advances and recommendations from original research articles, including modifications to centrifugation speed and time, the addition of sample filtering and freezing/storage conditions7,8, and how to reduce the &#34;background noise&#34;, improve the detection limit of PMN-MPs, and discriminate between different sizes of MPs.

<table border="0" cellpadding="0" cellspacing="0" style="width:968px;" width="967">
	<tbody>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">DMEM</td>
			<td style="width:181px;">Corning</td>
			<td style="width:181px;">10-041-CV</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">Fetal Bovine Serum (Heat inactivated)</td>
			<td style="width:181px;">Atlanta Biologics</td>
			<td style="width:181px;">S11150</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="44">
			<td height="44" style="height:44px;width:257px;">Penicillin/Streptomycin (10,000 units penicillin / 10,000 mg/ml strep)</td>
			<td style="width:181px;">GIBCO</td>
			<td style="width:181px;">15140</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">0.5 M EDTA</td>
			<td style="width:181px;">Fisher</td>
			<td style="width:181px;">BP2482</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="44">
			<td height="44" style="height:44px;width:257px;">Sodium chloride</td>
			<td style="width:181px;">Sigma</td>
			<td style="width:181px;">S9888</td>
			<td style="width:348px;">Sterile and filtered through a 0.1 micron filter unit</td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">Sterile filtered Histopaque 1077</td>
			<td style="width:181px;">Sigma</td>
			<td style="width:181px;">10771</td>
			<td></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">Sterile filtered Histopaque 1119</td>
			<td style="width:181px;">Sigma</td>
			<td style="width:181px;">11191</td>
			<td style="width:348px;">Density 1.119 g/ml.&#160; Solution of polysucrose and&#160; sodium diatrizoate. Bring to room temperature</td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">Phosphate Buffered Saline (PBS) without Calcium and Magnesium</td>
			<td style="width:181px;">Cellgro</td>
			<td style="width:181px;">210-40-CV</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">Ethyl Alcohol (200 proof)</td>
			<td style="width:181px;">Decon labs</td>
			<td style="width:181px;">2701</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">HBBS</td>
			<td style="width:181px;">Corning</td>
			<td style="width:181px;">21-022-CV</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">Trypan Blue Solution, 0.4%</td>
			<td style="width:181px;">Sigma</td>
			<td style="width:181px;">T8154&#160;SIGMA</td>
			<td style="width:348px;">Diluted 1:2 for cell viability counting</td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">Isoflurane</td>
			<td style="width:181px;">Abbot</td>
			<td style="width:181px;">50033</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">Anti-human CD11b-APC conjugated</td>
			<td style="width:181px;">Biolegend</td>
			<td style="width:181px;">301350</td>
			<td style="width:348px;">Clone ICRD44 (Final concetration: 1 &#181;g/mL)</td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;">BODIPY FL</td>
			<td>Invitrogen</td>
			<td>B10250</td>
			<td style="width:348px;">N-(2-aminoethyl) maleimide&#160;</td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;">Annexin V Binding Buffer</td>
			<td>Biolegend</td>
			<td>422201</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;">Calibration Beads for Flow Cytometry</td>
			<td>BioCytex</td>
			<td>7803</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;">FITC Annexin V</td>
			<td>Biolegend</td>
			<td>640906</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;">Polymorphprep</td>
			<td>Axis-Shield PoC AS</td>
			<td></td>
			<td style="width:348px;">Sodium diatrizoate/Dextran 500, density 1.113 g/ml</td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">C57BL/6 mice</td>
			<td style="width:181px;">Jackson Labs</td>
			<td style="width:181px;"></td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:257px;">15 ml centrifuge tubes</td>
			<td style="width:181px;">Corning</td>
			<td style="width:181px;">430053</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">50 ml centrifuge tubes</td>
			<td style="width:181px;">BD Falcon</td>
			<td style="width:181px;">352070</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">25 ml serological pipettes</td>
			<td style="width:181px;">Celltreat</td>
			<td style="width:181px;">229225B</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">10 ml serological pipettes</td>
			<td style="width:181px;">Celltreat</td>
			<td style="width:181px;">229210B</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">5 ml serological pipettes</td>
			<td style="width:181px;">Celltreat</td>
			<td style="width:181px;">229205B</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">Pasteur pipettes&#160;</td>
			<td style="width:181px;">BD Falcon</td>
			<td style="width:181px;">357575</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">25 G x 5/8 in. Needles (precision glide needles)</td>
			<td style="width:181px;">BD</td>
			<td style="width:181px;">305122</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">100 &#181;m cell strainers</td>
			<td style="width:181px;">Celltreat</td>
			<td style="width:181px;">229485</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;width:257px;">Vacutainer tubes</td>
			<td style="width:181px;">BD</td>
			<td style="width:181px;">367251</td>
			<td style="width:348px;"></td>
		</tr>
		<tr height="18">
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;">LSR Fortessa Special Order Research Product (SORP)</td>
			<td>BD</td>
			<td>(SORP)</td>
			<td></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:19px;">Swinging bucket centrifuge</td>
			<td>ThermoFisher Scientific</td>
			<td>75007210</td>
			<td></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;">Ultracentrifuge</td>
			<td>Beckman</td>
			<td>(L8-80 M)</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Micro-centrifuge</td>
			<td>ThermoFisher Scientific</td>
			<td>&#160;VV-17703-15 (Fresco 17)</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Swinging bucket centrifuge</td>
			<td>ThermoFisher Scientific</td>
			<td>&#160;75004503 (Megafuge 40R)</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Biopsy forceps, 28 cm</td>
			<td>Storz</td>
			<td>27071zj</td>
			<td></td>
		</tr>
		<tr height="17">
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Image J</td>
			<td>National Institute of Health</td>
			<td>Open source&#160;</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolation and characterization of neutrophil-derived microparticles for functional studies

Polymorphonuclear neutrophil-derived microparticles (PMN)-MPs) are lipid bilayer, spherical microvesicles with sizes ranging from 50&#8211;1,000 nm in diameter. MPs are a newly evolving, important part of cell-to-cell communication and signaling machinery. Because of their size and the nature of their release, until recently MP existence was overlooked. However, with improved technology and analytical methods their function in health and disease is now emerging. The protocols presented here are aimed at isolating and characterizing PMN-MPs by flow cytometry and immunoblotting. Moreover, several implementation examples are given. These protocols for MP isolation are fast, low-cost, and do not require the use of expensive kits. Furthermore, they allow for the labeling of MPs following isolation, as well as pre-labeling of source cells prior to MP release, using a membrane-specific fluorescent dye for visualization and analysis by flow cytometry. These methods, however, have several limitations including purity of PMNs and MPs and the need for sophisticated analytical instrumentation. A high-end flow cytometer is needed to reliably analyze MPs and minimize false positive reads due to noise or auto-fluorescence. The described protocols can be used to isolate and define MP biogenesis, and characterize their markers and variation in composition under different stimulating conditions. Size heterogeneity can be exploited to investigate whether the content of membrane particles versus exosomes is different, and whether they fulfill different roles in tissue homeostasis. Finally, following isolation and characterization of MPs, their function in cellular responses and various disease models (including, PMN-associated inflammatory disorders, such as Inflammatory Bowel Diseases or Acute Lung Injury) can be explored.

Representative flow cytometric analysis of MPs that were isolated from human and mouse PMNs are shown in Figure 1. The size heterogeneity of PMN-MPs can be assessed by comparison to known sized beads as shown in Figure 1A, B for human MPs. Note, no significant differences in size heterogeneity were observed between mouse and human MPs. Similarly, using flow cytometry and fluorescence labeling, expression of protein/s of choice by isolated MPs of either mouse or human origin can be determined. For example, expression of Phosphatidyl serine (PhS) can be examined by Annexin V staining. Murine bone marrow PMN-MPs are shown in Figure 1C, D. Expression of several markers of choice can also be assessed as shown for Annexin V and CD11b staining of human MPs, Figure 2A, B. Another method to detect PMN-MPs by flow cytometry is to stain freshly-isolated PMNs prior to stimulation as shown in Figure 2C. For example, PMN staining with the lipid marker N-(2-aminoethyl) maleimide-FITC prior to fMLF stimulation results in the release of green MPs, that can be readily detected by flow. Of note, while staining with N-(2-aminoethyl) maleimide-FITC has been previously used to label and detect MPs that were obtained from peripheral human blood10 or injected into animals11, the use of other markers for labeling purposes for in vitro or in vivo application should be determined by each investigator. In addition to flow cytometry, PMN-MPs can be analyzed by immunoblotting for proteins of interest. As shown in Figure 3, MPs derived from human PMNs that were stimulated with several known activators, were examined for the expression of key inflammatory (matrix metallopeptidase 9 (MMP-9) and myeloperoxidase (MPO)) and anti-inflammatory (Annexin A1) molecules. As evident from representative immunoblots, PMN stimulation with IFN&#947; (50 ng/mL), PMA (200 nM), and fMLF (1 &#181;M) resulted in MPs expressing varying levels of MMP-9. However, only perturbation of the actin cytoskeleton by Latrunculin B (1 &#181;M) prior to stimulation with fMLF (5 &#181;M) led to abundant presence of MPO in PMN-MPs. Similarly, varying levels of Annexin A1 (high to no detection) were detected on MPs following the described activating conditions. These results suggest that PMN-MP composition is stimulus-dependent.
Finally, Figure 4 shows how isolated PMN-MPs can be used to study wound healing in vitro and in vivo in colonic injury. PMN-MPs can be added to scratch-wounded epithelial monolayers in cultures, where healing can be monitored by imaging acquisition at pre-determined time points. MPs can further be microinjected directly into colonic wounds, that were generated by biopsy forceps and endoscopic imaging9, and their effect on healing can be assessed. For both in vitro and in vivo analysis of wound healing, images of inflicted wounds are acquired immediately post wounding (or otherwise as specified) and continuously through the healing process at pre-determined time points. Using commercially available image analysis software, changes in wound area (size) are measured and used to determine the wound closure rate. Application of PMN-MPs that contain MPO to either cultured epithelial monolayers or in vivo to colonic wounds has detrimental effects, leading to delayed healing9.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56949/56949fig1.jpg" /> 
Figure 1. Analysis of PMN-MP size and surface markers by flow cytometry. (A) Flow cytometer optimized control beads (see Table of Materials) of known sizes and scatter values are shown in the SSC and FSC orthogonal representation. The beads are used to get a relative size comparison with a PMN-MP sample shown in B. (B) Human PMN-MPs were isolated and analyzed by flow cytometry using conditions described for the beads. The heterogeneity in MP sizes can be seen. (C-D) MPs derived from fMLF-stimulated murine bone marrow PMNs were analyzed by flow cytometry. Representative flow diagrams show unstained (C) or Annexin V-FITC stained (D) MPs. Rectangular area shows Annexin V-positive MPs (FITC-positive MPs). SSC: Side Scatter; FSC: Forward Scatter; PhS: Phosphatidyl serine. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56949/56949fig1large.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/56949/56949fig2.jpg" /> 
Figure 2. PMN-MP staining and analysis by flow cytometry. (A) Unstained and (B) Annexin V-FITC- and hCD11b-APC-stained MPs. Square area encloses an Annexin V/CD11b double positive population of PMN-MPs. (C) Freshly isolated human PMNs (1 x 106) were stained with a fluorescent dye, N-(2-aminoethyl) maleimide-FITC and stimulated with fMLF. MPs were isolated from cell supernatants and analyzed by flow cytometry. Rectangular area encloses a N-(2-aminoethyl) maleimide-FITC positive MP population (M-FITC, Y-axis label). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56949/56949fig2large.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/56949/56949fig3.jpg" /> 
Figure 3. Composition of PMN-MPs is stimulus-dependent. (A) Human PMNs were stimulated with either IFN&#947;, TNF&#945;, fMLF, PMA, or a combination of latranculin B followed by fMLF (LtB- fMLF). MPs were isolated from the resulting cell supernatants by ultracentrifugation and protein lysates were prepared in 1% SDS buffer. Proteins were separated by size electrophoretically in a 10% polyacrylamide gel, transferred to a nitrocellulose membrane, and probed for either an MMP-9, MPO, or Annexin A1 primary antibody followed by the appropriate HRP-conjugated secondary antibodies. (B) Representative Electron Microscopy images of human PMN-MPs depict size heterogeneity. An exosome &#60;100 nm in size is indicated by the white arrow. Scale bar = 250 nm (left and right panels). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56949/56949fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56949/56949fig4.jpg" /> 
Figure 4: The use of isolated PMN-MPs in studying the role of PMNs in epithelial wound healing. (A) Caco-2 BBe human intestinal epithelial cells were plated to confluency, scratch-wounded, and subjected to MPs derived from 3 million PMNs, which were added immediately post-wounding. Representative images show wound closure (48 h post-wounding) in control (left panels) and PMN-MP-treated epithelial cells (right panels). Scale bar = 100 &#181;m. (B) To examine the effect of PMN-MPs on colonic wound healing in vivo, isolated PMN-MPs were injected directly into the wound area using an endoscopy-based microinjection system (at 24 h post-wounding, the colonic wound is outlined by a dashed line and the injection needle site is shown by a white arrow). Wound closure was assessed 3 days later (4 days post-wounding) by endoscopic imaging. Scale bar = 300 &#181;m. (C) 4 days post-wounding mice were euthanized and colonic mucosal wounds were extracted with scissors, embedded in optimum cutting temperature (O.C.T) compound and frozen with liquid nitrogen. Eight micrometer sections of the wounds were stained for E-cadherin (green) and the nuclear stain DAPI (blue) to assess the level of re-epithelialization (upper panels), or for DAPI (blue) and Ki67 (red) to visualize the total and proliferating epithelial cells at the wound edge (lower panels). Scale bar = 50 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56949/56949fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Isolation and Characterization of Neutrophil-derived Microparticles for Functional Studies at JoVE.com

Isolation and Characterization of Neutrophil-derived Microparticles for Functional Studies

New protocols are described here to isolate and characterize microparticles derived from human and mouse neutrophils. These protocols utilize ultracentrifugation, flow cytometry, and immunoblotting techniques to analyze microparticle content, and they can be used to study the role of microparticles derived from various cell types in cellular function.

Polymorphonuclear neutrophil-derived microparticles (PMN)-MPs) are lipid bilayer, spherical microvesicles with sizes ranging from 50&#8211;1,000 nm in ...

isolation-characterization-neutrophil-derived-microparticles-for

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

10.1K Views.  Northwestern University. New protocols are described here to isolate and characterize microparticles derived from human and mouse neutrophils. These protocols utilize ultracentrifugation, flow cytometry, and immunoblotting techniques to analyze microparticle content, and they can be used to study the role of microparticles derived from various cell types in cellular function.

Video: Isolation and Characterization of Neutrophil-derived Microparticles for Functional Studies

Isolation of Functional Cardiac Immune Cells

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5 These immune cells, which include mast cells, T-cells and macrophages; store and release a variety of biologically active mediators including cytokines and proteases such as tryptase.6-8 These mediators have been shown to be key players in extracellular matrix metabolism by activating matrix metalloproteinases or causing collagen accumulation by modulating the cardiac fibroblasts' function.9-11 However, available techniques for isolating cardiac immune cells have been problematic because they use bacterial collagenase to digest the myocardial tissue. This technique causes activation of the immune cells and thus a loss of function. For example, cardiac mast cells become significantly less responsive to compounds that cause degranulation.12 Therefore, we developed a technique that allows for the isolation of functional cardiac immune cells which would lead to a better understanding of the role of these cells in cardiac disease.13, 14
This method requires a familiarity with the anatomical location of the rat's xiphoid process, axilla and falciform ligament, and pericardium of the heart. These landmarks are important to increase success of the procedure and to ensure a higher yield of cardiac immune cells. These isolated cardiac immune cells can then be used for characterization of functionality, phenotype, maturity, and co-culture experiments with other cardiac cells to gain a better understanding of their interactions.

This method for isolating functional immune cells from the heart provides an alternative to the conventional methods of collagenase digestion, which causes unwanted immune cell activation, resulting in a decreased responsiveness of these cells. Our method of isolation yields functional cardiac immune cells by avoiding problems associated with enzymatic digestion.

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5  These immune cells, which ...

Flow Cytometric Isolation of Primary Murine Type II Alveolar Epithelial Cells for Functional and Molecular Studies

Throughout the last years, the contribution of alveolar type II epithelial cells (AECII) to various aspects of immune regulation in the lung has been increasingly recognized. AECII have been shown to participate in cytokine production in inflamed airways and to even act as antigen-presenting cells in both infection and T-cell mediated autoimmunity 1-8. Therefore, they are especially interesting also in clinical contexts such as airway hyper-reactivity to foreign and self-antigens as well as infections that directly or indirectly target AECII. However, our understanding of the detailed immunologic functions served by alveolar type II epithelial cells in the healthy lung as well as in inflammation remains fragmentary. Many studies regarding AECII function are performed using mouse or human alveolar epithelial cell lines 9-12. Working with cell lines certainly offers a range of benefits, such as the availability of large numbers of cells for extensive analyses. However, we believe the use of primary murine AECII allows a better understanding of the role of this cell type in complex processes like infection or autoimmune inflammation. Primary murine AECII can be isolated directly from animals suffering from such respiratory conditions, meaning they have been subject to all additional extrinsic factors playing a role in the analyzed setting. As an example, viable AECII can be isolated from mice intranasally infected with influenza A virus, which primarily targets these cells for replication 13. Importantly, through ex vivo infection of AECII isolated from healthy mice, studies of the cellular responses mounted upon infection can be further extended.
Our protocol for the isolation of primary murine AECII is based on enzymatic digestion of the mouse lung followed by labeling of the resulting cell suspension with antibodies specific for CD11c, CD11b, F4/80, CD19, CD45 and CD16/CD32. Granular AECII are then identified as the unlabeled and sideward scatter high (SSChigh) cell population and are separated by fluorescence activated cell sorting 3.
In comparison to alternative methods of isolating primary epithelial cells from mouse lungs, our protocol for flow cytometric isolation of AECII by negative selection yields untouched, highly viable and pure AECII in relatively short time. Additionally, and in contrast to conventional methods of isolation by panning and depletion of lymphocytes via binding of antibody-coupled magnetic beads 14, 15, flow cytometric cell-sorting allows discrimination by means of cell size and granularity. Given that instrumentation for flow cytometric cell sorting is available, the described procedure can be applied at relatively low costs. Next to standard antibodies and enzymes for lung disintegration, no additional reagents such as magnetic beads are required. The isolated cells are suitable for a wide range of functional and molecular studies, which include in vitro culture and T-cell stimulation assays as well as transcriptome, proteome or secretome analyses 3, 4.

We describe the rapid isolation of primary murine type II alveolar epithelial cells (AECII) by flow cytometric negative selection. These AECII show high viability and purity and are suitable for a wide range of functional and molecular studies regarding their role in respiratory conditions such as autoimmune or infectious diseases.

Throughout  the last years, the contribution of alveolar type II epithelial cells (AECII)  to various aspects of immune regulation in the lung has been ...

Isolation, Purification and Labeling of Mouse Bone Marrow Neutrophils for Functional Studies and Adoptive Transfer Experiments

Neutrophils are critical effector cells of the innate immune system. They are rapidly recruited at sites of acute inflammation and exert protective or pathogenic effects depending on the inflammatory milieu. Nonetheless, despite the indispensable role of neutrophils in immunity, detailed understanding of the molecular factors that mediate neutrophils' effector and immunopathogenic effects in different infectious diseases and inflammatory conditions is still lacking, partly because of their short half life, the difficulties with handling of these cells and the lack of reliable experimental protocols for obtaining sufficient numbers of neutrophils for downstream functional studies and adoptive transfer experiments. Therefore, simple, fast, economical and reliable methods are highly desirable for harvesting sufficient numbers of mouse neutrophils for assessing functions such as phagocytosis, killing, cytokine production, degranulation and trafficking. To that end, we present a reproducible density gradient centrifugation-based protocol, which can be adapted in any laboratory to isolate large numbers of neutrophils from the bone marrow of mice with high purity and viability. Moreover, we present a simple protocol that uses CellTracker dyes to label the isolated neutrophils, which can then be adoptively transferred into recipient mice and tracked in several tissues for at least 4 hr post-transfer using flow cytometry. Using this approach, differential labeling of neutrophils from wild-type and gene-deficient mice with different CellTracker dyes can be successfully employed to perform competitive repopulation studies for evaluating the direct role of specific genes in trafficking of neutrophils from the blood into target tissues in vivo.

We describe a protocol for isolation and purification of neutrophils from mouse bone marrow by density gradient centrifugation and for neutrophil labeling using CellTracker dyes. This represents a simple, fast, reproducible and economical method for obtaining large numbers of neutrophils for downstream functional studies or adoptive transfer and tracking experiments.

Neutrophils are critical effector cells of the innate immune system. They are rapidly recruited at sites of acute inflammation and exert protective or ...

Analysis of the Epithelial Damage Produced by Entamoeba histolytica Infection

Entamoeba histolytica is the causative agent of human amoebiasis, a major cause of diarrhea and hepatic abscess in tropical countries. Infection is initiated by interaction of the pathogen with intestinal epithelial cells. This interaction leads to disruption of intercellular structures such as tight junctions (TJ). TJ ensure sealing of the epithelial layer to separate host tissue from gut lumen. Recent studies provide evidence that disruption of TJ by the parasitic protein EhCPADH112 is a prerequisite for E. histolytica invasion that is accompanied by epithelial barrier dysfunction. Thus, the analysis of molecular mechanisms involved in TJ disassembly during E. histolytica invasion is of paramount importance to improve our understanding of amoebiasis pathogenesis. This article presents an easy model that allows the assessment of initial host-pathogen interactions and the parasite invasion potential. Parameters to be analyzed include transepithelial electrical resistance, interaction of EhCPADH112 with epithelial surface receptors, changes in expression and localization of epithelial junctional markers and localization of parasite molecules within epithelial cells.

Analysis of the Epithelial Damage Produced by Entamoeba histolytica Infection

Human infection by Entamoeba histolytica leads to amoebiasis, a major cause of diarrhea in tropical countries. Infection is initiated by pathogen interactions with intestinal epithelial cells, provoking the opening of cell-cell contacts and consequently diarrhea, sometimes followed by liver infection. This article provides a model to assess the early host-pathogen interactions to improve our understanding of amoebiasis pathogenesis.

Entamoeba histolytica is the causative agent of human amoebiasis, a major cause of diarrhea and hepatic abscess in tropical countries. Infection is ...

Detecting Glycogen in Peripheral Blood Mononuclear Cells with Periodic Acid Schiff Staining

Periodic acid Schiff (PAS) staining is an immunohistochemical technique used on muscle biopsies and as a diagnostic tool for blood samples. Polysaccharides such as glycogen, glycoproteins, and glycolipids stain bright magenta making it easy to enumerate positive and negative cells within the tissue. In muscle cells PAS staining is used to determine the glycogen content in different types of muscle cells, while in blood cell samples PAS staining has been explored as a diagnostic tool for a variety of conditions. Blood contains a proportion of white blood cells that belong to the immune system. The notion that cells of the immune system possess glycogen and use it as an energy source has not been widely explored. Here, we describe an adapted version of the PAS staining protocol that can be applied on peripheral blood mononuclear immune cells from human venous blood. Small cells with PAS-positive granules and larger cells with diffuse PAS staining were observed. Treatment of samples with amylase abrogates these patterns confirming the specificity of the stain. An alternate technique based on enzymatic digestion confirmed the presence and amount of glycogen in the samples. This protocol is useful for hematologists or immunologists studying polysaccharide content in blood-derived lymphocytes.

Periodic acid Schiff staining is a technique that visualizes the polysaccharide content of tissues. This article demonstrates periodic acid Schiff staining protocol adapted for use on peripheral blood mononuclear cells purified from human venous blood. Such samples are enriched for lymphocytes and other white blood cells of the immune system.

Periodic acid Schiff (PAS) staining is an immunohistochemical technique used on muscle biopsies and as a diagnostic tool for blood samples. ...

Isolation and Characterization of Neutrophils with Anti-Tumor Properties

Neutrophils, the most abundant of all white blood cells in the human circulation, play an important role in the host defense against invading microorganisms. In addition, neutrophils play a central role in the immune surveillance of tumor cells. They have the ability to recognize tumor cells and induce tumor cell death either through a cell contact-dependent mechanism involving hydrogen peroxide or through antibody-dependent cell-mediated cytotoxicity (ADCC). Neutrophils with anti-tumor activity can be isolated from peripheral blood of cancer patients and of tumor-bearing mice. These neutrophils are termed tumor-entrained neutrophils (TEN) to distinguish them from neutrophils of healthy subjects or na&#239;ve mice that show no significant tumor cytotoxic activity. Compared with other white blood cells, neutrophils show different buoyancy making it feasible to obtain a &#62; 98% pure neutrophil population when subjected to a density gradient. However, in addition to the normal high-density neutrophil population (HDN), in cancer patients, in tumor-bearing mice, as well as under chronic inflammatory conditions, distinct low-density neutrophil populations (LDN) appear in the circulation. LDN co-purify with the mononuclear fraction and can be separated from mononuclear cells using either positive or negative selection strategies. Once the purity of the isolated neutrophils is determined by flow cytometry, they can be used for in vitro and in vivo functional assays. We describe techniques for monitoring the anti-tumor activity of neutrophils, their ability to migrate and to produce reactive oxygen species, as well as monitoring their phagocytic capacity ex vivo. We further describe techniques to label the neutrophils for in vivo tracking, and to determine their anti-metastatic capacity in vivo. All these techniques are essential for understanding how to obtain and characterize neutrophils with anti-tumor function.

Neutrophils play an important role not only in host defense against invading microorganisms, but are also involved in the immune surveillance of tumor cells. Here, we describe techniques related to the isolation of neutrophils with anti-tumor properties and methods for monitoring anti-tumor neutrophil function in vitro and in vivo.

Neutrophils, the most abundant of all white blood cells in the human circulation, play an important role in the host defense against invading ...

A 3D Human Lung Tissue Model for Functional Studies on Mycobacterium tuberculosis Infection

Tuberculosis (TB) still holds a major threat to the health of people worldwide, and there is a need for cost-efficient but reliable models to help us understand the disease mechanisms and advance the discoveries of new treatment options. In vitro cell cultures of monolayers or co-cultures lack the three-dimensional (3D) environment and tissue responses. Herein, we describe an innovative in vitro model of a human lung tissue, which holds promise to be an effective tool for studying the complex events that occur during infection with Mycobacterium tuberculosis (M. tuberculosis). The 3D tissue model consists of tissue-specific epithelial cells and fibroblasts, which are cultured in a matrix of collagen on top of a porous membrane. Upon air exposure, the epithelial cells stratify and secrete mucus at the apical side. By introducing human primary macrophages infected with M. tuberculosis to the tissue model, we have shown that immune cells migrate into the infected-tissue and form early stages of TB granuloma. These structures recapitulate the distinct feature of human TB, the granuloma, which is fundamentally different or not commonly observed in widely used experimental animal models. This organotypic culture method enables the 3D visualization and robust quantitative analysis that provides pivotal information on spatial and temporal features of host cell-pathogen interactions. Taken together, the lung tissue model provides a physiologically relevant tissue micro-environment for studies on TB. Thus, the lung tissue model has potential implications for both basic mechanistic and applied studies. Importantly, the model allows addition or manipulation of individual cell types, which thereby widens its use for modelling a variety of infectious diseases that affect the lungs.

A 3D Human Lung Tissue Model for Functional Studies on Mycobacterium tuberculosis Infection

Human tuberculosis infection is a complex process, which is difficult to model in vitro. Here we describe a novel 3D human lung tissue model that recapitulates the dynamics that occur during infection, including the migration of immune cells and early granuloma formation in a physiological environment.

Tuberculosis (TB) still holds a major threat to the health of people worldwide, and there is a need for cost-efficient but reliable models to help us ...

Isolation, Characterization and Functional Examination of the Gingival Immune Cell Network

Immune cell networks in tissues play a vital role in mediating local immunity and maintaining tissue homeostasis, yet little is known of the resident immune cell populations in the oral mucosa and gingiva. We have established a technique for the isolation and study of immune cells from murine gingival tissues, an area of constant microbial exposure and a vulnerable site to a common inflammatory disease, periodontitis. Our protocol allows for a detailed phenotypic characterization of the immune cell populations resident in the gingiva, even at steady state. Our procedure also yields sufficient cells with high viability for use in functional studies, such as the assessment of cytokine secretion ex vivo. This combination of phenotypic and functional characterization of the gingival immune cell network should aid towards investigating the mechanisms involved in oral immunity and periodontal homeostasis, but will also advance our understanding of the mechanisms involved in local immunopathology.

We have established a technique for the isolation, phenotypic characterization and functional analysis of immune cells from murine gingiva.

Immune cell networks in tissues play a vital role in mediating local immunity and maintaining tissue homeostasis, yet little is known of the resident ...

Isolation and Flow Cytometric Characterization of Murine Small Intestinal Lymphocytes

The intestines &#8212; which contain the largest number of immune cells of any organ in the body &#8212; are constantly exposed to foreign antigens, both microbial and dietary. Given an increasing understanding that these luminal antigens help shape the immune response and that education of immune cells within the intestine is critical for a number of systemic diseases, there has been increased interest in characterizing the intestinal immune system. However, many published protocols are arduous and time-consuming. We present here a simplified protocol for the isolation of lymphocytes from the small-intestinal lamina propria, intraepithelial layer, and Peyer&#39;s patches that is rapid, reproducible, and does not require laborious Percoll gradients. Although the protocol focuses on the small intestine, it is also suitable for analysis of the colon. Moreover, we highlight some aspects that may need additional optimization depending on the specific scientific question. This approach results in the isolation of large numbers of viable lymphocytes that can subsequently be used for flow cytometric analysis or alternate means of characterization.

There is growing interest in the quantitative characterization of intestinal lymphocytes owing to increasing recognition that these cells play a critical role in a variety of intestinal and systemic diseases. In this protocol, we describe how to isolate single cell populations from different small-intestinal compartments for subsequent flow cytometric characterization.

The intestines &#8212; which contain the largest number of immune cells of any organ in the body &#8212; are constantly exposed to foreign antigens, both ...

Functional Characterization of Regulatory Macrophages That Inhibit Graft-reactive Immunity

Macrophage accumulation in transplanted organs has long been recognized as a feature of allograft rejection1. Immunogenic monocytes infiltrate the allograft early after transplantation, mount a graft reactive response against the transplanted organ, and initiate organ rejection2. Recent data suggest that suppressive macrophages facilitate successful long-term transplantation3 and are required for the induction of transplantation tolerance4. This suggests a multidimensional concept of macrophage ontogeny, activation, and function, which demands a new roadmap for the isolation and analysis of macrophage function5. Due to the plasticity of macrophages, it is necessary to provide a methodology to isolate and characterize macrophages, depending on the tissue environment, and to define their functions according to different scenarios. Here, we describe a protocol for immune characterization of graft-infiltrating macrophages and the methods we used to functionally evaluate their capacity to inhibit CD8+ T proliferation and to promote CD4+Foxp3+ Treg expansion in vitro.

Macrophages are plastic cells of the hematopoietic system that have a crucial role in protective immunity and homeostasis. In this report, we describe optimized in vitro techniques to phenotypically and functionally characterize graft-infiltrating regulatory macrophages that accumulate in the transplanted organ under tolerogenic conditions.

Isolation and Characterization of Neutrophil-derived Microparticles for Functional Studies

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Isolation of Functional Cardiac Immune Cells

Flow Cytometric Isolation of Primary Murine Type II Alveolar Epithelial Cells for Functional and Molecular Studies

Isolation, Purification and Labeling of Mouse Bone Marrow Neutrophils for Functional Studies and Adoptive Transfer Experiments

Analysis of the Epithelial Damage Produced by Entamoeba histolytica Infection

Detecting Glycogen in Peripheral Blood Mononuclear Cells with Periodic Acid Schiff Staining

Isolation and Characterization of Neutrophils with Anti-Tumor Properties

A 3D Human Lung Tissue Model for Functional Studies on Mycobacterium tuberculosis Infection

Isolation, Characterization and Functional Examination of the Gingival Immune Cell Network

Isolation and Flow Cytometric Characterization of Murine Small Intestinal Lymphocytes

Functional Characterization of Regulatory Macrophages That Inhibit Graft-reactive Immunity