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

Universidad San Sebastian

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

Neutrophil Isolation Protocol

Neutrophil polymorphonuclear granulocytes (PMN) are the most abundant leukocytes in humans and among the first cells to arrive on the site of inflammatory immune response. Due to their key role in inflammation, neutrophil functions such as locomotion, cytokine production, phagocytosis, and tumor cell combat are extensively studied. To characterize the specific functions of neutrophils, a clean, fast, and reliable method of separating them from other blood cells is desirable for in vitro studies, especially since neutrophils are short-lived and should be used within 2-4 hours of collection. Here, we demonstrate a standard density gradient separation method to isolate human neutrophils from whole blood using commercially available separation media that is a mixture of sodium metrizoate and Dextran 500. The procedure consists of layering whole blood over the density gradient medium, centrifugation, separation of neutrophil layer, and lysis of residual erythrocytes. Cells are then washed, counted, and resuspended in buffer to desired concentration. When performed correctly, this method has been shown to yield samples of >95% neutrophils with >95% viability.

Neutrophils are among the first cells to arrive on the site of inflammatory immune response, and their functions and mechanisms have been studied extensively in vitro. We demonstrate a standard density gradient separation method to isolate human neutrophils from whole blood using commercially available separation media.

Neutrophil polymorphonuclear granulocytes (PMN) are the most abundant leukocytes in humans and among the first cells to arrive on the site of inflammatory ...

Biology

Microfluidic Platform for Measuring Neutrophil Chemotaxis from Unprocessed Whole Blood

Neutrophils play an essential role in protection against infections and their numbers in the blood are frequently measured in the clinic. Higher neutrophil counts in the blood are usually an indicator of ongoing infections, while low neutrophil counts are a warning sign for higher risks for infections. To accomplish their functions, neutrophils also have to be able to move effectively from the blood where they spend most of their life, into tissues, where infections occur. Consequently, any defects in the ability of neutrophils to migrate can increase the risks for infections, even when neutrophils are present in appropriate numbers in the blood. However, measuring neutrophil migration ability in the clinic is a challenging task, which is time consuming, requires large volume of blood, and expert knowledge. To address these limitations, we designed a robust microfluidic assays for neutrophil migration, which requires a single droplet of unprocessed blood, circumvents the need for neutrophil separation, and is easy to quantify on a simple microscope. In this assay, neutrophils migrate directly from the blood droplet, through small channels, towards the source of chemoattractant. To prevent the granular flow of red blood cells through the same channels, we implemented mechanical filters with right angle turns that selectively block the advance of red blood cells. We validated the assay by comparing neutrophil migration from blood droplets collected from finger prick and venous blood. We also compared these whole blood (WB) sources with neutrophil migration from samples of purified neutrophils and found consistent speed and directionality between the three sources. This microfluidic platform will enable the study of human neutrophil migration in the clinic and the research setting to help advance our understanding of neutrophil functions in health and disease.

This protocol details an assay designed to measure human neutrophil chemotaxis from one droplet of whole blood with robust reproducibility. This approach circumvents the need for neutrophil separation and requires only a few minutes of assay preparation time. The microfluidic chip enables the repeated measure of neutrophil chemotaxis over time in infants or small mammals, where sample volume is limited.

Neutrophils play an essential role in protection against infections and their numbers in the blood are frequently measured in the clinic. Higher ...

Bioengineering

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 ...

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 ...

Neutrophil Isolation and Analysis to Determine their Role in Lymphoma Cell Sensitivity to Therapeutic Agents

Neutrophils are the most abundant (40% to 75%) type of white blood cells and among the first inflammatory cells to migrate towards the site of inflammation. They are key players in the innate immune system and play major roles in cancer biology. Neutrophils have been proposed as key mediators of malignant transformation, tumor progression, angiogenesis and in the modulation of the antitumor immunity; through their release of soluble factors or their interaction with tumor cells. To characterize the specific functions of neutrophils, a fast and reliable method is coveted for in vitro isolation of neutrophils from human blood. Here, a density gradient separation method is demonstrated to isolate neutrophils as well as mononuclear cells from the blood. The procedure consists of layering the density gradient solution such as Ficoll carefully above the diluted blood obtained from patients diagnosed with chronic lymphocytic leukemia (CLL), followed by centrifugation, isolation of mononuclear layer, separation of neutrophils from RBCsby dextran then lysis of residual erythrocytes. This method has been shown to isolate neutrophils &#8805; 90 % pure. To mimic the tumor microenvironment, 3-dimensional (3D) experiments were performed using basement membrane matrix such as Matrigel. Given the short half-life of neutrophils in vitro, 3D experiments with fresh human neutrophils cannot be performed. For this reason promyelocytic HL60 cells are differentiated along the granulocytic pathway using the differentiation inducers dimethyl sulfoxide (DMSO) and retinoic acid (RA). The aim of our experiments is to study the role of neutrophils on the sensitivity of lymphoma cells to anti-lymphoma agents. However these methods can be generalized to study the interactions of neutrophils or neutrophil-like cells with a large range of cell types in different situations.

Neutrophils are the most abundant type of white blood cells. The isolation of neutrophils from human blood by density gradient separation method and the differentiation of human promyelocytic (HL60) cells along the granulocytic pathway are described here; to test their role on sensitivity of lymphoma cells to anti-lymphoma agents.

Neutrophils are the most abundant (40% to 75%) type of white blood cells and among the first inflammatory cells to migrate towards the site of ...

Development and Identification of a Novel Subpopulation of Human Neutrophil-derived Giant Phagocytes In Vitro

Neutrophils (PMN) are best known for their phagocytic functions against invading pathogens and microorganisms. They have the shortest half-life amongst leukocytes and in their non-activated state are constitutively committed to apoptosis. When recruited to inflammatory sites to resolve inflammation, they produce an array of cytotoxic molecules with potent antimicrobial killing. Yet, when these powerful cytotoxic molecules are released in an uncontrolled manner they can damage surrounding tissues. In recent years however, neutrophil versatility is increasingly evidenced, by demonstrating plasticity and immunoregulatory functions. We have recently identified a new neutrophil-derived subpopulation, which develops spontaneously in standard culture conditions without the addition of cytokines/growth factors such as granulocyte colony-stimulating factor (GM-CSF)/interleukin (IL)-4. Their phagocytic abilities of neutrophil remnants largely contribute to increase their size immensely; therefore they were termed giant phagocytes (G&#981;). Unlike neutrophils, G&#981; are long lived in culture. They express the cluster of differentiation (CD) neutrophil markers CD66b/CD63/CD15/CD11b/myeloperoxidase (MPO)/neutrophil elastase (NE), and are devoid of the monocytic lineage markers CD14/CD16/CD163 and the dendritic CD1c/CD141 markers. They also take-up latex and zymosan, and respond by oxidative burst to stimulation with opsonized-zymosan and PMA. G&#981; also express the scavenger receptors CD68/CD36, and unlike neutrophils, internalize oxidized-low density lipoprotein (oxLDL). Moreover, unlike fresh neutrophils, or cultured monocytes, they respond to oxLDL uptake by increased reactive oxygen species (ROS) production. Additionally, these phagocytes contain microtubule-associated protein-1 light chain 3B (LC3B) coated vacuoles, indicating the activation of autophagy. Using specific inhibitors it is evident that both phagocytosis and autophagy are prerequisites for their development and likely NADPH oxidase dependent ROS. We describe here a method for the preparation of this new subpopulation of long-lived, neutrophil-derived phagocytic cells in culture, their identification and their currently known characteristics. This protocol is essential for obtaining and characterizing G&#981; in order to further investigate their significance and functions.

Development and Identification of a Novel Subpopulation of Human Neutrophil-derived Giant Phagocytes In Vitro

We describe here a method for obtaining and identifying a newly characterized subpopulation of neutrophil-derived giant phagocytes. These cells develop in culture from fresh human blood neutrophils, and are characterized by phagocytosis, autophagy, immensely large size, and extended lifespan. This method is essential to further investigate this unique neutrophil-derived subpopulation.

Neutrophils (PMN) are best known for their phagocytic functions against invading pathogens and microorganisms. They have the shortest half-life amongst ...

Label-free Neutrophil Enrichment from Patient-derived Airway Secretion Using Closed-loop Inertial Microfluidics

Airway secretions contain a large number of immune-related cells, e.g., neutrophils, macrophages, and lymphocytes, which can be used as a major resource to evaluate a variety of pulmonary diseases, both for research and clinical purposes. However, due to the heterogeneous and viscous nature of patient mucus, there is currently no reliable dissociation method that does not damage the host immune cells in the patient airway secretion. In this research, we introduce a sample preparation method that uses inertial microfluidics for the patient&#39;s immune assessment. Regardless of the heterogeneous fluidic properties of the clinical samples, the proposed method recovers more than 95% of neutrophils from airway secretion samples that are diluted 1,000-fold with milliliters of clean saline. By recirculating the concentrated output stream to the initial sample reservoir, a high concentration, recovery, and purity of the immune cells are provided; recirculation is considered a trade-off to the single-run syringe-based operation of inertial microfluidics. The closed-loop operation of spiral microfluidics provides leukocytes without physical or chemical disturbance, as demonstrated by the phorbol 12-myristate 13-acetate (PMA)-induced elastase release of sorted neutrophils.

In this research, we demonstrate a label-free neutrophil separation method from clinical airway secretions using closed-loop operation of spiral inertial microfluidics. The proposed method would expand the clinical in vitro assays for various respiratory diseases.

Airway secretions contain a large number of immune-related cells, e.g., neutrophils, macrophages, and lymphocytes, which can be used as a major resource ...

Neutrophil Functional Assays for Screening Different Signaling Pathways

A Set of Screening Techniques for a Quick Overview of the Neutrophil Function

Neutrophils are known as one of the first lines of defense in the innate immune response and can perform many particular cellular functions, such as chemotaxis, reverse migration, phagocytosis, degranulation of cytotoxic enzymes and metabolites, and release of DNA as neutrophil extracellular traps (NETs). Neutrophils not only have tightly regulated signaling themselves, but also participate in the regulation of other components of the immune system. As fresh neutrophils are terminally differentiated, short-lived, and highly variable among individuals, it is important to make the most of the collected samples. Researchers often need to perform screening assays to assess an overview of the many neutrophil functions that may be affected by specific conditions under evaluation. A set of tests following a single isolation process of normal density neutrophils was developed to address this need, seeking a balance between speed, comprehensiveness, cost, and accuracy. The results can be used to reason and guide in-depth follow-up studies. This procedure can be carried out in an average time of 4 h and includes the evaluation of cell viability, reactive oxygen species (ROS) production, real-time migration, and phagocytosis of yeast on glass slides, leaving enough cells for more detailed approaches like omics studies. Moreover, the procedure includes a way to easily observe a preliminary suggestion of NETs after fast panoptic staining observed by light microscopy, with a lack of specific markers, albeit enough to indicate if further efforts in that way would be worthwhile. The diversity of functions tested combines common points among tests, reducing the analysis time and expenses. The procedure was named NeutroFun Screen, and although having&#160;limitations, it balances the aforementioned factors. Furthermore, the aim of this work is not a definite test set, but rather a guideline that can easily be adjusted to each lab&#39;s resources and demands.

Author Spotlight: Balancing Speed, Cost, and Accuracy in Neutrophil Function Assessment with NeutroFun Screen Protocol 

This protocol features a set of neutrophil functional assays to be used as a screening method to cover functions from different signaling pathways. The protocol includes an initial and simple evaluation of cell viability, purity, reactive oxygen species production, real-time migration, phagocytosis, and a preliminary suggestion of neutrophil extracellular traps.

Neutrophils are known as one of the first lines of defense in the innate immune response and can perform many particular cellular functions, such as ...

The Isolation and Characterization of Low- and Normal- Density Neutrophils from Whole Blood

Emerging research shows that the circulating neutrophil population in humans consists of diverse subtypes and should not be studied as a single population, as has been done historically. In particular, low-density and normal-density neutrophils (LDNs, NDNs) have been shown to have functionally and metabolically distinct profiles, a factor that must be considered when publishing neutrophil research. Here, we present a modified method for the untouched isolation and separation of LDNs and NDNs from whole blood.
The density gradient medium (1.135 g/mL) is combined at 9:10 with 10x PBS. Specific density gradients of 55%, 70%, and 81% are subsequently made by combining the 100% density gradient medium with 1x phosphate-buffered saline (PBS). Neutrophils isolated from 12 mL of peripheral whole blood obtained from consented donors using a negative selection-based magnetic isolation kit are resuspended in the 55% fraction. A volume of 3 mL of the 81% and 70% fractions is layered into a 15 mL tube, followed by the 55% fraction containing total neutrophils. The density gradients are then centrifuged at 720 x&#160;g for 30 min. Two distinct bands are obtained at the 55%/70% interface (LDNs) and 70%/81% interface (NDNs). The cells are carefully pipetted into separate tubes and washed using PBS. The purity of the isolated fractions is determined using flow cytometry. Both LDNs and NDNs were defined as CD14lo CD15+ SSChi by flow cytometry. Isolation purity was calculated at &#8805;93% of viable cells for both types.
This method provides a reliable and efficient approach for separating LDN and NDNs from peripheral blood, ensuring high purity and viability of the isolated cells. Enhancing the precision of neutrophil isolation facilitates more accurate downstream analyses of these distinct neutrophil subpopulations. These are critical for advancing our understanding of neutrophil heterogeneity and its implications in various physiological and pathological contexts.

Here, we provide a reliable approach for isolating low- and normal-density neutrophils from whole blood using magnetic isolation (negative selection) and discontinuous density gradient medium. It ensures untouched isolation of high-purity cells (&#8805;93%), facilitating accurate downstream analysis of neutrophil subpopulations, crucial for understanding their roles in health and disease.

Emerging research shows that the circulating neutrophil population in humans consists of diverse subtypes and should not be studied as a single ...

Isolation of Human Neutrophils from Whole Blood and Buffy Coats

Neutrophils (PMNs) are the most abundant leukocytes in human circulation, ranging from 40 to 70% of total blood leukocytes. They are the first cells recruited at the site of inflammation via rapid extravasation through vessels. There, neutrophils perform an array of functions to kill invading pathogens and mediate immune signaling. Freshly purified neutrophils from human blood are the model of choice for study, as no cell line fully replicates PMN functions and biology. However, neutrophils are short-lived, terminally differentiated cells and are highly susceptible to activation in response to physical (temperature, centrifugation speed) and biological (endotoxin, chemo- and cytokines) stimuli. Therefore, it is crucial to follow a standardized, reliable, and fast method to obtain pure and non-activated cells. This protocol presents an updated protocol combining density gradient centrifugation, red blood cell (RBC) sedimentation, and RBC lysis to obtain high PMN purity and minimize cell activation. Furthermore, methods to assess neutrophil isolation quality, viability, and purity are also discussed.

This protocol details a method for the isolation of neutrophils from whole blood, buffy coats, or leukapheresis membranes, achieving good yield, high purity, and minimal cell activation. We utilize gradient purification, red blood cell (RBC) sedimentation, and RBC lysis to obtain a high-quality/purity neutrophil preparation.

Neutrophils (PMNs) are the most abundant leukocytes in human circulation, ranging from 40 to 70% of total blood leukocytes. They are the first cells ...

Isolation and Characterization of Neutrophil-derived Microparticles for Functional Studies

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The Isolation and Characterization of Low- and Normal- Density Neutrophils from Whole Blood

Isolation of Human Neutrophils from Whole Blood and Buffy Coats