This work was supported by funding from the William G. Lowrie Department of Chemical and Biomolecular Engineering and the Comprehensive Cancer Center at The Ohio State University. Data presented in this report came from images processed using Imaris x64 (ver. 9.3.0 Bitplane) available at the Campus Microscopy and Imaging Facility, The Ohio State University. This facility is supported in part by grant P30 CA016058, National Cancer Institute, Bethesda, MD.

The authors acknowledge the healthy volunteers who kindly donated their blood. Blood specimens were obtained after informed volunteer consent according to institutional review board (IRB) protocol #2018H0268 reviewed by the Biomedical Sciences Committee at The Ohio State University.
1. Microfabrication of bioparticle microarray
<ol>
	<li>Using standard photolithography procedures, generate the master silicon wafer.
	<ol>
		<li>Generate a proof of the desired design using a computer-aided design (CAD) software, then send to a photomask manufacturer to produce a chrome photomask. The design used here is 4 mm x 4 mm rectangular arrays of 30 &#181;m diameter filled in circles with a 150 &#181;m center-to-center spacing. This design can be modified as desired for different applications.</li>
		<li>Spincoat a 40 &#181;m thick layer of a negative photoresist onto a silicon wafer. Bake wafer at 65 &#176;C for 5 min and 95 &#176;C for 10 min.</li>
		<li>Expose wafer to UV light through a chrome photomask with 150&#8211;160 mJ/cm2 (Figure 1A).</li>
		<li>Bake wafer at 65 &#176;C for 5 min and 95 &#176;C for 10 min. Submerge wafer in photoresist developer for 10 min and rinse with isopropyl alcohol. At this stage, the pattern should be visible on the wafer (Figure 1B).</li>
	</ol>
	</li>
	<li>Thoroughly mix a 10:1 ratio of polydimethylsiloxane prepolymer and its curing agent (i.e., 20 g of prepolymer and 2 g of curing agent) and pour the uncured polydimethylsiloxane (PDMS) mixture over the master wafer in a Petri dish (Figure 1C).</li>
	<li>Vacuum treat the uncured PDMS mixture until no air bubbles are present over the master wafer. Cure at 65 &#176;C overnight.</li>
	<li>Use a scalpel to cut around the exterior of the patterned section of the wafer, and slowly remove the cured PDMS slab. Place the PDMS slab on a clean cutting board with the patterned side facing up (Figure 1D).</li>
	<li>Punch out individual stamps from the PDMS slab with an 8 mm biopsy punch (Figure 1E). For one glass slide, eight stamps will be needed.</li>
	<li>Place each stamp face down on the adhesive tape to remove any debris.</li>
	<li>In advance, prepare a 1.6 mg/mL solution of CP in water.
	<ol>
		<li>Add the proper amount of the CP powder to water (e.g., 0.8 g to 500 mL).</li>
		<li>Mix on a stir plate at room temperature overnight, or until all the solid is dissolved into the water. The CP solution can be stored at room temperature for 6 months.</li>
		<li>If desired, make the CP solution fluorescent by adding poly-L-lysine labelled with fluorescein isothiocyanate (PLL-FITC).
		<ol>
			<li>Aliquot about 10 mL of the CP solution. Add a small amount of PLL-FITC (0.05 mg) to the aliquoted volume. The amount can be altered to adjust the brightness of fluorescence as desired.</li>
			<li>Vortex the CP solution labeled with FITC for 20 s, or until the solution is a uniform, pale yellow color. Protect from light and store at 4 &#176;C for up to 1 month.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>With the stamps face-up, prime each stamp with 100 &#181;L of 1.6 mg/mL solution of CP, ensuring no air bubbles form between the CP solution and the stamp (Figure 1F).</li>
	<li>Invert the stamps onto a layer of CP solution (Figure 1G).</li>
	<li>Remove the stamps from the CP solution after 1 h.</li>
	<li>Dab each wet stamp face down onto a clean glass slide 6&#8211;8x to remove excess liquid.</li>
	<li>Vacuum treat the stamps for 1&#8211;2 min.</li>
	<li>Adhere an eight well, 9 mm diameter imaging spacer on the top of a clean glass slide as a guide for stamp placement. With this spacer, each glass slide can have eight microarrays.</li>
	<li>Place a stamp face down on the glass slide in the center of each well of the imaging spacer (i.e., use eight stamps total).</li>
	<li>Place a 5.6 &#177; 0.1 g balanced weight on top of each stamp and allow 10 min for stamping (Figure 1H). A balanced weight is required to ensure the stamp is pressed evenly onto the glass slide and promote even transfer of the CP to the glass slide.</li>
	<li>Remove the weights and stamps from the glass slide (Figure 1I). Allow the CP layer to dry at room temperature for 24 h before adding the bioparticles, as described in step 1.17. If the CP is tagged with FITC, the effectiveness of the stamping can be checked at this point with a fluorescent microscope at 488 nm before proceeding to step 1.17 (Figure 1M).</li>
	<li>Cut a blank PDMS slab to the size of the imaging spacer and use the 8 mm biopsy punch to create wells in the PDMS that align with the wells of an imaging spacer. Adhere the PDMS slab to the glass slide with the imaging spacer (Figure 1J).</li>
	<li>Thaw a solution of bioparticles (e.g., E. coli or zymosan) and dilute to 500 &#956;g/mL in water for injection (WFI). 
	NOTE: The bioparticles do not need to be opsonized. Neutrophil surface receptors directly recognize molecules on these bioparticles19,20,21.</li>
	<li>Add 100 &#956;L of bioparticle solution to each PDMS well on the glass slide (Figure 1K).</li>
	<li>Rock the glass slide for 30 min.</li>
	<li>Rinse the wells thoroughly with water. The bioparticle microarray can be stored in a dust-free environment at 4 &#176;C for up to 3 months. At this point, the pattern should be checked with a fluorescent microscope at 594 nm before proceeding to step 2.1 (Figure 1N).</li>
</ol>
2. Sample Preparation
<ol>
	<li>Collect at least 2 mL of fresh blood in K2-EDTA tubes from the desired donor. The expected yield of neutrophils is 1&#8211;2 x 106 cells/1 mL whole blood. The imaging assay requires approximately 1.5 x 105 neutrophils, and the analysis of the supernatant requires 1 x 106 neutrophils. Use the blood within 4 h.</li>
	<li>Separate red blood cells (RBCs) by adding an erythrocyte aggregation agent in a 1:5 ratio to the whole blood. Wait 45 min for a translucent layer (buffy coat) to separate from the layer of RBCs.</li>
	<li>Remove the buffy coat and wash with phosphate buffered saline (PBS) using 1 mL buffy coat: 9 mL PBS.</li>
	<li>Centrifuge for 5 min at 190 x g and 20 &#176;C.</li>
	<li>Aspirate the supernatant and resuspend the pellet at 5 x 107 cells/mL.</li>
	<li>Use a negative immunomagnetic selection kit to isolate neutrophils.
	<ol>
		<li>Add 50 &#956;L of antibody cocktail/1 mL cell suspension. Wait 10 min.</li>
		<li>Add 100 &#956;L of magnetic beads/1 mL cell suspension. Wait 10 min.</li>
		<li>Add cell suspension to a round-bottom tube and place in a cylindrical magnet. Wait 10 min.</li>
	</ol>
	</li>
	<li>Pour supernatant into a centrifuge tube. Add up to 10 mL of PBS. Centrifuge for 5 min at 190 x g and 20 &#176;C.</li>
	<li>Aspirate supernatant. Resuspend white pellet in IMDM with 0.4% human serum albumin.</li>
	<li>Stain nuclei with 20 &#181;g/mL Hoechst 33342 for 10 min at 37 &#176;C.</li>
	<li>Add 5 mL of IMDM with 0.4% human serum albumin to rinse. Centrifuge for 5 min at 190 x g and 20 &#176;C.</li>
	<li>Resuspend cells at 7.5 x 105 cells/mL in IMDM with 0.4% human serum albumin.</li>
	<li>Add 100 &#181;L of the cell suspension to a PDMS well containing a bioparticle microarray. Ensure the cell suspension is convex over the top of the PDMS well and does not contain any bubbles.</li>
	<li>Seal with a 12 mm diameter coverslip. Cover the opening of the PDMS well with a 12 mm diameter coverslip. Press down gently onto the coverslip with tweezers so the excess cell suspension escapes to the edge of the well. Use a tissue to remove the excess cell suspension.</li>
</ol>
3. Running the assay and image analysis
<ol>
	<li>Load microparticle array with cells on the live cell imaging station with a microscope equipped with a cage incubator set to 37 &#176;C, 5% CO2, and 90% relative humidity.</li>
	<li>Use time-lapse fluorescent and brightfield microscopy to record images at 10x magnification every 10 s at 405 nm, 594 nm, and brightfield. In a typical experiment, images are collected up to 2 h.</li>
	<li>Use an automated cell tracking software to track the migration of individual neutrophils toward the bioparticle cluster.
	<ol>
		<li>Use the autoregression mode of a spot detection cell tracking software. Set the spot radius to 5 &#181;m (the approximate size of a neutrophil nucleus). Set the minimum track length to 120 s and a maximum gap size of one frame.</li>
		<li>From the data generated by the cell tracking software, extract the files that contain the neutrophil position and speed. These files can be used with a graphing software to generate neutrophil migration tracks (Figure 2C) and a heat map of speed vs. time (Figure 2D), respectively.</li>
	</ol>
	</li>
	<li>Use the 405 nm fluorescent images to track swarm size over time on an image analysis software of your choice.
	<ol>
		<li>Define regions of interest (ROIs) around each bioparticle cluster where neutrophils will swarm. Keep the same size ROI to analyze each bioparticle cluster.</li>
		<li>Analyze the mean fluorescent intensity of the 405 nm images within each ROI over time.</li>
		<li>Generate a calibration curve of mean fluorescent intensity to swarm size by taking manual measurements at various swarm sizes from 0 &#181;m2 to the maximum swarm size. Use this calibration to calculate the swarm size over time.</li>
	</ol>
	</li>
</ol>
4. Supernatant collection and protein detection
<ol>
	<li>Incubate the neutrophils in the wells containing the bioparticle microarray at 37 &#176;C and 5% CO2 for 3 h. Take samples at desired time points. Typically, samples will be taken at 0, 0.5, 1, and 3 h. To overcome the limit of detection of the protein array assay, the entire volume of supernatant of a single well (200 &#181;L) was used for each time point. Each time point was analyzed in triplicate.</li>
	<li>Using a brightfield microscope, verify that swarms are formed on the microarray.</li>
	<li>Aspirate the supernatant with a 200 &#181;L pipette and load in a 0.45 &#956;m centrifuge filter tube.</li>
	<li>Centrifuge the supernatant at 190 x g and 20 &#176;C for 5 min and collect the filtrated volume.</li>
	<li>Store samples at -80 &#176;C until the processing time.</li>
	<li>Use a microarray kit that detects a range of human proteins to process samples.
	<ol>
		<li>Add 200 &#956;L of each sample to a separate dialysis tube provided with the kit.</li>
		<li>Place the dialysis tubes in a beaker containing at least 500 mL of PBS (pH = 8.0). Stir gently on a stir plate for at least 3 h at 4 &#176;C. Change the PBS in the beaker and repeat this step.</li>
		<li>Transfer each sample to a clean centrifuge tube and centrifuge at 9,000 x g for 5 min to remove any precipitates. Transfer each supernatant to a clean tube.</li>
		<li>Biotinylate each sample by adding 36 &#181;L of 1x labeling reagent from the kit per 1 mg of total protein in the dialyzed sample to 180 &#181;L of dialyzed sample. Incubate at 20 &#176;C for 30 min. Mix gently every 5 min.</li>
		<li>Add 3 &#181;L of stop solution provided with the kit into each sample tube. Transfer each sample to a fresh dialysis tube and repeat steps 4.6.2&#8211;4.6.3. At this stage, the sample can be stored at -20 &#176;C or -80 &#176;C until you are ready to proceed.</li>
		<li>The glass slide provided with the kit is stored at -20 &#176;C. Allow it to come to room temperature. Place the assembled glass slide in a laminar flow hood for 1&#8211;2 h at room temperature.</li>
		<li>Add 400 &#956;L of the blocking buffer provided with the kit into each well of the assembled glass slide. Incubate at room temperature for 30 min.</li>
		<li>Centrifuge the prepared samples for 5 min at 9,000 x g to remove precipitates or particulates. Dilute 5x with blocking buffer.</li>
		<li>Remove the blocking buffer from each well. Add 400 &#956;L of the diluted samples into the appropriate wells. Incubate for 2 h at room temperature while rocking.</li>
		<li>Decant the samples from each well. Wash 3x with 800 &#956;L of the 1x wash buffer I provided with the kit at room temperature for 5 min each while rocking.</li>
		<li>In a clean container, submerge the assembled glass slide in 1x wash buffer I. Wash 2x at room temperature for 5 min each while rocking.</li>
		<li>Add 400 &#956;L of 1x Cy3-conjugated streptavidin to each sub-array. Cover with plastic adhesive strips. Protect from light for the remainder of the protocol.</li>
		<li>Incubate for 2 h at room temperature while rocking.</li>
		<li>Decant the solution and disassemble the glass slide from the sample chambers.</li>
		<li>In the 30 mL centrifuge tube provided with the kit, carefully add the glass slide and enough 1x wash buffer I to cover the glass slide. Wash 3x for 10 min each at room temperature while rocking.</li>
		<li>In the 30 mL centrifuge tube, wash 2x with 1x wash buffer II for 5 min each at room temperature while rocking.</li>
		<li>Wash the glass slide with 30 mL of ddH2O for 5 min. Remove the glass slide from the centrifuge tube and allow to dry for 20 min in a laminar flow hood. The prepared glass slide may be stored at -20 &#176;C until ready to scan.</li>
		<li>Scan the glass slide with a microarray scanner at a fluorescence emission of 555 nm.</li>
	</ol>
	</li>
</ol>

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The authors declare no conflicts of interest.

We developed a microstamping platform to generate uniform arrays of bioparticles to stimulate in vitro neutrophil swarming. The in vitro nature of our platform allows us to circumvent the complications that arise with in vivo swarming experiments, namely the poor ability to analyze mediators released by swarming neutrophils5. Additionally, in vivo models are typically performed in rodents11,12,13,15,22,23, or zebrafish11,12,15,23. Our platform uses human neutrophils, which enables us to more directly interpret our results in the context of human disease, though certain similarities between mouse and human neutrophils have been observed5,11. Additionally, we maintain a spatially-controlled swarming environment that distinguishes our platform from in vivo models by providing a high level of reproducibility that facilitates the analysis of human neutrophil collective migration as well as the collection and analysis of mediators released by swarming neutrophils.
During the development of our microstamping protocol, several challenges arose that required careful troubleshooting. First, the CP used for the microstamping is highly hydrophilic, and the PDMS stamps are hydrophobic. Because the CP does not have a high affinity for PDMS, our procedure was carefully designed to avoid the formation of bubbles and promote wetting. By first priming the stamp with CP while face-up (step 1.8), we minimize the formation of bubbles between the CP and the stamp. The stamp is then inverted onto a layer of CP and incubated for 1 h. This long incubation time ensures that every section of the stamp is wetted. Second, the process of removing excess CP before stamping on a clean glass slide (step 1.11) can be inconsistent. While performing step 1.11, the stamp must be carefully examined. When the pattern begins to become visible, the stamp is ready for step 1.12. Additionally, the required vacuum time to dry the stamps (step 1.12) can vary. This is primarily dependent on the weather. On a warm, humid day, 2 min of vacuum time is required. On a cool, dry day, 1 min of vacuum time is sufficient.
We have shown that neutrophils from healthy donors form stable swarms around bioparticle clusters. With time-lapse fluorescent microscopy, we can quantify swarm size and track neutrophil migration, which enables us to analyze neutrophil chemotaxis quantitatively5. For example, we have previously shown that this platform can be used to calculate the chemotactic index (CI, the cosine of the angle between the neutrophil velocity vector and the position vector between the neutrophil and the nearest bioparticle cluster), speed (distance the neutrophil travels divided by time), radial velocity (the speed multiplied by CI), and the total distance traveled (the difference between initial and final neutrophil position) of individual migrating neutrophils5. Unlike most in vitro studies24,25,26, our platform does not have an artificial chemotactic gradient, so neutrophil intercellular communication is the sole driving force of neutrophil migration. Additionally, the supernatant generated by swarming neutrophils is easily accessible. We can collect and analyze the supernatant for intercellular mediators released by neutrophils without interference from other cell types that are present in vivo. 16 proteins were observed whose expression during swarming could be described as one of three trends: increase (10 proteins), decrease (two proteins), and spike (four proteins). Six of these proteins confirmed previously reported trends during swarming over time5. Some of the identified proteins were previously shown to be swarming-specific, while others were expressed differentially by activated non-swarming neutrophils5. Proteins that increase in concentration over time are likely associated with the pro-inflammation response. Some of the proteins that increase in concentration over time are already known to be involved in the pro-inflammatory response (e.g., galectin-3 and MMP-9)27,28. The relationship between other proteins and inflammation is less well understood. The proteins that spike or decrease during swarming may be involved in the regulation of inflammation. However, further research is necessary to understand the role in inflammation of many of the proteins that are differentially expressed during swarming. Analyzing released mediators along with neutrophil migration can help us better understand the complex picture of inflammation and how neutrophils impact the surrounding tissue during swarming.
In future studies, our platform can be used to thoroughly study the ability of unhealthy neutrophils to generate stable swarms around patterned bioparticle clusters. Various medical conditions have been related to neutrophils, including sepsis3, trauma6, and cancer1,8,17,18, which suggests neutrophils in these patients may have altered function. This platform can be used to examine differences between the intercellular mediators released by healthy and unhealthy neutrophils. Additionally, this platform could be modified to pattern live microbes and used to analyze neutrophil response to live microbes in vitro.
In conclusion, we have developed a novel platform for analyzing neutrophil swarming in vitro. The highly controlled nature of our platform allows us to mitigate issues that arise during in vivo neutrophil swarming experiments. Neutrophil swarming on a bioparticle microarray is easily quantifiable via time-lapse fluorescent microscopy. Additionally, we can collect the mediators released by the neutrophils without the interference of other tissues that are present in vivo. This platform can be used in future research to quantify differences between the migration behaviors of neutrophils from healthy and unhealthy donors.

Neutrophils, the most abundant white blood cell in the bloodstream1, are gaining attention as potential diagnostic and therapeutic targets2,3 because they may be involved in a variety of medical conditions including gout4, sepsis3, trauma5,6, cancer1,7,8, and various autoimmune diseases5,9. Neutrophil swarming is a multistage, tightly regulated process with a complexity that makes it a particularly interesting focus of study5,10,11. During swarming, neutrophils isolate a site of inflammation from the surrounding healthy tissue5,10,11. Proper regulation of neutrophil swarming is essential to promote wound healing and ultimately inflammation resolution5,12. Neutrophil swarming has primarily been studied in vivo in rodent12,13,14,15 and zebrafish10,11,12,15 models. However, the nature of these in vivo animal models gives rise to limitations5. For example, the mediators released by neutrophils during swarming are not easily accessible for analysis5. Additionally, there are many potential sources for a given mediator in vivo, so an in vivo experiment must introduce a genetic deficiency to inhibit cellular production and/or interaction in order to investigate the role of that mediator in a given process13. An in vitro experiment circumvents this complication by enabling neutrophil observation without the context of additional cells. Additionally, research describing human neutrophil coordinated migration is limited16. On an in vitro swarming platform, human neutrophils can be directly analyzed. An in vitro swarming platform could expand upon the knowledge gained from in vivo studies by providing opportunities to fill the gaps left by the limitations of in vivo studies.
To address the need for an in vitro platform that mimics in vivo neutrophil swarming, we developed a microstamping platform that enables us to pattern bioparticle microarrays that stimulate neutrophil swarming in a spatially controlled manner. We generate bioparticle microarrays on glass slides in a two-step process. First, we use microstamping to generate a microarray of cationic polyelectrolyte (CP) spots. Second, we add a solution of bioparticles that adhere to the CP spots via electrostatic interaction. By first patterning the CP layer, we can selectively pattern negatively charged bioparticles to generate the desired neutrophil swarming pattern. The positively charged layer holds the negatively charged bioparticles through the vigorous washing step that removes the bioparticles from the areas on the glass slide that do not have the CP. Additionally, the CP used here, a copolymer of acrylamide and quaternized cationic monomer, is biocompatible, so it does not induce a response from the neutrophils. It has a very high surface charge that immobilizes the micron-sized bioparticles to the glass slide, thus inhibiting neutrophils from removing the particles from the patterned position on the glass slide. This results in bioparticle clusters arranged in a microarray. When we added neutrophils to the microarray, they formed stable swarms around the bioparticle clusters. Through tracking neutrophil migration, we found that swarming neutrophils actively migrate toward the bioparticle clusters. Furthermore, we used this platform to analyze certain mediators that neutrophils release during swarming. We found 16 mediators that are differentially expressed during swarming. Their concentrations follow three general trends over time: increase, decrease, or spike. Our in vitro neutrophil swarming platform facilitates the analysis of spatially controlled human neutrophil swarming, as well as the collection and analysis of mediators released by neutrophil swarming. In a previous publication, we demonstrated that patients with certain medical conditions (trauma, autoimmune disease, and sepsis), had neutrophils that functioned differently than those from healthy donors5. In future research studies, our platform could be used to analyze neutrophil function among a variety of patient populations. This platform can quantitatively analyze the complex coordination involved in neutrophil swarming. Additional studies can be done to provide insight on the neutrophil function of a specific patient population or neutrophil response to a pathogen of interest.

<table><tbody><tr><td>&quot;The Big Easy&quot; EasySep Magnet</td><td>STEMCELL Technologies</td><td>18001</td><td>Magnet to use with neutrophil isolation kit</td></tr><tr><td>Cell Incubator</td><td>Okolab</td><td>777057437 / 77057343</td><td>Okolab cage incubator for temperature and CO2 control</td></tr><tr><td>EasySep Human Neutrophil Isolation Kit</td><td>STEMCELL Technologies</td><td>17957</td><td>Kit for immunomagnetic negative selection of human neutrophils</td></tr><tr><td>Eclipse Ti2</td><td>Nikon Instruments</td><td>MEA54010 / MEF55037</td><td>Inverted research microscope</td></tr><tr><td>Escherichia coli (K-12 strain) BioParticles Texas Red conjugate</td><td>Invitrogen</td><td>E2863</td><td>Bioparticle powder, dissolve in water prior to addition to Zetag&amp;reg; array</td></tr><tr><td>Harris Uni-Core 8-mm biopsy punch</td><td>Sigma Aldrich</td><td>Z708925</td><td>To cut PDMS stamps</td></tr><tr><td>HetaSep</td><td>STEMCELL Technologies</td><td>7906</td><td>Erythrocyte aggregation agent for separating buffy coat from red blood cells in fresh human blood</td></tr><tr><td>Hoechst 33342</td><td>Life Technologies</td><td>H3570</td><td>Nucleus fluorescent stain</td></tr><tr><td>Human L1000 Array</td><td>Raybiotech Inc.</td><td>AAH-BLG-1000-4</td><td>High density array to detect 1000 human proteins</td></tr><tr><td>Human Serum Albumin (HSA)</td><td>Sigma Aldrich</td><td>A5843</td><td>Low endotoxin HSA, to prepare 2 % solutions in IMDM for isolated neutrophils</td></tr><tr><td>Iscove&#039;s Modified Dulbeccos&#039; Medium (IMDM)</td><td>Thermo Fisher Scientific</td><td>12440053</td><td>To resuspend isolated neutrophils</td></tr><tr><td>K2-EDTA tubes</td><td>Thermo Fisher Scientific</td><td>02-657-32</td><td>Tubes for blood collection</td></tr><tr><td>Low Reflective Chrome Photomask</td><td>Front Range Photomask</td><td>N/A</td><td>Dimensions 5&quot; x 5&quot; x 0.09&quot; (L x W x D)</td></tr><tr><td>Microarray Scanner</td><td>Perkin Elmer</td><td>ASCNGX00</td><td>Fluorescence reader of protein patterned microdomains</td></tr><tr><td>Microscopy Image Analsysis Software - Imaris</td><td>Bitplane</td><td>9.3.0</td><td>Software for automatic cell tracking analysis</td></tr><tr><td>NiS Elements Advanced Research Software Package</td><td>Nikon Instruments</td><td>MQS31100</td><td>Software for automatic live cell imaging and swarm size calculation</td></tr><tr><td>Poly-L-lysine fluorescein isothiocyanate (PLL-FITC)</td><td>Sigma Aldrich</td><td>P3069-10MG</td><td>30,000 - 70,000 MW PLL labeled with FITC, used to fluorescently label CP solution</td></tr><tr><td>SecureSeal 8-well Imaging Spacer</td><td>Grace Bio-Labs</td><td>654008</td><td>8-well, 9-mm diameter, adhesive imaging spacer</td></tr><tr><td>Silicon Wafer</td><td>University Wafer</td><td>590</td><td>Silicon 100 mm N/P (100) 0- 100 ohm-cm 500 &amp;mu;m SSP test</td></tr><tr><td>Spin Coater</td><td>Laurell</td><td>WS-650MZ-23NPPB</td><td>Used to spincoat a 40-&amp;micro;m layer of photoresist onto silicon wafer</td></tr><tr><td>SU-8 2025</td><td>MicroChem</td><td>2025</td><td>Negative photoresist to make silicon master wafer</td></tr><tr><td>SU-8 Developer</td><td>MicroChem</td><td>Y020100</td><td>Photoresist developer. Remove non-crosslinked SU-8 2025 from silicon wafer</td></tr><tr><td>Sylgard 184 (polydimethylsiloxane, PDMS)</td><td>Dow</td><td>1673921</td><td>2-part silicone elastomer kit for making microstamps and PDMS wells</td></tr><tr><td>UV Exposure Masking System</td><td>Klo&amp;eacute;</td><td>UV-KUB 2</td><td>Used to crosslink photoresist on silicon wafer through chrome mask with UV light</td></tr><tr><td>Water</td><td>Thermo Fisher Scientific</td><td>A1287303</td><td>High quality water to dilute bioparticles</td></tr><tr><td>Zetag 8185</td><td>BASF</td><td>8185</td><td>Cationic polyelectrolyte (CP), powder, Copolymer of acrylamide and quaternized cationic monomer, forms &quot;inking solution&quot; for microstamping when dissolved in water</td></tr><tr><td>Zymosan A S. cerevisiae BioParticles Texas Red conjugate</td><td>Invitrogen</td><td>Z2843</td><td>Bioparticle powder, dissolve in water prior to addition to Zetag array</td></tr></tbody></table>

bioparticle microarrays for chemotactic and molecular analysis of human neutrophil swarming in vitro

Neutrophil swarming is a cooperative process by which neutrophils seal off a site of infection and promote tissue reorganization. Swarming has classically been studied in vivo in animal models showing characteristic patterns of cell migration. However, in vivo models have several limitations, including intercellular mediators that are difficult to access and analyze, as well as the inability to directly analyze human neutrophils. Because of these limitations, there is a need for an in vitro platform that studies swarming with human neutrophils and provides easy access to the molecular signals generated during swarming. Here, a multistep microstamping process is used to generate a bioparticle microarray that stimulates swarming by mimicking an in vivo infection. The bioparticle microarray induces neutrophils to swarm in a controlled and stable manner. On the microarray, neutrophils increase in speed and form stable swarms around bioparticle clusters. Additionally, supernatant generated by the neutrophils was analyzed and 16 proteins were discovered to have been differentially expressed over the course of swarming. This in vitro swarming platform facilitates direct analysis of neutrophil migration and protein release in a reproducible, spatially controlled manner.

When neutrophils are added to the bioparticle microarray, neutrophils that contact the bioparticle clusters become activated and initiate the swarming response. The bioparticle microarray was validated using time-lapse fluorescent microscopy to track neutrophil migration toward the bioparticle clusters (Video S1). The migration of individual neutrophil nuclei is tracked as they migrate toward the bioparticle cluster. When neutrophils reach the bioparticle cluster, their nuclei overlap with other nuclei in the cluster. Thus, it is not possible to accurately track a neutrophil within the cluster using this method. Zymosan and E. coli bioparticle clusters both result in the generation of neutrophil swarms. For our results, Figure 2B uses data from neutrophil swarms generated by E. coli particles. Figure 3 and the other panels of Figure 2 use neutrophil swarms generated by zymosan particles. The results obtained demonstrate that bioparticle clusters stimulates neutrophil activation when a neutrophil contacted the cluster, which ultimately led to the formation of stable neutrophil swarms around each cluster after 30&#8211;60 min (Figure 2A, top). In contrast, neutrophils did not show collective migration in the absence of bioparticle clusters (Figure 2A, bottom, and Video S2). Using the fluorescent intensity of stained neutrophil nuclei at 405 nm, the average neutrophil swarm size around 30 &#181;m diameter E. coli bioparticle clusters was found to be 1,490 &#177; 680 &#181;m2 (mean &#177; SD, Figure 2B, top). The fluorescence intensity of a given region of interest where no bioparticle clusters are present was approximately constant over time, which confirmed the absence of collective migration in this setting (Figure 2B, bottom). Tracks of neutrophil migration show that neutrophils converged on a bioparticle cluster when one was present (Figure 2C, top). Conversely, no convergence was observed in the control system (Figure 2C, bottom). The speed (distance travelled/time) of swarming and nonactivated neutrophils was measured and a statistically significant difference in the speed distributions was found (ANOVA, p &#60; 0.0001), as shown in Figure 2D. The average speed for swarming neutrophils was 20.6 &#177; 13.0 &#181;m/min (mean &#177; SD), and the average speed for control neutrophils was 2.0 &#177; 2.2 &#181;m/min.
Additionally, the concentration of 16 proteins that neutrophils released during swarming were analyzed (Figure 3). The normalized concentration of each protein at different time points in the swarming process (t = 0, 0.5, 1, and 3 h) were calculated, where the normalized concentration is (C &#8211; Cmin) / (Cmax &#8211; Cmin). 10 proteins increased in concentration throughout swarming. These proteins were adipsin, galectin-3, GROa, IL-6R, MIP-1a, MMP-8, MMP-9, Nidogen-1, TIMP-1, and TLR2. Two proteins (pentraxin 3 and RANK) decreased in concentration throughout swarming. The remaining four proteins (clusterin, PF4, RANTES, and Trappin-2) increased during the first hour of swarming but decreased thereafter. In other words, the concentrations of those proteins &#34;spiked&#34; during swarming. Of the 16 proteins identified, 12 proteins were identified in our previous publication5. Adipsin, galectin-3, nidogen-1, pentraxin 3, TIMP-1, and TLR2 were shown to be swarming-specific, while clusterin, IL-6R, MMP-8, and MMP-9, RANK, and trappin-2 were not5.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60544/60544fig01.jpg" /> 
Figure 1: Production of bioparticle microarray. A silicon wafer coated with negative photoresist is exposed to UV light through a chrome photomask (A). After the silicon wafer is baked and developed, a photoresist pattern remains on the surface of the silicon wafer. This is the master wafer (B). In a Petri dish, a PDMS mixture is added to the top of the master wafer. The PDMS is cured overnight at 65 &#176;C to form the PDMS mold (C). The PDMS mold is cut from the master wafer and a biopsy punch is used to punch out individual PDMS stamps (D). A PDMS stamp (E) is coated with CP solution (F). The stamp is inverted onto a thin layer of CP solution (G). After incubating in the CP solution for 1 h, the stamp is blotted onto a glass slide to remove excess CP. Eight stamps are then pressed onto a clean glass slide with a 5.6 &#177; 0.1 g weight, aligned with an imaging spacer (H). When the stamp is removed, a CP pattern remains (I). A PDMS slab with precut wells is adhered to the glass slide (J). Then, a bioparticle solution is added over the CP pattern (K). The negatively charged bioparticles bind to the positively charged polyelectrolyte via electrostatic interaction. The excess bioparticle solution is washed away, leaving bioparticles patterned on top of the CP pattern (L). (M) Fluorescent image of the CP layer labeled with FITC (Scale bars = 50 &#181;m, large image; Scale bar = 25 &#181;m, inset). (N) Fluorescent image of patterned zymosan bioparticles conjugated with Texas Red (Scale bar = 100 &#181;m, large image; Scale bar = 50 &#181;m, inset). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60544/60544fig01large.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/60544/60544fig02.jpg" /> 
Figure 2: Neutrophil swarm growth around bioparticle clusters. In the presence of bioparticle clusters, neutrophils undergo collective migration toward the bioparticle clusters (bottom, control neutrophils). When no bioparticles are present, the neutrophils do not perform collective migration. (A) In the presence of zymosan bioparticle clusters, neutrophils form swarms in 30 min. Without bioparticle clusters, no swarms form (Scale bars = 50 &#181;m). (B) Neutrophil swarms grow to an average size of 1,490 &#177; 680 &#181;m2 (mean &#177; SD) around 30 &#181;m diameter E. coli bioparticle clusters. Control neutrophils exhibit a constant density that corresponds to no swarm growth (ANOVA, p &#60; 0.0001, n = 32 neutrophil swarms, N = 1 donor, error bars = standard deviation). (C) Tracks of swarming neutrophils converge on the zymosan bioparticle clusters, while control neutrophils show no converging point (Scale bars = 50 &#181;m). (D) Neutrophils swarming toward a zymosan target have a mean speed of 20.6 &#177; 13.0 &#181;m/min (mean &#177; SD), while control neutrophils have a mean speed of 2.0 &#177; 2.2 &#181;m/min. Each count on the heat map represents a neutrophil with the given instantaneous speed at the given time point. These heat maps are representative of one experiment (n = 1 swarm; N = 1 donor; ANOVA, 6,114 = swarming neutrophils, 32,116 = control neutrophils, p &#60; 0.0001). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60544/60544fig02large.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/60544/60544fig03.jpg" /> 
Figure 3: Free mediators released by swarming neutrophils. Neutrophil swarming affects the production of various proteins over time. Protein concentration was measured at 0, 0.5, 1, and 3 h. The data points were fitted with smoothing splines (&#955; = 0.05). These proteins tend to follow one of three characteristic trends: decreasing over time, increasing over time, or spiking around 1 h and then decreasing (Error bars = standard deviation; n = 3 replicates; N = 1 donor). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60544/60544fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video S1: Neutrophil swarming toward zymosan bioparticle clusters. Neutrophil nuclei are shown in blue. Zymosan targets are marked with red circles (Scale bar = 50 &#181;m; original acquisition time = 60 min). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60544/Supplementary_Video_1.mp4" target="_blank">Please click here to download this video.</a>
Video S2: Nonactivated neutrophil random migration. Neutrophil nuclei are shown in blue (Scale bar = 50 &#181;m; original acquisition time = 60 min). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60544/Supplementary_Video_2.mp4" target="_blank">Please click here to download this video.</a>

Watch this Scientific Journal Video about Bioparticle Microarrays for Chemotactic and Molecular Analysis of Human Neutrophil Swarming in vitro at JoVE.com

Bioparticle Microarrays for Chemotactic and Molecular Analysis of Human Neutrophil Swarming in vitro

This protocol generates bioparticle microarrays that provide spatially controlled neutrophil swarming. It provides easy access to the mediators that neutrophils release during migration and allows for quantitative imaging analysis.

Neutrophil swarming is a cooperative process by which neutrophils seal off a site of infection and promote tissue reorganization. Swarming has classically ...

bioparticle-microarrays-for-chemotactic-molecular-analysis-human

Nanyang Technological University

Research

JoVE Journal

Bioengineering

5.0K Views.  The Ohio State University. Dit protocol is belangrijk omdat het een in vitro analyse van neutrofiele zwermen mogelijk maakt, die vele beperkingen van ons huidige onvermogen om te experimenteren overwint. Het belangrijkste voordeel van deze techniek is dat het gemakkelijke toegang biedt tot de uitgescheiden cytokines en lipide bemiddelaars die neutrofielen vrijlaten tijdens het zwermen en kwantitatieve beeldvormingsanalyse mogelijk maakt. Terwijl we deze techniek toepasten op neutrofielen, kon het worden uitgebreid om d...

Video: Bioparticle Microarrays for Chemotactic and Molecular Analysis of Human Neutrophil Swarming in vitro

In vitro Assembly of Semi-artificial Molecular Machine and its Use for Detection of DNA Damage

Naturally occurring bio-molecular machines work in every living cell and display a variety of designs 1-6. Yet the development of artificial molecular machines centers on devices capable of directional motion, i.e. molecular motors, and on their scaled-down mechanical parts (wheels, axels, pendants etc) 7-9. This imitates the macro-machines, even though the physical properties essential for these devices, such as inertia and momentum conservation, are not usable in the nanoworld environments 10. Alternative designs, which do not follow the mechanical macromachines schemes and use mechanisms developed in the evolution of biological molecules, can take advantage of the specific conditions of the nanoworld. Besides, adapting actual biological molecules for the purposes of nano-design reduces potential dangers the nanotechnology products may pose. Here we demonstrate the assembly and application of one such bio-enabled construct, a semi-artificial molecular device which combines a naturally-occurring molecular machine with artificial components. From the enzymology point of view, our construct is a designer fluorescent enzyme-substrate complex put together to perform a specific useful function. This assembly is by definition a molecular machine, as it contains one 12. Yet, its integration with the engineered part - fluorescent dual hairpin - re-directs it to a new task of labeling DNA damage12.
Our construct assembles out of a 32-mer DNA and an enzyme vaccinia topoisomerase I (VACC TOPO). The machine then uses its own material to fabricate two fluorescently labeled detector units (Figure 1). One of the units (green fluorescence) carries VACC TOPO covalently attached to its 3'end and another unit (red fluorescence) is a free hairpin with a terminal 3'OH. The units are short-lived and quickly reassemble back into the original construct, which subsequently recleaves. In the absence of DNA breaks these two units continuously separate and religate in a cyclic manner. In tissue sections with DNA damage, the topoisomerase-carrying detector unit selectively attaches to blunt-ended DNA breaks with 5'OH (DNase II-type breaks)11,12, fluorescently labeling them. The second, enzyme-free hairpin formed after oligonucleotide cleavage, will ligate to a 5'PO4 blunt-ended break (DNase I-type breaks)11,12, if T4 DNA ligase is present in the solution 13,14 . When T4 DNA ligase is added to a tissue section or a solution containing DNA with 5'PO4 blunt-ended breaks, the ligase reacts with 5'PO4 DNA ends, forming semi-stable enzyme-DNA complexes. The blunt ended hairpins will interact with these complexes releasing ligase and covalently linking hairpins to DNA, thus labeling 5'PO4 blunt-ended DNA breaks.
This development exemplifies a new practical approach to the design of molecular machines and provides a useful sensor for detection of apoptosis and DNA damage in fixed cells and tissues.

In vitro Assembly of Semi-artificial Molecular Machine and its Use for Detection of DNA Damage

We demonstrate the assembly and application of a molecular-scale device powered by a topoisomerase protein. The construct is a bio-molecular sensor which labels two major types of DNA breaks in tissue sections by attaching two different fluorophores to their ends.

Naturally occurring bio-molecular machines work in every living cell and display a variety of designs 1-6. Yet the development of artificial molecular ...

Polymer Microarrays for High Throughput Discovery of Biomaterials

The discovery of novel biomaterials that are optimized for a specific biological application is readily achieved using polymer microarrays, which allows a combinatorial library of materials to be screened in a parallel, high throughput format1. Herein is described the formation and characterization of a polymer microarray using an on-chip photopolymerization technique 2. This involves mixing monomers at varied ratios to produce a library of monomer solutions, transferring the solution to a glass slide format using a robotic printing device and curing with UV irradiation. This format is readily amenable to many biological assays, including stem cell attachment and proliferation, cell sorting and low bacterial adhesion, allowing the ready identification of 'hit' materials that fulfill a specific biological criterion3-5. Furthermore, the use of high throughput surface characterization (HTSC) allows the biological performance to be correlated with physio-chemical properties, hence elucidating the biological-material interaction6. HTSC makes use of water contact angle (WCA) measurements, atomic force microscopy (AFM), X-ray photoelectron spectroscopy (XPS) and time-of-flight secondary ion mass spectrometry (ToF-SIMS). In particular, ToF-SIMS provides a chemically rich analysis of the sample that can be used to correlate the cell response with a molecular moiety. In some cases, the biological performance can be predicted from the ToF-SIMS spectra, demonstrating the chemical dependence of a biological-material interaction, and informing the development of hit materials5,3.

A description of the formation of a polymer microarray using an on-chip photopolymerization technique. The high throughput surface characterization using atomic force microscopy, water contact angle measurements, X-ray photoelectron spectroscopy and time of flight secondary ion mass spectrometry and a cell attachment assay is also described.

The discovery of novel biomaterials that are optimized for a specific biological application is readily achieved using polymer microarrays, which allows a ...

Silk Film Culture System for in vitro Analysis and Biomaterial Design

Silk films are promising protein-based biomaterials that can be fabricated with high fidelity and economically within a research laboratory environment 1,2 . These materials are desirable because they possess highly controllable dimensional and material characteristics, are biocompatible and promote cell adhesion, can be modified through topographic patterning or by chemically altering the surface, and can be used as a depot for biologically active molecules for drug delivery related applications 3-8 . In addition, silk films are relatively straightforward to custom design, can be designed to dissolve within minutes or degrade over years in vitro or in vivo, and are produce with the added benefit of being transparent in nature and therefore highly suitable for imaging applications 9-13. The culture system methodology presented here represents a scalable approach for rapid assessments of cell-silk film surface interactions. Of particular interest is the use of surface patterned silk films to study differences in cell proliferation and responses of cells for alignment 12,14 . The seeded cultures were cultured on both micro-patterned and flat silk film substrates, and then assessed through time-lapse phase-contrast imaging, scanning electron microscopy, and biochemical assessment of metabolic activity and nucleic acid content. In summary, the silk film in vitro culture system offers a customizable experimental setup suitable to the study of cell-surface interactions on a biomaterial substrate, which can then be optimized and then translated to in vivo models. Observations using the culture system presented here are currently being used to aid in applications ranging from basic cell interactions to medical device design, and thus are relevant to a broad range of biomedical fields.

Silk Film Culture System for in vitro Analysis and Biomaterial Design

Silk films are a novel class of biomaterials readily customizable for an array of biomedical applications. The presented silk film culture system is highly adaptable to a variety of in vitro analyses. This system represents a biomaterial design platform offering in vitro optimization before direct translation to in vivo models.

Silk films are promising protein-based biomaterials that can be fabricated with high fidelity and economically within a research laboratory environment ...

Tracking Neutrophil Intraluminal Crawling, Transendothelial Migration and Chemotaxis in Tissue by Intravital Video Microscopy

The recruitment of circulating leukocytes from blood stream to the inflamed tissue is a crucial and complex process of inflammation1,2. In the postcapillary venules of inflamed tissue, leukocytes initially tether and roll on the luminal surface of venular wall. Rolling leukocytes arrest on endothelium and undergo firm adhesion in response to chemokine or other chemoattractants on the venular surface. Many adherent leukocytes relocate from the initial site of adhesion to the junctional extravasation site in endothelium, a process termed intraluminal crawling3. Following crawling, leukocytes move across endothelium (transmigration) and migrate in extravascular tissue toward the source of chemoattractant (chemotaxis)4. Intravital microscopy is a powerful tool for visualizing leukocyte-endothelial cell interactions in vivo and revealing cellular and molecular mechanisms of leukocyte recruitment2,5. In this report, we provide a comprehensive description of using brightfield intravital microscopy to visualize and determine the detailed processes of neutrophil recruitment in mouse cremaster muscle in response to the gradient of a neutrophil chemoattractant. To induce neutrophil recruitment, a small piece of agarose gel (~1-mm3 size) containing neutrophil chemoattractant MIP-2 (CXCL2, a CXC chemokine) or WKYMVm (Trp-Lys-Tyr-Val-D-Met, a synthetic analog of bacterial peptide) is placed on the muscle tissue adjacent to the observed postcapillary venule. With time-lapsed video photography and computer software ImageJ, neutrophil intraluminal crawling on endothelium, neutrophil transendothelial migration and the migration and chemotaxis in tissue are visualized and tracked. This protocol allows reliable and quantitative analysis of many neutrophil recruitment parameters such as intraluminal crawling velocity, transmigration time, detachment time, migration velocity, chemotaxis velocity and chemotaxis index in tissue. We demonstrate that using this protocol, these neutrophil recruitment parameters can be stably determined and the single cell locomotion conveniently tracked in vivo.

We describe a protocol of brightfield intravital microscopy for measuring dynamic neutrophil-endothelial cell interactions during neutrophil recruitment in response to the source of a neutrophil chemoattractant in vivo. Neutrophil intraluminal crawling, transendothelial migration and chemotaxis in mouse cremaster muscle tissue are visualized with time-lapsed video photography and tracked with ImageJ.

The recruitment of circulating leukocytes from blood stream to the inflamed tissue is a crucial and complex process of inflammation1,2. In the ...

Immunology and Infection

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 ...

An All-on-chip Method for Rapid Neutrophil Chemotaxis Analysis Directly from a Drop of Blood

Neutrophil migration and chemotaxis are critical for our body's immune system. Microfluidic devices are increasingly used for investigating neutrophil migration and chemotaxis owing to their advantages in real-time visualization, precise control of chemical concentration gradient generation, and reduced reagent and sample consumption. Recently, a growing effort has been made by the microfluidic researchers toward developing integrated and easily operated microfluidic chemotaxis analysis systems, directly from whole blood. In this direction, the first all-on-chip method was developed for integrating the magnetic negative purification of neutrophils and the chemotaxis assay from small blood volume samples. This new method permits a rapid sample-to-result neutrophil chemotaxis test in 25 min. In this paper, we provide detailed construction, operation and data analysis method for this all-on-chip chemotaxis assay with a discussion on troubleshooting strategies, limitations and future directions. Representative results of the neutrophil chemotaxis assay testing a defined chemoattractant, N-Formyl-Met-Leu-Phe (fMLP), and sputum from a chronic obstructive pulmonary disease (COPD) patient, using this all-on-chip method are shown. This method is applicable to many cell migration-related investigations and clinical applications.

This article provides the detailed method of performing a rapid neutrophil chemotaxis assay by integrating the on-chip neutrophil isolation from whole blood and the chemotaxis test on a single microfluidic chip.

Neutrophil migration and chemotaxis are critical for our body's immune system. Microfluidic devices are increasingly used for investigating neutrophil ...

Quantifying Human Monocyte Chemotaxis In Vitro and Murine Lymphocyte Trafficking In Vivo

Chemotaxis is migration along a specific chemical gradient1. Chemokines are chemotactic cytokines that promote cellular trafficking with anatomic and temporal specificity2. Chemotaxis is a critical function of lymphocytes and other immune cells that can be quantitatively assessed in vitro. This manuscript describes methods that permit the evaluation of chemotaxis, both in vitro and in vivo, for diverse cell types including cell lines and native cells. The in vitro, plate-based format permits the comparison of several conditions simultaneously in real-time, and can be completed within 1-4 h. In vitro assay conditions can be manipulated to introduce agonists and antagonists, as well as differentiate chemotaxis from chemokinesis, which is random movement. For in vivo trafficking assessments, immune cells can be labeled with multiple fluorescent dyes and used for adoptive transfer. The differential labeling of cells allows for mixed cell populations to be introduced into the same animal, thereby decreasing variance and reducing the number of animals required for an adequately powered experiment. Migration into lymphoid tissue occurs in as little as 1 h, and multiple tissue compartments can be sampled. Flow cytometry following tissue harvest allows for a rapid and quantitative analysis of the migratory patterns of multiple cell types.

Quantifying Human Monocyte Chemotaxis In Vitro and Murine Lymphocyte Trafficking In Vivo

Protocols for quantitative assessment of lymphocyte chemotaxis and migration are important tools for immunology research. Here, an in vitro protocol is described that permits real-time, multiplexed evaluation of cell migration, as well as a complementary in vivo technique enabling tracking of native cells to spleen.

Chemotaxis is migration along a specific chemical gradient1. Chemokines are chemotactic cytokines that promote cellular trafficking with anatomic and ...

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

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

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

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

Imaging G Protein-coupled Receptor-mediated Chemotaxis and its Signaling Events in Neutrophil-like HL60 Cells

Eukaryotic cells sense and move towards a chemoattractant gradient, a cellular process referred as chemotaxis. Chemotaxis plays critical roles in many physiological processes, such as embryogenesis, neuron patterning, metastasis of cancer cells, recruitment of neutrophils to sites of inflammation, and the development of the model organism Dictyostelium discoideum. Eukaryotic cells sense chemo-attractants using G protein-coupled receptors. Visual chemotaxis assays are essential for a better understanding of how eukaryotic cells control chemoattractant-mediated directional cell migration. Here, we describe detailed methods for: 1) real-time, high-resolution monitoring of multiple chemotaxis assays, and 2) simultaneously visualizing the chemoattractant gradient and the spatiotemporal dynamics of signaling events in neutrophil-like HL60 cells.

Visual chemotaxis assays are essential for a better understanding of how eukaryotic cells control chemoattractant-mediated directional cell migration. Here, we describe detailed methods for: 1) real-time, high-resolution monitoring of multiple chemotaxis assays, and 2) simultaneously visualizing the chemoattractant gradient and the spatiotemporal dynamics of signaling events in neutrophil-like HL60 cells.

Eukaryotic cells sense and move towards a chemoattractant gradient, a cellular process referred as chemotaxis. Chemotaxis plays critical roles in many ...

Biology

A Microfluidic Chip-Based Method for Rapid Neutrophil Chemotaxis Analysis Using Whole Blood

Source: Yang, K., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/55615">An All-on-chip Method for Rapid Neutrophil Chemotaxis Analysis Directly from a Drop of Blood.</a> J. Vis. Exp. (124), (2017)
This video demonstrates neutrophil chemotaxis assay using an all-on-chip method. The method allows direct isolation of neutrophils from the whole blood by immunomagnetic negative selection followed by chemotaxis assay on a microfluidic chip.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Microfluidic device preparation.

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Bioparticle Microarrays for Chemotactic and Molecular Analysis of Human Neutrophil Swarming in vitro

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