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,</s...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#34;The Big Easy&#34; 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&#174; 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&#39;s Modified Dulbeccos&#39; 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&#34; x 5&#34; x 0.09&#34; (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 &#956;m SSP test</td>
		</tr>
		<tr>
			<td>Spin Coater</td>
			<td>Laurell</td>
			<td>WS-650MZ-23NPPB</td>
			<td>Used to spincoat a 40-&#181;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&#233;</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 &#34;inking solution&#34; 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 i...

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

Research

JoVE Journal

Bioengineering

5.0K Views.  The Ohio State University. 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.

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 ...

Self-reporting Scaffolds for 3-Dimensional Cell Culture

Culturing cells in 3D on appropriate scaffolds is thought to better mimic the in vivo microenvironment and increase cell-cell interactions. The resulting 3D cellular construct can often be more relevant to studying the molecular events and cell-cell interactions than similar experiments studied in 2D. To create effective 3D cultures with high cell viability throughout the scaffold the culture conditions such as oxygen and pH need to be carefully controlled as gradients in analyte concentration can exist throughout the 3D construct. Here we describe the methods of preparing biocompatible pH responsive sol-gel nanosensors and their incorporation into poly(lactic-co-glycolic acid) (PLGA) electrospun scaffolds along with their subsequent preparation for the culture of mammalian cells. The pH responsive scaffolds can be used as tools to determine microenvironmental pH within a 3D cellular construct. Furthermore, we detail the delivery of pH responsive nanosensors to the intracellular environment of mammalian cells whose growth was supported by electrospun PLGA scaffolds. The cytoplasmic location of the pH responsive nanosensors can be utilized to monitor intracellular pH (pHi) during ongoing experimentation.

Biocompatible pH responsive sol-gel nanosensors can be incorporated into poly(lactic-co-glycolic acid) (PLGA) electrospun scaffolds. The produced self-reporting scaffolds can be used for in&#160;situ monitoring of microenvironmental conditions whilst culturing cells upon the scaffold. This is beneficial as the 3D cellular construct can be monitored in real-time without disturbing the experiment.

Culturing cells in 3D on appropriate scaffolds is thought to better mimic the in vivo microenvironment and increase cell-cell interactions. The resulting ...

Eye Irritation Test (EIT) for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial (RhCE) Tissue Model

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and accurate assessment of ocular toxicity in man are needed. To address this need, we have developed an eye irritation test (EIT) which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model that is based on normal human cells. The EIT is able to separate ocular&#160;irritants and corrosives (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category). The test utilizes two separate protocols, one designed for liquid chemicals and a second, similar protocol for solid test articles. The EIT prediction model uses a single exposure period (30 min for liquids, 6 hr for solids) and a single tissue viability cut-off (60.0% as determined by the MTT assay). Based on the results for 83 chemicals (44 liquids and 39 solids) EIT achieved 95.5/68.2/ and 81.8% sensitivity/specificity and accuracy (SS&#38;A) for liquids, 100.0/68.4/ and 84.6% SS&#38;A for solids, and 97.6/68.3/ and 83.1% for overall SS&#38;A. The EIT will contribute significantly to classifying the ocular irritation potential of a wide range of liquid and solid chemicals without the use of animals to meet regulatory testing requirements. The EpiOcular EIT method was implemented in 2015 into the OECD Test Guidelines as TG 492.

Eye Irritation Test EIT for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial RhCE Tissue Model

We have developed an eye irritation test which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model. The test is able to discriminate between ocular irritant and corrosive materials (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category).

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and ...

Adapting the Electrospinning Process to Provide Three Unique Environments for a Tri-layered In Vitro Model of the Airway Wall

Electrospinning is a highly adaptable method producing porous 3D fibrous scaffolds that can be exploited in in vitro cell culture. Alterations to intrinsic parameters within the process allow a high degree of control over scaffold characteristics including fiber diameter, alignment and porosity. By developing scaffolds with similar dimensions and topographies to organ- or tissue-specific extracellular matrices (ECM), micro-environments representative to those that cells are exposed to in situ can be created. 
The airway bronchiole wall, comprised of three main micro-environments, was selected as a model tissue. Using decellularized airway ECM as a guide, we electrospun the non-degradable polymer, polyethylene terephthalate (PET), by three different protocols to produce three individual electrospun scaffolds optimized for epithelial, fibroblast or smooth muscle cell-culture. Using a commercially available bioreactor system, we stably co-cultured the three cell-types to provide an in vitro model of the airway wall over an extended time period.
This model highlights the potential for such methods being employed in in vitro diagnostic studies investigating important inter-cellular cross-talk mechanisms or assessing novel pharmaceutical targets, by providing a relevant platform to allow the culture of fully differentiated adult cells within 3D, tissue-specific environments.

Adapting the Electrospinning Process to Provide Three Unique Environments for a Tri-layered In Vitro Model of the Airway Wall

Advancements in biomaterial technologies enable the development of three-dimensional multi-cell-type constructs. We have developed electrospinning protocols to produce three individual scaffolds to culture the main structural cells of the airway to provide a 3D in vitro model of the airway bronchiole wall.

Electrospinning is a highly adaptable method producing porous 3D fibrous scaffolds that can be exploited in in vitro cell culture. Alterations to ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Traction Microscopy Integrated with Microfluidics for Chemotactic Collective Migration

Cells change migration patterns in response to chemical stimuli, including the gradients of the stimuli. Cellular migration in the direction of a chemical gradient, known as chemotaxis, plays an important role in development, the immune response, wound healing, and cancer metastasis. While chemotaxis modulates the migration of single cells as well as collections of cells in vivo, in vitro research focuses on single-cell chemotaxis, partly due to the lack of the proper experimental tools. To fill that gap, described here is a unique experimental system that combines microfluidics and micropatterning to demonstrate the effects of chemical gradients on collective cell migration. Furthermore, traction microscopy and monolayer stress microscopy are incorporated into the system to characterize changes in cellular force on the substrate as well as between neighboring cells. As proof-of-concept, the migration of micropatterned circular islands of Madin-Darby canine kidney (MDCK) cells is tested under a gradient of hepatocyte growth factor (HGF), a known scattering factor. It is found that cells located near the higher concentration of HGF migrate faster than those on the opposite side within a cell island. Within the same island, cellular traction is similar on both sides, but intercellular stress is much lower on the side of higher HGF concentration. This novel experimental system can provide new opportunities to studying the mechanics of chemotactic migration by cellular collectives.

Collective cell migration in development, wound healing, and cancer metastasis is often guided by the gradients of growth factors or signaling molecules. Described here is an experimental system combining traction microscopy with a microfluidic system and a demonstration of how to quantify the mechanics of collective migration under biochemical gradient.

Cells change migration patterns in response to chemical stimuli, including the gradients of the stimuli. Cellular migration in the direction of a chemical ...

Production of Autologous Platelet-Rich Plasma for Boosting In Vitro Human Fibroblast Expansion

There is currently great clinical interest in the use of autologous fibroblasts for skin repair. In most cases, culture of skin cells in vitro is required. However, cell culture using xenogenic or allogenic culture media has some disadvantages (i.e., risk of infectious agent transmission or slow cell expansion). Here, an autologous culture system is developed for the expansion of human skin fibroblast cells in vitro using a patient&#8217;s own platelet-rich plasma (PRP). Human dermal fibroblasts are isolated from the patient while undergoing abdominoplasty. Cultures are followed for up to 7 days using a medium supplemented with either fetal bovine serum (FBS) or PRP. Blood cell content in PRP preparations, proliferation, and fibroblast differentiation are assessed. This protocol describes the method for obtaining a standardized, non-activated preparation of PRP using a dedicated medical device. The preparation requires only a medical device (CuteCell-PRP) and centrifuge. This device is suitable under sufficient medical practice conditions and is a one-step, apyrogenic, and sterile closed system that requires a single, soft spin centrifugation of 1,500 x g for 5 min. After centrifugation, the blood components are separated, and the platelet-rich plasma is easily collected. This device allows a quick, consistent, and standardized preparation of PRP that can be used as a cell culture supplement for in vitro expansion of human cells. The PRP obtained here contains a 1.5-fold platelet concentration compared to whole blood together, with a preferential removal of red and white blood cells. It is shown that PRP presents a boosting effect in cell proliferation compared to FBS (7.7x) and that fibroblasts are activated upon PRP treatment.

This protocol presents a device that produces PRP to boost the in vitro expansion of cells in a 100% autologous fibroblast culture system.

There is currently great clinical interest in the use of autologous fibroblasts for skin repair. In most cases, culture of skin cells in vitro is ...

In Vitro Model of Human Cutaneous Hypertrophic Scarring using Macromolecular Crowding

It has been shown that in vivo tissues are highly crowded by proteins, nucleic acids, ribonucleoproteins, polysaccharides, etc. The following protocol applies a macromolecular crowding (MMC) technique to mimic this physiological crowding through the addition of neutral polymers (crowders) to cell cultures in vitro. Previous studies using Ficoll or dextran as crowders demonstrate that the expression of collagen I and fibronectin in WI38 and WS-1 cell lines are significantly enhanced using the MMC technique. However, this technique has not been validated in primary hypertrophic scar-derived human skin fibroblasts (hHSFs). As hypertrophic scarring arises from the excessive deposition of collagen, this protocol aims to construct a collagen-rich in vitro hypertrophic scar model by applying the MMC technique with hHSFs. This optimized MMC model has been shown to possess more similarities with in vivo scar tissue compared to traditional 2-dimensional (2-D) cell culture systems. In addition, it is cost-effective, time-efficient, and ethically desirable compared to animal models. Therefore, the optimized model reported here offers an advanced &#8220;in vivo-like&#8221; model for hypertrophic scar-related studies.

This protocol describes the use of macromolecular crowding to create an in vitro human hypertrophic scar tissue model that resembles in vivo conditions. When cultivated in a crowded macromolecular environment, human skin fibroblasts exhibit phenotypes, biochemistry, physiology, and functional characteristics resembling scar tissue.

It has been shown that in vivo tissues are highly crowded by proteins, nucleic acids, ribonucleoproteins, polysaccharides, etc. The following protocol ...