This work was supported by the National Institutes of Health, Grant Number: GM114359 and GM134701 (M.F.K. and L.E.K.), 1F31AI164870-01 (J.C.L.), and Defense Threat Reduction Agency, Grant Number: HDTRA11910012 (M.F.K. and L.E.K.).

Heparinized human blood is obtained for neutrophil isolation from healthy adult donors (males and females, aged between 21 and 60 years old), following informed consent as approved by the Institutional Review Board of Temple University (Philadelphia, PA, USA).
1. Priming and coating the device with human fibronectin
NOTE: The bMFA has two inlet ports and two outlet ports connected to the vascular compartment. It also has one port connected to the tissue compartment (Figure 2).
<ol>
	<li>Insert tubing (O.D. of 0.06&#34; and I.D. of 0.02&#34;) (Table of Materials) to all the ports except for one inlet port with fine-point forceps. Clamp the two outlet ports and the tissue compartment port with jaw clamps. Ensure that each tubing is ~1 inch in length.</li>
	<li>Dilute human fibronectin to 100 &#181;g/mL with PBS from fibronectin stock solution (1mg/mL). Load a 1 mL syringe with prepared fibronectin solution. Connect the syringe to a 24 G blunt needle and a ~4-inch long tubing. 
	NOTE: To prevent crosslinking, do not over mix/pipette human fibronectin. Other extracellular matrices (ECM) such as collagen may be used for different cell types.</li>
	<li>Insert the tubing into the open inlet port, push the plunger until human fibronectin comes out of the other inlet port, then clamp it. Repeat this process for the rest of the ports until all the channels, tissue compartment and tubing are filled with the human fibronectin solution. Remove the needle but keep the 4-inch long tubing inserted and unclamped. 
	NOTE: Make sure the tubing is filled with fibronectin before it is clamped.</li>
	<li>For degassing, connect the unclamped tubing to&#160;a compressed nitrogen tank through the Pneumatic Primer, adjust the pressure to ~5 psi and run for ~15 min.</li>
	<li>After 15 min, check under the microscope to make sure there are no air bubbles trapped in the device. If there are air bubbles trapped in the channel or tissue compartment, reconnect the device to the Pneumatic Primer for degassing again, i.e., repeat step 1.4. 
	NOTE: Inverted microscope is used in all the steps involving microscopy in this protocol.</li>
	<li>Remove the device from the Pneumatic Primer. Incubate the device at 37 &#176;C for 1 h, then flush all the channels and tissue compartment with pre-warmed HLMVEC culture media (Table of Materials), similar to step 1.3. The bMFA is now ready for cell seeding. 
	&#8203;NOTE: Before removing/inserting a tubing from port, first place a drop of water with a pipette around the base of the inlet port tubing to prevent air from entering the device.</li>
</ol>
2. Seeding bMFA with HLMVEC
<ol>
	<li>Culture HLMVEC to 60%- 80% confluency according to the vendor&#39;s protocol in a T-25 flask.</li>
	<li>Aspirate HLMVEC culture media from cell culture flask. Wash twice with PBS.</li>
	<li>Add 2 mL of pre-warmed trypsin-EDTA solution (37 &#176;C) in the T-25 flask. Incubate for 3-5 min in an incubator (37 &#176;C, 5% CO2) until most cells are detached from the flask. 
	NOTE: Incubation time may vary for different cell types and trypsin-EDTA concentration.</li>
	<li>Add 2 mL of Trypsin Neutralization Solution (TNS, 37 &#176;C) to the flask to neutralize trypsin-EDTA.</li>
	<li>Gently tap the flask to help dissociate the cells. Then, transfer the cell suspension solution to a 15 mL conical tube.</li>
	<li>Centrifuge for 5 min at 150 x g to pellet the endothelial cells. Remove the supernatant and resuspend cells in 3 mL of cell culture media. Count the number of cells with a hemocytometer.</li>
	<li>Centrifuge again for 5 min at 150 x g to pellet the cells. Remove the supernatant and resuspend the cells to the desired seeding density of 20 x 106/mL. 
	NOTE: Depending on the cell types and confluency, about 2 x 106 endothelial cells can be collected from two T-25 flasks, and with the seeding density of 20 x 106/mL, 100 &#181;L of the cell suspension (2 x 106 endothelial cells), is sufficient to seed 5 devices.</li>
	<li>Mount a 1 mL syringe in the programmable syringe pump, attach tubing to the blunt needle.</li>
	<li>Draw ~20 &#181;L of the cell suspension into the tubing, making sure the cell suspension is only in the tubing, not in the syringe barrel.</li>
	<li>Take the clamp off an outlet port of the device. Connect the tubing to one inlet of the device. Make sure no air bubbles are introduced into the channel.</li>
	<li>Start the pump with a flow rate of 4-8 &#181;L/min. Observe the device under a microscope.</li>
	<li>When the channels are filled with cells, stop the pump, clamp the outlet, and cut the inlet tubing. 
	NOTE: Take care not to introduce air bubbles into the device.</li>
	<li>Put the device in the incubator for 4 h at 5% CO2 and 37 &#176;C.</li>
	<li>After 4 h incubation, prepare another syringe filled with fresh cell culture media. Mount the syringe on a syringe pump, connect the syringe to the inlet port&#160;and remove the clamp from an outlet port.</li>
	<li>Run fresh media through the device for about 5 min at 4-8 &#181;L/min to wash out floating/unattached cells.</li>
</ol>
3. Cell culture under flow for 48 h
<ol>
	<li>Prepare a syringe filled with fresh cell culture media. Mount the syringe in a syringe pump, connect the syringe to one inlet port&#160;and keep an outlet open. Put the device in the incubator (5% CO2, 37 &#176;C).</li>
	<li>Program the syringe pump for culturing endothelial cells under flow using the following instructions mentioned in Table 1.</li>
	<li>Check bMFA under a microscope after 48 h of culture under flow. HLMVEC should cover vascular channels forming a complete lumen (Figure 6). 
	NOTE: Other flow regimes may be required for different cell types.</li>
</ol>
4. Cytokine and/or therapeutic treatment
NOTE: As an example, this section describes using bMFA to study the impact of treating the cells with Tumor Necrosis Factor-alpha (TNF-&#945;) and a novel anti-inflammatory inhibitor (Protein Kinase C delta-TAT peptide inhibitor, PKC&#948;-i)18,27,32,33,34,35,36.
<ol>
	<li>Prepare three different bMFA devices using the protocol above (section 1-3).</li>
	<li>Load three 1 mL syringe with cell culture media,&#160;(TNF-&#945;) (10 ng/mL), TNF-&#945; (10 ng/mL) + PKC&#948;-i (5 &#181;M), respectively. 
	NOTE: Cytokines are reconstituted in cell culture media.</li>
	<li>Connect the three syringes to three bMFA devices. Treat HLMVEC with buffer TNF-&#945; or TNF-&#945; + inhibitor for 4 h at 0.1 &#181;L/min. 
	&#8203;NOTE: Based on specific study requirements, other cytokines or cytokine cocktails (e.g., TNF-&#945; + Interleukin 1 beta (IL1-&#946;) + Interferon-gamma (IFN-&#947;)), may be used to treat endothelial cells.</li>
</ol>
5. Human neutrophil isolation
<ol>
	<li>Use all reagents at room temperature (RT). Reagents include leukocyte isolation media , 1x HEPES buffer with Ca2+/Mg2+ (Table 2), 6% Dextran in normal saline (0.9% NaCl), 3.6% NaCl solution and de-ionized water (DI) water.</li>
	<li>Pipette 20 mL of leukocyte isolation media into a 50 mL conical tube and carefully layer 25 mL of blood over the leukocyte isolation media. Centrifuge for 40 min at 427 x g at RT. 
	NOTE: Perform this step slowly and carefully to avoid mixing the blood and leukocyte isolation media.</li>
	<li>Aspirate down to ~5 mL above red blood cells (RBC) layer. The white buffy layer on top of the RBC is predominantly neutrophils.</li>
	<li>Add HEPES buffer to double the current volume (existing volume: HEPES buffer = 1:1).</li>
	<li>Add an amount of 6% Dextran, which will be 20% of the final volume. (current volume/4 = volume of added 6% Dextran).</li>
	<li>Invert the tube gently a few times to mix well and let the RBC sediment (~25 min).</li>
	<li>Carefully pipette the upper layer (neutrophil-rich layer) and transfer to a new 50 mL tube, be careful not to contaminate with sedimented RBC.</li>
	<li>Centrifuge for 10 min at 315 x g at RT. Remove the supernatant and resuspend the pellet in 8 mL of HEPES buffer.</li>
	<li>To lyse the residual RBC, add 24 mL of DI water and gently mix for 50 s. Immediately, add 8 mL of 3.6% NaCl solution, mix and centrifuge for 10 min at 315 x g at RT.</li>
	<li>Remove the supernatant, wash the cell pellet with 15 mL of HEPES buffer and centrifuge for 5 min at 315 x g. Resuspend the pellet in HEPES buffer.</li>
</ol>
6. Neutrophil adhesion and migration experiment with bMFA
<ol>
	<li>Resuspend the isolated human neutrophils (15 million) in 999 &#181;L of HEPES buffer.</li>
	<li>Add 1 &#181;L CFDA-SE (carboxyfluorescein diacetate succinimidyl ester) dye stock solution (10 mM) to neutrophil suspension to obtain a working concentration of 10 &#181;M CFDA-SE and incubate for 10 min at RT. Wash the cells twice with HEPES buffer by centrifuging for 5 min at 315 x g.</li>
	<li>Count the neutrophils using a hemocytometer and resuspend at 2 million neutrophils/mL with cell culture media, TNF-&#945; (10 ng/mL) and TNF-&#945; (10 ng/mL) +PKC&#948;-i (5 &#956;M), respectively; incubate for 15 min at RT before the start of the experiment.</li>
	<li>Prepare a syringe filled with 1 &#181;M N-formyl-met-leu-phe (fMLP), the chemoattractant for the study, in cell culture media. Take bMFA and open one inlet port and one outlet port. 
	NOTE: These experimental conditions are used to mimic the conditions of sepsis that include a systemic inflammatory response with high levels of circulating cytokine/chemokines. Furthermore, fMLP, a Gram-negative bacteria-derived peptide, is used to model the presence of bacteria in the lung (i.e., tissue compartment).</li>
	<li>Remove the tissue compartment port tubing and insert the fMLP tubing. Inject ~20 &#181;L of fMLP into the tissue compartment for all bMFA, except the one treated with cell culture media alone. Then cut the tubing and clamp it.</li>
	<li>Fill the syringe with ~200 &#181;L of the neutrophil suspension and mount the syringe on the syringe pump.</li>
	<li>Put the device on the inverted microscope stage (Table of Materials).</li>
	<li>Start the pump at a flow rate is 1 &#181;L/min and wait until a small drop of the neutrophil suspension comes out of the tubing. Insert the tubing into the inlet port. See Figure 7 for the setup.</li>
</ol>
7. Acquire images
<ol>
	<li>Use the Scan Large Image function in the microscope image analysis software (Table of Materials) to obtain the adhesion map in bMFA (Figure 8) 10 min after starting the experiment. Most of the neutrophils enter bMFA during the first 10 min. 
	NOTE: Keep the fluorescence light off when not taking images to reduce photobleaching.</li>
	<li>During the next hour, take timelapse images (1 image every 5 min) of the tissue compartment (Figure 8B,C) for migration analysis to obtain the migration map.</li>
</ol>
8. Digital Image analysis
<ol>
	<li>For the adhesion map, count the number of adhered neutrophils in each vessel. For each bifurcation, create a circular region of interest (ROI) with a diameter equal to two times the vessel diameter and count the number of adhered neutrophils.</li>
	<li>Use manual count or the automated Object Count function to quantify the number of adhering neutrophils in each vessel and bifurcation region.</li>
	<li>Use the shear rate map of the network generated using CFD simulation17 to determine the shear rate in each vessel. Then plot the number of adhered neutrophils in a given vessel against the shear rate in that vessel to create a shear rate map in bMFA as shown in Figure 9A.</li>
	<li>Neutrophils are counted as migrated if they have crossed through the endothelium into the tissue compartment. Count the number of migrated neutrophils at 10 min, 15 min, 30 min and 60 min. Plot the number of migrated neutrophils versus time as shown in Figure 9B.</li>
</ol>

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J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biodistribution+and+efficacy+of+targeted+pulmonary+delivery+of+a+protein+kinase+C-d+inhibitory+peptide:+Impact+on+Indirect+lung+Injury.">Biodistribution and efficacy of targeted pulmonary delivery of a protein kinase C-d inhibitory peptide: Impact on Indirect lung Injury.</a> Journal of Pharmacology and Experimental Therapeutics. 355, 86-98 (2015).</li><li>Liverani, E., Mondrinos, M. J., Sun, S., Kunapuli, S. P., Kilpatrick, L. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+protein+kinase+C-delta+in+regulating+platelet+activation+and+platelet-leukocyte+interaction+during+sepsis.">Role of protein kinase C-delta in regulating platelet activation and platelet-leukocyte interaction during sepsis.</a> PloS one. 13, 0195379 (2018).</li><li>Smith, A. M., Prabhakarpandian, B., Pant, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+shear+adhesion+map+using+SynVivo+synthetic+microvascular+networks.">Generation of shear adhesion map using SynVivo synthetic microvascular networks.</a> Journal of Visualized Experiments: JoVE. (87), e51025 (2014).</li></ol>

The authors declare no conflict of interest.

The bMFA reproduces the topography and flow conditions of the in vivo microvascular networks and can be used to study leukocyte-endothelial cell interaction and endothelial function in vitro under physiologically realistic conditions. In the microvasculature of either mouse or human, the geometry of the microvascular networks are self-similar and fractal, and the Reynolds number &#60;&#60; 1, indicating that vascular geometry does not significantly impact flow patterns. Therefore, the bFMA can be used t...

Inflammation is the host response to infection and injury, and the endothelium plays an important role in the inflammatory response1,2,3. Inflammatory dysregulation is the underlying cause of a number of disease pathologies such as sepsis, cardiovascular diseases, asthma, inflammatory bowel disease, cancer&#160;and COVID-19. Leukocyte-endothelial cell interactions play a central role in these inflammatory diseases. During inflammation, the release of PAMPS (pathogen-associated molecular patterns) from pathogens or DAMPS (damage-associated molecular patterns) from injured tissues activate immune cells to release cytokines/chemokines and other proinflammatory mediators that lead to the activation of endothelium, resulting in alterations in vascular endothelium barrier function and increased permeability3,4. Increased activation of endothelial cells during inflammation results in enhanced leukocyte-endothelial cell interaction leading to excessive migration of activated leukocytes across the vascular endothelium into key organs1,5,6,7.
The recruitment of leukocytes is initiated by chemically diverse chemoattractants composed of bioactive lipids, cytokines, chemokines&#160;and complement components8,9. Leukocyte recruitment is a multi-step process that includes five discrete steps: 1) leukocyte margination and capture/attachment, 2) rolling, 3) firm arrest, 4) spreading and crawling and 5) extravasation/migration (Figure 1). Each step of this process requiring crosstalk between leukocytes and endothelial cells to orchestrate this dynamic phenomenon1,9. Ultimately, arrested leukocytes extravasate to inflamed tissues across endothelium via a multi-step process controlled by concurrent chemoattractant-dependent signals, adhesive events and hemodynamic shear forces1,9,10,11,12.
Given the central role of shear stress in regulating endothelial cell function and the significance of the leukocyte-endothelium cell interactions13, several in vitro models have been developed during the last few decades to study various aspects of the leukocyte migration cascade in a more controlled environment14. Traditional fluidic devices to study leukocyte-endothelial cell interactions can be classified into two broad categories14: a) devices for studying leukocyte rolling, adhesion&#160;and adhesion molecule expression such as parallel plate flow chambers and b) devices for studying leukocyte migration under static conditions such as transwell chambers. Systems like parallel plate flow chambers have been used to study the roles of adhesion molecules and their ligands in the adhesion cascade under shear forces15. However, a significant drawback is that these simplistic, idealized devices (e.g., straight channel) are not able to reproduce the scale and geometry of the in vivo microvasculature (e.g., successive vascular bifurcations, vascular morphology)&#160;and the resulting flow conditions (e.g., converging or diverging flows at bifurcations). As a result, these devices&#160;can only model adhesion but not transmigration. Transwell chambers can only study transmigration under&#160;static conditions without considering the in vivo geometrical features and flow conditions. Thus, these traditional models do not mimic the microenvironment of living tissues or resolve adhesion and migration cascade in a single assay6.
To address this limitation, we have developed and extensively validated a novel 3D biomimetic microfluidic assay (bMFA) (Figure 2), which realistically reproduces in vivo microvascular networks on a chip16,17,18. The protocol for microfabrication of this device has been published previously17&#160;and is only briefly described here. The microvasculature of mouse cremaster muscle was digitized using a modified Geographic Information System (GIS) approach19. Then, the synthetic microvascular network was generated on polydimethylsiloxane (PDMS) using soft-lithography processes based on the digitized microvascular network14,17,20,21,22. Briefly, the digitized network images were printed on Mylar film, which was then used as a mask to pattern a SU-8 positive photoresist on top of a silicon wafer to create the masters for fabrication. Microfabricated pillars (10 &#181;m diameter, 3 &#181;m tall) were used to create the 3 &#181;m high and 100 &#181;m wide pores, an optimum size for leukocyte migration23,24,25, connecting the vascular channels and tissue compartments. PDMS was prepared according to the manufacturer&#39;s instructions and poured over the developed masters. Further, the PDMS was degassed and allowed to cure overnight in an oven (65 &#176;C) to create complementary microchannels in PDMS. Subsequently, the cured PDMS was peeled from the SU-8 master, followed by punching ports for inlets/outlets. Then, the PMDS was plasma bonded to a glass slide. The surface of the microfluidic device comprises native glass and PDMS. In order to promote cell attachment, spreading&#160;and proliferation, extracelluar matrix (ECM) coating is required. The bMFA includes a microvascular network and a tissue compartment connected via 3 &#181;m high and 100 &#181;m wide pores (Figure 2). This microfluidic system reproduces the entire leukocyte adhesion/migration cascade in a physiologically relevant 3D environment of a complete microvascular network with interconnecting vessels and bifurcations, including circulation, margination, rolling, adhesion&#160;and migration of leukocytes into the extra-vascular tissue compartment in a single system14,16,17,21,26.
It should be noted that even when the flow rate at the inlet of bMFA is fixed, the flow conditions in the network vary at different locations and cannot be calculated by a simple mathematical formula. A computational fluid dynamics (CFD)-based model was developed to calculate different flow parameters (e.g., shear stress, shear rate, velocity) at different locations in the network. This CFD model was used to simulate the dye perfusion patterns and flow parameters in the bMFA. Cross-validation with experimental results suggested that the flow resistances across the network are well predicted by the computational model (Figure 3)17. This CFD model was then used to estimate velocity and shear rate profile in every vessel of bMFA (Figure 4), allowing analysis of the effects of shear flow and geometry on leukocyte rolling, adhesion and migration16. Leukocytes preferentially adhere near bifurcations and in low shear regions in vivo, and these spatial patterns of leukocyte adhesion were successfully demonstrated in bMFA using neutrophils (Figure 5)16. This paper describes the protocol for preparing bMFA to study leukocyte-endothelial cell interaction under inflammatory conditions using human lung microvascular endothelial cells (HLMVEC) and human neutrophils. Microphysiological systems, such as the bMFA, can be used to study endothelial cell interactions with different types of cells such as neutrophils, monocytes, lymphocytes&#160;and tumor cells18,27,28,29,30. The bMFA can be seeded with primary endothelial cells from different organs (e.g., lung vs. brain) and different species (e.g., human vs. murine endothelial cells), as well as endothelial cell lines21,27,31,32. The bMFA can be used to study multiple cellular responses, cell-cell interactions, barrier function, drug delivery&#160;and drug toxicity.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mL syringe</td>
			<td>Fisher Scientific</td>
			<td>14-823-30</td>
			<td></td>
		</tr>
		<tr>
			<td>Biomimetic microfluidic assay (bMFA)</td>
			<td>SynVivo</td>
			<td>SMN1-C001</td>
			<td>Exclusive at SynVivo</td>
		</tr>
		<tr>
			<td>Blunt needle</td>
			<td>Jensen Global</td>
			<td>JG24-0.5</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride</td>
			<td>Fisher Scientific</td>
			<td>C70-500</td>
			<td></td>
		</tr>
		<tr>
			<td>CFDA, SE</td>
			<td>ThermoFisher</td>
			<td>C1157</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextran, 250,000, Powder</td>
			<td>Spectrum Chemical Mfg. Corp</td>
			<td>DE-130</td>
			<td></td>
		</tr>
		<tr>
			<td>Ficoll-Paque Premium</td>
			<td>GE Health Care</td>
			<td>17-5442-02</td>
			<td>Leukocyte isolation media</td>
		</tr>
		<tr>
			<td>fMLP</td>
			<td>Sigma-Aldrich</td>
			<td>F3506</td>
			<td></td>
		</tr>
		<tr>
			<td>Hepes</td>
			<td>Fisher Scientific</td>
			<td>AAJ1692630</td>
			<td></td>
		</tr>
		<tr>
			<td>Human fibronectin</td>
			<td>Fisher Scientific</td>
			<td>33-016-015</td>
			<td>use vendor recommended ECM for different cell lines</td>
		</tr>
		<tr>
			<td>Microvascular Endothelial Cell Growth Medium-2 BulletKit</td>
			<td>Lonza</td>
			<td>cc-3202</td>
			<td>Human lung microvascular endothelial cell culture medium (HLMVEC).</td>
		</tr>
		<tr>
			<td>Human lung microvascular endothelial cells</td>
			<td>Lonza</td>
			<td>cc-2527</td>
			<td>use vedor remommended trypsin-EDTA and TNS</td>
		</tr>
		<tr>
			<td>Magnesium Chloride</td>
			<td>Fisher Scientific</td>
			<td>BP214-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon Eclipse Ti2</td>
			<td>Nikon Instruments Inc.</td>
			<td></td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>NIS-elements, 5.20.01</td>
			<td>Nikon Instruments Inc.</td>
			<td></td>
			<td>Imaging software</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Fisher Scientific</td>
			<td>MT21040CV</td>
			<td></td>
		</tr>
		<tr>
			<td>PhD Ultra Syringe Pump</td>
			<td>Harvard Apparatus</td>
			<td>70-3007</td>
			<td>Syringe Pump</td>
		</tr>
		<tr>
			<td>Potassium Hydroxide</td>
			<td>Fisher Scientific</td>
			<td>02-003-763</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Human TNF-alpha</td>
			<td>R&#38;D Systems</td>
			<td>210-TA</td>
			<td></td>
		</tr>
		<tr>
			<td>Slide clamp</td>
			<td>SynVivo</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Fisher Scientific</td>
			<td>S640-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Synvivo Pneumatic Primer</td>
			<td>SynVivo</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA, Trypsin Neutralization Solution(TNS)</td>
			<td>Lonza</td>
			<td>cc-5034</td>
			<td></td>
		</tr>
		<tr>
			<td>Tygon tubing</td>
			<td>Fisher Scientific</td>
			<td>50-206-8921</td>
			<td>Tubing</td>
		</tr>
	</tbody>
</table>

a microphysiological system to study leukocyte-endothelial cell interaction during inflammation

Leukocyte-endothelial cell interactions play an important role in inflammatory diseases such as sepsis. During inflammation, excessive migration of activated leukocytes across the vascular endothelium into key organs can lead to organ failure. A physiologically relevant biomimetic microfluidic assay (bMFA) has been developed and validated using several experimental and computational techniques, which can reproduce the entire leukocyte rolling/adhesion/migration cascade to study leukocyte-endothelial cell interactions. Microvascular networks obtained from in vivo images in rodents were digitized using a Geographic Information System (GIS) approach and microfabricated with polydimethylsiloxane (PDMS) on a microscope slide. To study the effect of shear rate and vascular topology on leukocyte-endothelial cell interactions, a Computational Fluid Dynamics (CFD) model was developed to generate a corresponding map of shear rates and velocities throughout the network. The bMFA enables the quantification of leukocyte-endothelial cells interactions, including rolling velocity, number of adhered leukocytes in response to different shear rates, number of migrated leukocytes, endothelial cell permeability, adhesion molecule expression and other important variables. Furthermore, by using human-related samples, such as human endothelial cells and leukocytes, bMFA provides a tool for rapid screening of potential therapeutics to increase their clinical translatability.

After 48 h of culture under shear flow in bMFA, endothelial cells covered the surface of the vascular channels in bMFA and aligned in the direction of flow (Figure&#160;6). Confocal microscopy indicated that all surfaces of the vascular channels were covered by endothelial cells, forming a complete 3D lumen in bMFA18.
Using this protocol, a neutrophil adhesion map can be acquired, showing that there is significant adhesion of neutrophils to...

Watch this Scientific Journal Video about A Microphysiological System to Study Leukocyte-Endothelial Cell Interaction during Inflammation at JoVE.com

A Microphysiological System to Study Leukocyte-Endothelial Cell Interaction during Inflammation

In this protocol, a biomimetic microfluid assay, which can reproduce a physiologically relevant microvascular environment and reproduce the entire leukocyte adhesion/migration cascade, is employed to study leukocyte-endothelial cell interactions in inflammatory disease.

Leukocyte-endothelial cell interactions play an important role in inflammatory diseases such as sepsis. During inflammation, excessive migration of ...

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JoVE Journal

Bioengineering

3.1K Views.  Temple University. In this protocol, a biomimetic microfluid assay, which can reproduce a physiologically relevant microvascular environment and reproduce the entire leukocyte adhesion/migration cascade, is employed to study leukocyte-endothelial cell interactions in inflammatory disease. 

Video: A Microphysiological System to Study Leukocyte-Endothelial Cell Interaction during Inflammation

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Constructing a Low-budget Laser Axotomy System to Study Axon Regeneration in C. elegans

Laser axotomy followed by time-lapse microscopy is a sensitive assay for axon regeneration phenotypes in C. elegans1. The main difficulty of this assay is the perceived cost ($25-100K) and technical expertise required for implementing a laser ablation system2,3. However, solid-state pulse lasers of modest costs (&lt;$10K) can provide robust performance for laser ablation in transparent preparations where target axons are "close" to the tissue surface. Construction and alignment of a system can be accomplished in a day. The optical path provided by light from the focused condenser to the ablation laser provides a convenient alignment guide. An intermediate module with all optics removed can be dedicated to the ablation laser and assures that no optical elements need be moved during a laser ablation session. A dichroic in the intermediate module allows simultaneous imaging and laser ablation. Centering the laser beam to the outgoing beam from the focused microscope condenser lens guides the initial alignment of the system. A variety of lenses are used to condition and expand the laser beam to fill the back aperture of the chosen objective lens. Final alignment and testing is performed with a front surface mirrored glass slide target. Laser power is adjusted to give a minimum size ablation spot (&lt;1um). The ablation spot is centered with fine adjustments of the last kinematically mounted mirror to cross hairs fixed in the imaging window. Laser power for axotomy will be approximately 10X higher than needed for the minimum ablation spot on the target slide (this may vary with the target you use). Worms can be immobilized for laser axotomy and time-lapse imaging by mounting on agarose pads (or in microfluidic chambers4). Agarose pads are easily made with 10% agarose in balanced saline melted in a microwave. A drop of molten agarose is placed on a glass slide and flattened with another glass slide into a pad approximately 200 um thick (a single layer of time tape on adjacent slides is used as a spacer). A "Sharpie" cap is used to cut out a uniformed diameter circular pad of 13mm. Anesthetic (1ul Muscimol 20mM) and Microspheres (Chris Fang-Yen personal communication) (1ul 2.65% Polystyrene 0.1 um in water) are added to the center of the pad followed by 3-5 worms oriented so they are lying on their left sides. A glass coverslip is applied and then Vaseline is used to seal the coverslip and prevent evaporation of the sample.

Constructing a Low-budget Laser Axotomy System to Study Axon Regeneration in C. elegans

Laser axotomy followed by time-lapse imaging is a sensitive way to assay the effects of mutations in C. elegans on axon regeneration. A high quality, but inexpensive, laser ablation system can be easily added to most microscopes. Time lapse imaging over 15 hours requires careful immobilization of the worm.

Laser axotomy followed by time-lapse microscopy is a sensitive assay for axon regeneration phenotypes in C. elegans1.  The main difficulty of this assay ...

An Experimental System to Study Mechanotransduction in Fetal Lung Cells

Mechanical forces generated in utero by repetitive breathing-like movements and by fluid distension are critical for normal lung development. A key component of lung development is the differentiation of alveolar type II epithelial cells, the major source of pulmonary surfactant. These cells also participate in fluid homeostasis in the alveolar lumen, host defense, and injury repair. In addition, distal lung parenchyma cells can be directly exposed to exaggerated stretch during mechanical ventilation after birth. However, the precise molecular and cellular mechanisms by which lung cells sense mechanical stimuli to influence lung development and to promote lung injury are not completely understood. Here, we provide a simple and high purity method to isolate type II cells and fibroblasts from rodent fetal lungs. Then, we describe an in vitro system, The Flexcell Strain Unit, to provide mechanical stimulation to fetal cells, simulating mechanical forces in fetal lung development or lung injury. This experimental system provides an excellent tool to investigate molecular and cellular mechanisms in fetal lung cells exposed to stretch. Using this approach, our laboratory has identified several receptors and signaling proteins that participate in mechanotransduction in fetal lung development and lung injury.

Mechanical forces play a key role in lung development and lung injury. Here, we describe a method to isolate rodent fetal lung type II epithelial cells and fibroblasts and to expose them to mechanical stimulation using an in vitro system.

Mechanical forces generated in utero by repetitive breathing-like movements and by fluid distension are critical for normal lung development. A key ...

Microfabrication of Nanoporous Gold Patterns for Cell-material Interaction Studies

Nanostructured materials with feature sizes in tens of nanometers have enhanced the performance of several technologies, including fuel cells, biosensors, biomedical device coatings, and drug delivery tools. Nanoporous gold (np-Au), produced by a nano-scale self-assembly process, is a relatively new material that exhibits large effective surface area, high electrical conductivity, and catalytic activity. These properties have made np-Au an attractive material to scientific community. Most studies on np-Au employ macro-scale specimens and focus on fundamental science of the material and its catalytic and sensor applications. The macro-scale specimens limit np-Au's potential in miniaturized systems, including biomedical devices. In order to address these issues, we initially describe two different methods to micropattern np-Au thin films on rigid substrates. The first method employs manually-produced stencil masks for creating millimeter-scale np-Au patterns, while the second method uses lift-off photolithography to pattern sub-millimeter-scale patterns. As the np-Au thin films are obtained by sputter-deposition process, they are compatible with conventional microfabrication techniques, thereby amenable to facile integration into microsystems. These systems include electrically-addressable biosensor platforms that benefit from high effective surface area, electrical conductivity, and gold-thiol-based surface bioconjugation. We describe cell culture, immunostaining, and image processing techniques to quantify np-Au's interaction with mammalian cells, which is an important performance parameter for some biosensors. We expect that the techniques illustrated here will assist the integration of np-Au in platforms at various length-scales and in numerous applications, including biosensors, energy storage systems, and catalysts.

We report on techniques to micropattern nanoporous gold thin films via stencil printing and photolithography, as well as methods to culture cells on the microfabricated patterns. In addition, we describe image analysis methods to characterize morphology of the material and the cultured cells using scanning electron and fluorescence microscopy techniques.

Nanostructured materials with feature sizes in tens of nanometers have enhanced the performance of several technologies, including fuel cells, biosensors, ...

Assessing Leukocyte-endothelial Interactions Under Flow Conditions in an Ex Vivo Autoperfused Microflow Chamber Assay

Leukocyte-endothelial interactions are early and critical events in acute and chronic inflammation and can, when dysregulated, mediate tissue injury leading to permanent pathological damage. Existing conventional assays allow the analysis of leukocyte adhesion molecules only after the extraction of leukocytes from the blood. This requires the blood to undergo several steps before peripheral blood leukocytes (PBLs) can be ready for analysis, which in turn can stimulate PBLs influencing the research findings. The autoperfused micro flow chamber assay, however, allows scientists to study early leukocytes functional dysregulation using the systemic flow of a live mouse while having the freedom of manipulating a coated chamber. Through a disease model, the functional expression of leukocyte adhesion molecules can be assessed and quantified in a micro-glass chamber coated with immobilized endothelial adhesion molecules ex vivo. In this model, the blood flows between the right common carotid artery and left external jugular vein of a live mouse under anesthesia, allowing the interaction of native PBLs in the chamber. Real-time experimental analysis is achieved with the assistance of an intravital microscope as well as a Harvard Apparatus pressure device. The application of a flow regulator at the input point of the glass chamber allows comparable physiological flow conditions amongst the experiments. Leukocyte rolling velocity is the main outcome and is measured using the National Institutes of Health open-access software ImageJ. In summary, the autoperfused micro flow chamber assay provides an optimal physiological environment to study leukocytes endothelial interaction and allows researchers to draw accurate conclusions when studying inflammation.

Assessing Leukocyte-endothelial Interactions Under Flow Conditions in an Ex Vivo Autoperfused Microflow Chamber Assay

Here, we present a protocol that allows the investigator to assess leukocyte recruitment dynamics ex vivo by connecting a chamber coated with endothelial-derived adhesion molecules to the circulatory system of a mouse. This method offers significant advantages since it allows for leukocyte assessment under relative biological conditions.

Leukocyte-endothelial interactions are early and critical events in acute and chronic inflammation and can, when dysregulated, mediate tissue injury ...

Concentric Gel System to Study the Biophysical Role of Matrix Microenvironment on 3D Cell Migration

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to tumor and cancer metastasis. Despite intense research efforts, the basic biochemical and biophysical principles of cell migration are still not fully understood, especially in the physiologically relevant three-dimensional (3D) microenvironments. Here, we describe an in vitro assay designed to allow quantitative examination of 3D cell migration behaviors. The method exploits the cell&#8217;s mechanosensing ability and propensity to migrate into previously unoccupied extracellular matrix (ECM). We use the invasion of highly invasive breast cancer cells, MDA-MB-231, in collagen gels as a model system. The spread of cell population and the migration dynamics of individual cells over weeks of culture can be monitored using live-cell imaging and analyzed to extract spatiotemporally-resolved data. Furthermore, the method is easily adaptable for diverse extracellular matrices, thus offering a simple yet powerful way to investigate the role of biophysical factors in the microenvironment on cell migration.

The mechanical properties and microstructure of the extracellular matrix strongly affect 3D migration of cells. An in vitro method to study the spatiotemporal cell migration behavior in biophysically variable environments, at both population and individual cell levels, is described.

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to ...

Live Cell Imaging during Mechanical Stretch

There is currently a significant interest in understanding how cells and tissues respond to mechanical stimuli, but current approaches are limited in their capability for measuring responses in real time in live cells or viable tissue. A protocol was developed with the use of a cell actuator to distend live cells grown on or tissues attached to an elastic substrate while imaging with confocal and atomic force microscopy (AFM). Preliminary studies show that tonic stretching of human bronchial epithelial cells caused a significant increase in the production of mitochondrial superoxide. Moreover, using this protocol, alveolar epithelial cells were stretched and imaged, which showed direct damage to the epithelial cells by overdistention simulating one form of lung injury in vitro. A protocol to conduct AFM nano-indentation on stretched cells is also provided.

A novel imaging protocol was developed using a custom motor-driven mechanical actuator to allow the measurement of real time responses to mechanical strain in live cells. Relevant to mechanobiology, the system can apply strains up to 20% while allowing near real-time imaging with confocal or atomic force microscopy.

There is currently a significant interest in understanding how cells and tissues respond to mechanical stimuli, but current approaches are limited in ...

Construction of Defined Human Engineered Cardiac Tissues to Study Mechanisms of Cardiac Cell Therapy

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that replicate the biofidelity of three-dimensional native human myocardium, while also enabling a controlled level of biological complexity, and allowing non-destructive longitudinal monitoring of tissue contractile function. Initially, human engineered cardiac tissues (hECT) were created using the entire cell population obtained from directed differentiation of human pluripotent stem cells, which typically yielded less than 50% cardiomyocytes. However, to create reliable predictive models of human myocardium, and to elucidate mechanisms of heterocellular interaction, it is essential to accurately control the biological composition in engineered tissues. 
To address this limitation, we utilize live cell sorting for the cardiac surface marker SIRP&#945; and the fibroblast marker CD90 to create tissues containing a 3:1 ratio of these cell types, respectively, that are then mixed together and added to a collagen-based matrix solution. Resulting hECTs are, thus, completely defined in both their cellular and extracellular matrix composition.
Here we describe the construction of defined hECTs as a model system to understand mechanisms of cell-cell interactions in cell therapies, using an example of human bone marrow-derived mesenchymal stem cells (hMSC) that are currently being used in human clinical trials. The defined tissue composition is imperative to understand how the hMSCs may be interacting with the endogenous cardiac cell types to enhance tissue function. A bioreactor system is also described that simultaneously cultures six hECTs in parallel, permitting more efficient use of the cells after sorting.

This manuscript describes the creation of defined engineered cardiac tissues using surface marker expression and cell sorting. The defined tissues can then be used in a multi-tissue bioreactor to investigate mechanisms of cardiac cell therapy in order to provide a functional, yet controlled, model system of the human heart.

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that ...

A Microfluidic System with Surface Patterning for Investigating Cavitation Bubble(s)&#8211;Cell Interaction and the Resultant Bioeffects at the Single-cell Level

In this manuscript, we first describe the fabrication protocol of a microfluidic chip, with gold dots and fibronectin-coated regions on the same glass substrate, that precisely controls the generation of tandem bubbles and individual cells patterned nearby with well-defined locations and shapes. We then demonstrate the generation of tandem bubbles by using two pulsed lasers illuminating a pair of gold dots with a few-microsecond time delay. We visualize the bubble-bubble interaction and jet formation by high-speed imaging and characterize the resultant flow field using particle image velocimetry (PIV). Finally, we present some applications of this technique for single cell analysis, including cell membrane poration with macromolecule uptake, localized membrane deformation determined by the displacements of attached integrin-binding beads, and intracellular calcium response from ratiometric imaging. Our results show that a fast and directional jetting flow is produced by the tandem bubble interaction, which can impose a highly localized shear stress on the surface of a cell grown in close proximity. Furthermore, different bioeffects can be induced by altering the strength of the jetting flow by adjusting the standoff distance from the cell to the tandem bubbles.

A Microfluidic System with Surface Patterning for Investigating Cavitation Bubble(s)&amp;#8211;Cell Interaction and the Resultant Bioeffects at the Single-cell Level

A microfluidic chip was fabricated to produce pairs of gold dots for tandem bubble generation and fibronectin-coated islands for single-cell patterning nearby. The resultant flow field was characterized by particle image velocimetry and was employed to study various bioeffects, including cell membrane poration, membrane deformation, and intracellular calcium response.

In this manuscript, we first describe the fabrication protocol of a microfluidic chip, with gold dots and fibronectin-coated regions on the same glass ...

DNA Nanotubes as a Versatile Tool to Study Semiflexible Polymers

Mechanical properties of complex, polymer-based soft matter, such as cells or biopolymer networks, can be understood in neither the classical frame of flexible polymers nor of rigid rods. Underlying filaments remain outstretched due to their non-vanishing backbone stiffness, which is quantified via the persistence length (lp), but they are also subject to strong thermal fluctuations. Their finite bending stiffness leads to unique, non-trivial collective mechanics of bulk networks, enabling the formation of stable scaffolds at low volume fractions while providing large mesh sizes. This underlying principle is prevalent in nature (e.g., in cells or tissues), minimizing the high molecular content and thereby facilitating diffusive or active transport. Due to their biological implications and potential technological applications in biocompatible hydrogels, semiflexible polymers have been subject to considerable study. However, comprehensible investigations remained challenging since they relied on natural polymers, such as actin filaments, which are not freely tunable. Despite these limitations and due to the lack of synthetic, mechanically tunable, and semiflexible polymers, actin filaments were established as the common model system. A major limitation is that the central quantity lp cannot be freely tuned to study its impact on macroscopic bulk structures. This limitation was resolved by employing structurally programmable DNA nanotubes, enabling controlled alteration of the filament stiffness. They are formed through tile-based designs, where a discrete set of partially complementary strands hybridize in a ring structure with a discrete circumference. These rings feature sticky ends, enabling the effective polymerization into filaments several microns in length, and display similar polymerization kinetics as natural biopolymers. Due to their programmable mechanics, these tubes are versatile, novel tools to study the impact of lp on the single-molecule as well as the bulk scale. In contrast to actin filaments, they remain stable over weeks, without notable degeneration, and their handling is comparably straightforward.

Semiflexible polymers display unique mechanical properties that are extensively applied by living systems. However, systematic studies on biopolymers are limited since properties such as polymer rigidity are inaccessible. This manuscript describes how this limitation is circumvented by programmable DNA nanotubes, enabling experimental studies on the impact of filament rigidity.

Mechanical properties of complex, polymer-based soft matter, such as cells or biopolymer networks, can be understood in neither the classical frame of ...