Method Article

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

DOI:

10.3791/63312

⸱

December 9th, 2021

 ,  ,  ,  ,  , 
1Department of Mechanical Engineering, Temple University, 2Department of Bioengineering, Temple University, 3Department of Bioengineering, University of Toledo, 4Biomedical Technology, CFD Research Corporation, 5Center for Inflammation and Lung Research, Department of Microbiology, Immunology and Inflammation, Lewis Katz School of Medicine, Temple University, 6Department of Radiation Oncology, Lewis Katz School of Medicine, Temple University

In This Article

Summary

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

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.

Abstract

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

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.

Introduction

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

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 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 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 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) and the resulting flow conditions (e.g., converging or diverging flows at bifurcations). As a result, these devices can only model adhesion but not transmigration. Transwell chambers can only study transmigration under 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 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 µm diameter, 3 µm tall) were used to create the 3 µm high and 100 µm wide pores, an optimum size for leukocyte migration23,24,25, connecting the vascular channels and tissue compartments. PDMS was prepared according to the manufacturer's instructions and poured over the developed masters. Further, the PDMS was degassed and allowed to cure overnight in an oven (65 °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 and proliferation, extracelluar matrix (ECM) coating is required. The bMFA includes a microvascular network and a tissue compartment connected via 3 µm high and 100 µm 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 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 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 and drug toxicity.

Access restricted. Please log in or start a trial to view this content.

Protocol

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

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).

  1. Insert tubing (O.D. of 0.06" and I.D. of 0.02") (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.
  2. Dilute human fibronectin to 100 µg/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.
  3. 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.
  4. For degassing, connect the unclamped tubing to a compressed nitrogen tank through the Pneumatic Primer, adjust the pressure to ~5 psi and run for ~15 min.
  5. 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.
  6. Remove the device from the Pneumatic Primer. Incubate the device at 37 °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.
    ​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.

2. Seeding bMFA with HLMVEC

  1. Culture HLMVEC to 60%- 80% confluency according to the vendor's protocol in a T-25 flask.
  2. Aspirate HLMVEC culture media from cell culture flask. Wash twice with PBS.
  3. Add 2 mL of pre-warmed trypsin-EDTA solution (37 °C) in the T-25 flask. Incubate for 3-5 min in an incubator (37 °C, 5% CO2) until most cells are detached from the flask.
    NOTE: Incubation time may vary for different cell types and trypsin-EDTA concentration.
  4. Add 2 mL of Trypsin Neutralization Solution (TNS, 37 °C) to the flask to neutralize trypsin-EDTA.
  5. Gently tap the flask to help dissociate the cells. Then, transfer the cell suspension solution to a 15 mL conical tube.
  6. 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.
  7. 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 µL of the cell suspension (2 x 106 endothelial cells), is sufficient to seed 5 devices.
  8. Mount a 1 mL syringe in the programmable syringe pump, attach tubing to the blunt needle.
  9. Draw ~20 µL of the cell suspension into the tubing, making sure the cell suspension is only in the tubing, not in the syringe barrel.
  10. 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.
  11. Start the pump with a flow rate of 4-8 µL/min. Observe the device under a microscope.
  12. 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.
  13. Put the device in the incubator for 4 h at 5% CO2 and 37 °C.
  14. 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 and remove the clamp from an outlet port.
  15. Run fresh media through the device for about 5 min at 4-8 µL/min to wash out floating/unattached cells.

3. Cell culture under flow for 48 h

  1. Prepare a syringe filled with fresh cell culture media. Mount the syringe in a syringe pump, connect the syringe to one inlet port and keep an outlet open. Put the device in the incubator (5% CO2, 37 °C).
  2. Program the syringe pump for culturing endothelial cells under flow using the following instructions mentioned in Table 1.
  3. 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.

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-α) and a novel anti-inflammatory inhibitor (Protein Kinase C delta-TAT peptide inhibitor, PKCδ-i)18,27,32,33,34,35,36.

  1. Prepare three different bMFA devices using the protocol above (section 1-3).
  2. Load three 1 mL syringe with cell culture media, (TNF-α) (10 ng/mL), TNF-α (10 ng/mL) + PKCδ-i (5 µM), respectively.
    NOTE: Cytokines are reconstituted in cell culture media.
  3. Connect the three syringes to three bMFA devices. Treat HLMVEC with buffer TNF-α or TNF-α + inhibitor for 4 h at 0.1 µL/min.
    ​NOTE: Based on specific study requirements, other cytokines or cytokine cocktails (e.g., TNF-α + Interleukin 1 beta (IL1-β) + Interferon-gamma (IFN-γ)), may be used to treat endothelial cells.

5. Human neutrophil isolation

  1. 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.
  2. 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.
  3. Aspirate down to ~5 mL above red blood cells (RBC) layer. The white buffy layer on top of the RBC is predominantly neutrophils.
  4. Add HEPES buffer to double the current volume (existing volume: HEPES buffer = 1:1).
  5. Add an amount of 6% Dextran, which will be 20% of the final volume. (current volume/4 = volume of added 6% Dextran).
  6. Invert the tube gently a few times to mix well and let the RBC sediment (~25 min).
  7. Carefully pipette the upper layer (neutrophil-rich layer) and transfer to a new 50 mL tube, be careful not to contaminate with sedimented RBC.
  8. Centrifuge for 10 min at 315 x g at RT. Remove the supernatant and resuspend the pellet in 8 mL of HEPES buffer.
  9. 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.
  10. 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.

6. Neutrophil adhesion and migration experiment with bMFA

  1. Resuspend the isolated human neutrophils (15 million) in 999 µL of HEPES buffer.
  2. Add 1 µL CFDA-SE (carboxyfluorescein diacetate succinimidyl ester) dye stock solution (10 mM) to neutrophil suspension to obtain a working concentration of 10 µM 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.
  3. Count the neutrophils using a hemocytometer and resuspend at 2 million neutrophils/mL with cell culture media, TNF-α (10 ng/mL) and TNF-α (10 ng/mL) +PKCδ-i (5 μM), respectively; incubate for 15 min at RT before the start of the experiment.
  4. Prepare a syringe filled with 1 µM 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).
  5. Remove the tissue compartment port tubing and insert the fMLP tubing. Inject ~20 µL 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.
  6. Fill the syringe with ~200 µL of the neutrophil suspension and mount the syringe on the syringe pump.
  7. Put the device on the inverted microscope stage (Table of Materials).
  8. Start the pump at a flow rate is 1 µL/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.

7. Acquire images

  1. 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.
  2. 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.

8. Digital Image analysis

  1. 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.
  2. Use manual count or the automated Object Count function to quantify the number of adhering neutrophils in each vessel and bifurcation region.
  3. 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.
  4. 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.

Access restricted. Please log in or start a trial to view this content.

Results

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

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 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 endo...

Access restricted. Please log in or start a trial to view this content.

Discussion

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

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 << 1, indicating that vascular geometry does not significantly impact flow patterns. Therefore, the bFMA can be used t...

Access restricted. Please log in or start a trial to view this content.

Disclosures

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

The authors declare no conflict of interest.

Acknowledgements

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

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.).

Access restricted. Please log in or start a trial to view this content.

Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1 mL syringeFisher Scientific14-823-30
Biomimetic microfluidic assay (bMFA)SynVivoSMN1-C001Exclusive at SynVivo
Blunt needleJensen GlobalJG24-0.5
Calcium ChlorideFisher ScientificC70-500
CFDA, SEThermoFisherC1157
Dextran, 250,000, PowderSpectrum Chemical Mfg. CorpDE-130
Ficoll-Paque PremiumGE Health Care17-5442-02Leukocyte isolation media
fMLPSigma-AldrichF3506
HepesFisher ScientificAAJ1692630
Human fibronectinFisher Scientific33-016-015use vendor recommended ECM for different cell lines
Microvascular Endothelial Cell Growth Medium-2 BulletKitLonzacc-3202Human lung microvascular endothelial cell culture medium (HLMVEC).
Human lung microvascular endothelial cellsLonzacc-2527use vedor remommended trypsin-EDTA and TNS
Magnesium ChlorideFisher ScientificBP214-500
Nikon Eclipse Ti2Nikon Instruments Inc.Microscope
NIS-elements, 5.20.01Nikon Instruments Inc.Imaging software
PBSFisher ScientificMT21040CV
PhD Ultra Syringe PumpHarvard Apparatus70-3007Syringe Pump
Potassium HydroxideFisher Scientific02-003-763
Recombinant Human TNF-alphaR&D Systems210-TA
Slide clampSynVivo
Sodium ChlorideFisher ScientificS640-500
Synvivo Pneumatic PrimerSynVivo
Trypsin-EDTA, Trypsin Neutralization Solution(TNS)Lonzacc-5034
Tygon tubingFisher Scientific50-206-8921Tubing

References

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,
  1. Kolaczkowska, E., Kubes, P. Neutrophil recruitment and function in health and inflammation. Nature Reviews Immunology. 13 (3), 159-175 (2013).
  2. Yang, Q., Wijerathne, H., Langston, J. C., Kiani, M. F., Kilpatrick, L. E. Emerging approaches to understanding microvascular endothelial heterogeneity: A roadmap for developing anti-inflammatory therapeutics. International Journal of Molecular Sciences. 22 (15), 7770(2021).
  3. Joffre, J., Hellman, J., Ince, C., Ait-Oufella, H. Endothelial responses in sepsis. American Journal of Respiratory and Critical Care Medicine. 202, 361-370 (2020).
  4. Ince, C., et al. The endothelium in sepsis. Shock. 45, Augusta, Ga. 259-270 (2016).
  5. Ruparelia, N., Chai, J. T., Fisher, E. A., Choudhury, R. P. Inflammatory processes in cardiovascular disease: a route to targeted therapies. Nature Reviews Cardiology. 14 (3), 133-144 (2017).
  6. Kilpatrick, L. E., Kiani, M. F. Experimental approaches to evaluate leukocyte-endothelial cell interactions in sepsis and inflammation. Shock. 53, Augusta, Ga. 585-595 (2020).
  7. Langer, H. F., Chavakis, T. Leukocyte-endothelial interactions in inflammation. Journal of Cellular and Molecular Medicine. 13 (7), 1211-1220 (2009).
  8. Sadik, C. D., Luster, A. D. Lipid-cytokine-chemokine cascades orchestrate leukocyte recruitment in inflammation. Journal of Leukocyte Biology. 91 (2), 207-215 (2012).
  9. Phillipson, M., Kubes, P. The neutrophil in vascular inflammation. Nature Medicine. 17, 1381-1390 (2011).
  10. Maniatis, N. A., Orfanos, S. E. The endothelium in acute lung injury/acute respiratory distress syndrome. Current Opinion in Critical Care. 14, 22-30 (2008).
  11. Ley, K., Laudanna, C., Cybulsky, M. I., Nourshargh, S. Getting to the site of inflammation: the leukocyte adhesion cascade updated. Nature Reviews Immunology. 7, 678-689 (2007).
  12. Molteni, R., Fabbri, M., Bender, J. R., Pardi, R. Pathophysiology of leukocyte-tissue interactions. Current Opinion in Cell Biology. 18, 491-498 (2006).
  13. Chistiakov, D. A., Orekhov, A. N., Bobryshev, Y. V. Effects of shear stress on endothelial cells: go with the flow. Acta Physiologica. 219 (2), 382-408 (2017).
  14. Prabhakarpandian, B., Shen, M. -C., Pant, K., Kiani, M. F. Microfluidic devices for modeling cell-cell and particle-cell interactions in the microvasculature. Microvascular Research. 82, 210-220 (2011).
  15. Zou, X., et al. PSGL-1 derived from human neutrophils is a high-efficiency ligand for endothelium-expressed E-selectin under flow. American Journal of Physiology-Cell Physiology. 289, 415-424 (2005).
  16. Lamberti, G., et al. Bioinspired microfluidic assay for in vitro modeling of leukocyte-endothelium interactions. Analytical Chemistry (Journal). 86, 8344-8351 (2014).
  17. Prabhakarpandian, B., et al. Synthetic microvascular networks for quantitative analysis of particle adhesion. Biomedical Microdevices. 10 (4), 585-595 (2008).
  18. Soroush, F., et al. A novel microfluidic assay reveals a key role for protein kinase C delta in regulating human neutrophil-endothelium interaction. Journal of Leukocyte Biology. 100, 1027-1035 (2016).
  19. Roth, N. M., Kiani, M. F. A "geographic information systems" based technique for the study of microvascular networks. Annals of Biomedical Engineering. 27, 42-47 (1999).
  20. Prabhakarpandian, B., Wang, Y. I., Rea-Ramsey, A., Sundaram, S., Kiani, M. F., Pant, K. Bifurcations: Focal points of particle adhesion in microvascular networks. Microcirculation. 18, 380-389 (2011).
  21. Rosano, J., et al. A physiologically realistic in vitro model of microvascular networks. Biomedical Microdevices. 11, 1051-1057 (2009).
  22. Tousi, N., Wang, B., Pant, K., Kiani, M. F., Prabhakarpandian, B. Preferential adhesion of leukocytes near bifurcations is endothelium independent. Microvascular Research. 80, 384-388 (2010).
  23. Yona, S., Hayhoe, R., Avraham-Davidi, I. Monocyte and neutrophil isolation and migration assays. Current Protocols in Immunology. 88 (1), 11-14 (2010).
  24. Entschladen, F., et al. Analysis methods of human cell migration. Experimental Cell Research. 307 (2), 418-426 (2005).
  25. Chen, H. -C. Boyden chamber assay. Methods in Molecular Biology. Cell Migration: Developmental Methods and Protocols. , Humana Press Inc. Totowa, NJ. (2005).
  26. Deosarkar, S. P., Prabhakarpandian, B., Wang, B., Sheffield, J. B., Krynska, B., Kiani, M. F. A novel dynamic neonatal blood-brain barrier on a chip. PloS one. 10, 142725(2015).
  27. Soroush, F., et al. Neutrophil-endothelial interactions of murine cells is not a good predictor of their interactions in human cells. The FASEB Journal. 34, 2691-2702 (2020).
  28. Soroush, F., et al. Protein Kinase C-Delta (PKCδ) tyrosine phosphorylation is a critical regulator of neutrophil-endothelial cell interaction in inflammation. Shock. 51 (5), 538-547 (2019).
  29. Soroush, F., Tang, Y., Zaidi, H. M., Sheffield, J. B., Kilpatrick, L. E., Kiani, M. F. PKCδ inhibition as a novel medical countermeasure for radiation-induced vascular damage. The FASEB Journal. 32, 6436-6444 (2018).
  30. Pradhan, S., et al. A microvascularized tumor-mimetic platform for assessing anti-cancer drug efficacy. Scientific Reports. 8 (1), 3171(2018).
  31. Tang, Y., et al. A biomimetic microfluidic tumor microenvironment platform mimicking the EPR effect for rapid screening of drug delivery systems. Scientific Reports. 7 (1), 1-14 (2017).
  32. Tang, Y., et al. Protein kinase C-delta inhibition protects blood-brain barrier from sepsis-induced vascular damage. Journal of Neuroinflammation. 15, 309(2018).
  33. Mondrinos, M. J., et al. Pulmonary endothelial protein kinase C-delta (PKCd) regulates neutrophil migration in acute lung inflammation. The American Journal of Pathology. 184, 200-213 (2014).
  34. Kilpatrick, L. E., et al. Protection against sepsis-induced lung injury by selective inhibition of protein kinase C-d (d-PKC). Journal of Leukocyte Biology. 89, 3-10 (2011).
  35. Mondrinos, M. J., et al. Biodistribution and efficacy of targeted pulmonary delivery of a protein kinase C-d inhibitory peptide: Impact on Indirect lung Injury. Journal of Pharmacology and Experimental Therapeutics. 355, 86-98 (2015).
  36. Liverani, E., Mondrinos, M. J., Sun, S., Kunapuli, S. P., Kilpatrick, L. E. Role of protein kinase C-delta in regulating platelet activation and platelet-leukocyte interaction during sepsis. PloS one. 13, 0195379(2018).
  37. Smith, A. M., Prabhakarpandian, B., Pant, K. Generation of shear adhesion map using SynVivo synthetic microvascular networks. Journal of Visualized Experiments: JoVE. (87), e51025(2014).

Access restricted. Please log in or start a trial to view this content.

Reprints and Permissions

Request permission to reuse the text or figures of this JoVE article

Request Permission

Tags

Leukocyte Endothelial InteractionBiomimetic Microfluidic AssayInflammatory Disease ModelNeutrophil MigrationEndothelial Cell PermeabilityConfocal MicroscopyShear Rate AnalysisAdhesion Molecule ExpressionComputational Fluid Dynamics

Related Articles