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Research Papers

In Vitro Evaluation of the Link Between Cell Activation State and Its Rheological Impact on the Microscale Flow of Neutrophil Suspensions

[+] Author and Article Information
Michael L. Akenhead, Nolan M. Horrall

Department of Biomedical Engineering,
University of Kentucky,
Lexington, KY 40506-0070

Dylan Rowe

Math, Science and Technology Center,
Paul L. Dunbar High School,
Lexington, KY 40513

Palaniappan Sethu

Division of Cardiovascular Disease,
University of Alabama-Birmingham,
Birmingham, AL 35294-0006

Hainsworth Y. Shin

Department of Biomedical Engineering,
University of Kentucky,
Lexington, KY 40506-0070
e-mail: hy.shin@uky.edu

1M. L. Akenhead and N. M. Horrall contributed equally to this work.

2Corresponding author.

Manuscript received November 17, 2014; final manuscript received June 6, 2015; published online July 9, 2015. Assoc. Editor: Mohammad Mofrad.

J Biomech Eng 137(9), 091003 (Sep 01, 2015) (10 pages) Paper No: BIO-14-1569; doi: 10.1115/1.4030824 History: Received November 17, 2014; Revised June 06, 2015; Online July 09, 2015

Activated neutrophils have been reported to affect peripheral resistance, for example, by plugging capillaries or adhering to the microvasculature. In vivo and ex vivo data indicate that activated neutrophils circulating in the blood also influence peripheral resistance. We used viscometry and microvascular mimics for in vitro corroboration. The rheological impact of differentiated neutrophil-like HL-60 promyelocytes (dHL60s) or human neutrophil suspensions stimulated with 10 nM fMet-Leu-Phe (fMLP) was quantified using a cone-plate rheometer (450 s−1 shear rate). To evaluate their impact on microscale flow resistance, we used 10-μm Isopore® membranes to model capillaries as well as single 200 × 50 μm microchannels and networks of twenty 20 × 50 μm microfluidic channels to mimic noncapillary microvasculature. Stimulation of dHL60 and neutrophil populations significantly altered their flow behavior as evidenced by their impact on suspension viscosity. Notably, hematocrit abrogated the impact of leukocyte activation on blood cell suspension viscosity. In micropore filters, activated cell suspensions enhanced flow resistance. This effect was further enhanced by the presence of erythrocytes. The resistance of our noncapillary microvascular mimics to flow of activated neutrophil suspensions was significantly increased only with hematocrit. Notably, it was elevated to a higher extent within the micronetwork chambers compared to the single-channel chambers. Collectively, our findings provide supportive evidence that activated neutrophils passing through the microcirculation may alter hemodynamic resistance due to their altered rheology in the noncapillary microvasculature. This effect is another way neutrophil activation due to chronic inflammation may, at least in part, contribute to the elevated hemodynamic resistance associated with cardiovascular diseases (e.g., hypertension and hypercholesterolemia).

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References

Granger, D. N., and Senchenkova, E., 2010, “Leukocyte-Endothelial Cell Adhesion,” Inflammation and the Microcirculation (Integrated Systems Physiology—From Cell to Function), Morgan & Claypool Life Sciences, San Rafael, CA. [PubMed] [PubMed]
Baskurt, O. K., and Meiselman, H. J., 2003, “Blood Rheology and Hemodynamics,” Semin. Thromb. Hemostasis, 29(5), pp. 435–450. [CrossRef]
Popel, A. S., and Johnson, P. C., 2005, “Microcirculation and Hemorheology,” Annu. Rev. Fluid Mech., 37, pp. 43–69. [CrossRef] [PubMed]
Sethu, P., Moldawer, L. L., Mindrinos, M. N., Scumpia, P. O., Tannahill, C. L., Wilhelmy, J., Efron, P. A., Brownstein, B. H., Tompkins, R. G., and Toner, M., 2006, “Microfluidic Isolation of Leukocytes From Whole Blood for Phenotype and Gene Expression Analysis,” Anal. Chem., 78(15), pp. 5453–5461. [CrossRef] [PubMed]
Schmid-Schonbein, G. W., 2006, “Analysis of Inflammation,” Annu. Rev. Biomed. Eng., 8, pp. 93–131. [CrossRef] [PubMed]
Lipowsky, H. H., 2005, “Microvascular Rheology and Hemodynamics,” Microcirculation, 12(1), pp. 5–15. [CrossRef] [PubMed]
Mazzoni, M. C., and Schmid-Schonbein, G. W., 1996, “Mechanisms and Consequences of Cell Activation in the Microcirculation,” Cardiovas. Res., 32(4), pp. 709–719. [CrossRef]
Arndt, H., Smith, C. W., and Granger, D. N., 1993, “Leukocyte-Endothelial Cell Adhesion in Spontaneously Hypertensive and Normotensive Rats,” Hypertension, 21(5), pp. 667–673. [CrossRef] [PubMed]
Williams, S. A., and Tooke, J. E., 1992, “Noninvasive Estimation of Increased Structurally-Based Resistance to Blood Flow in the Skin of Subjects With Essential Hypertension,” Int. J. Microcirc., Clin. Exp., 11(1), pp. 109–116.
Worthen, G. S., Schwab, B., 3rd, Elson, E. L., and Downey, G. P., 1989, “Mechanics of Stimulated Neutrophils: Cell Stiffening Induces Retention in Capillaries,” Science, 245(4914), pp. 183–186. [CrossRef] [PubMed]
Schmid-Schonbein, G. W., Usami, S., Skalak, R., and Chien, S., 1980, “The Interaction of Leukocytes and Erythrocytes in Capillary and Postcapillary Vessels,” Microvas. Res., 19(1), pp. 45–70. [CrossRef]
Eppihimer, M. J., and Lipowsky, H. H., 1996, “Effects of Leukocyte-Capillary Plugging on the Resistance to Flow in the Microvasculature of Cremaster Muscle for Normal and Activated Leukocytes,” Microvas. Res., 51(2), pp. 187–201. [CrossRef]
Adams, R. A., Evans, S. A., and Jones, J. G., 1994, “Characterization of Leukocytes by Filtration of Diluted Blood,” Biorheology, 31(6), pp. 603–615. [PubMed]
Kikuchi, Y., 1995, “Effect of Leukocytes and Platelets on Blood Flow Through a Parallel Array of Microchannels: Micro- and Macroflow Relation and Rheological Measures of Leukocyte and Platelet Activities,” Microvas. Res., 50(2), pp. 288–300. [CrossRef]
Kikuchi, Y., Sato, K., and Mizuguchi, Y., 1994, “Modified Cell-Flow Microchannels in a Single-Crystal Silicon Substrate and Flow Behavior of Blood Cells,” Microvas. Res., 47(1), pp. 126–139. [CrossRef]
Schmid-Schonbein, G. W., 1987, “Capillary Plugging by Granulocytes and the No-Reflow Phenomenon in the Microcirculation,” Fed. Proc., 46(7), pp. 2397–2401. [PubMed]
Karnis, A., Goldsmith, H. L., and Mason, S. G., 1963, “Axial Migration of Particles in Poiseuille Flow,” Nature, 200(4902), pp. 159–160. [CrossRef]
Pries, A. R., Secomb, T. W., and Gaehtgens, P., 1996, “Biophysical Aspects of Blood Flow in the Microvasculature,” Cardiovas. Res., 32(4), pp. 654–667. [CrossRef]
Fåhræus, R., and Lindqvist, T., 1931, “The Viscosity of the Blood in Narrow Capillary Tubes,” 96(3), pp. 562–568.
Kim, S., Ong, P. K., Yalcin, O., Intaglietta, M., and Johnson, P. C., 2009, “The Cell-Free Layer in Microvascular Blood Flow,” Biorheology, 46(3), pp. 181–189. [CrossRef] [PubMed]
Freund, J. B., and Orescanin, M. M., 2011, “Cellular Flow in a Small Blood Vessel,” J. Fluid Mech., 671, pp. 466–490. [CrossRef]
Harris, A. G., and Skalak, T. C., 1993, “Effects of Leukocyte Activation on Capillary Hemodynamics in Skeletal Muscle,” Am. J. Physiol., 264(3 Pt 2), pp. H909–916. [PubMed]
Helmke, B. P., Bremner, S. N., Zweifach, B. W., Skalak, R., and Schmid-Schonbein, G. W., 1997, “Mechanisms for Increased Blood Flow Resistance Due to Leukocytes,” Am. J. Physiol., 273(6 Pt 2), pp. H2884–2890. [PubMed]
Helmke, B. P., Sugihara-Seki, M., Skalak, R., and Schmid-Schonbein, G. W., 1998, “A Mechanism for Erythrocyte-Mediated Elevation of Apparent Viscosity by Leukocytes In Vivo Without Adhesion to the Endothelium,” Biorheology, 35(6), pp. 437–448. [CrossRef] [PubMed]
Makino, A., Glogauer, M., Bokoch, G. M., Chien, S., and Schmid-Schonbein, G. W., 2005, “Control of Neutrophil Pseudopods by Fluid Shear: Role of Rho Family GTPases,” Am. J. Physiol. Cell Physiol., 288(4), pp. C863–871. [CrossRef] [PubMed]
Makino, A., Prossnitz, E. R., Bunemann, M., Wang, J. M., Yao, W., and Schmid-Schonbein, G. W., 2006, “G Protein-Coupled Receptors Serve as Mechanosensors for Fluid Shear Stress in Neutrophils,” Am. J. Physiol. Cell Physiol., 290(6), pp. C1633–1639. [CrossRef] [PubMed]
Rainger, G. E., Buckley, C. D., Simmons, D. L., and Nash, G. B., 1999, “Neutrophils Sense Flow-Generated Stress and Direct Their Migration Through AlphaVbeta3-Integrin,” Am. J. Physiol., 276(3 Pt 2), pp. H858–864. [PubMed]
Usami, S., Chen, H. H., Zhao, Y., Chien, S., and Skalak, R., 1993, “Design and Construction of a Linear Shear Stress Flow Chamber,” Ann. Biomed. Eng., 21(1), pp. 77–83. [CrossRef] [PubMed]
Carrigan, S. O., Weppler, A. L., Issekutz, A. C., and Stadnyk, A. W., 2005, “Neutrophil Differentiated HL-60 Cells Model Mac-1 (CD11b/CD18)-Independent Neutrophil Transepithelial Migration,” Immunology, 115(1), pp. 108–117. [CrossRef] [PubMed]
Hauert, A. B., Martinelli, S., Marone, C., and Niggli, V., 2002, “Differentiated HL-60 Cells Are a Valid Model System for the Analysis of Human Neutrophil Migration and Chemotaxis,” Int. J. Biochem. Cell Biol., 34(7), pp. 838–854. [CrossRef] [PubMed]
Sham, R. L., Packman, C. H., Abboud, C. N., and Lichtman, M. A., 1991, “Signal Transduction and the Regulation of Actin Conformation During Myeloid Maturation: Studies in HL60 Cells,” Blood, 77(2), pp. 363–370. [PubMed]
Zhelev, D. V., Alteraifi, A. M., and Chodniewicz, D., 2004, “Controlled Pseudopod Extension of Human Neutrophils Stimulated With Different Chemoattractants,” Biophys. J., 87(1), pp. 688–695. [CrossRef] [PubMed]
Neelamegham, S., Taylor, A. D., Burns, A. R., Smith, C. W., and Simon, S. I., 1998, “Hydrodynamic Shear Shows Distinct Roles for LFA-1 and Mac-1 in Neutrophil Adhesion to Intercellular Adhesion Molecule-1,” Blood, 92(5), pp. 1626–1638. [PubMed]
Russo, R. G., Liotta, L. A., Thorgeirsson, U., Brundage, R., and Schiffmann, E., 1981, “Polymorphonuclear Leukocyte Migration Through Human Amnion Membrane,” J. Cell Biol., 91(2 Pt 1), pp. 459–467. [CrossRef] [PubMed]
Rochon, Y. P., and Frojmovic, M. M., 1991, “Dynamics of Human Neutrophil Aggregation Evaluated by Flow Cytometry,” J. Leukocyte Biol., 50(5), pp. 434–443.
Fukuda, S., Yasu, T., Predescu, D. N., and Schmid-Schonbein, G. W., 2000, “Mechanisms for Regulation of Fluid Shear Stress Response in Circulating Leukocytes,” Circ. Res., 86(1), pp. E13–E18. [CrossRef] [PubMed]
Moazzam, F., DeLano, F. A., Zweifach, B. W., and Schmid-Schonbein, G. W., 1997, “The Leukocyte Response to Fluid Stress,” Proc. Natl. Acad. Sci. U. S. A., 94(10), pp. 5338–5343. [CrossRef] [PubMed]
Taylor, A. D., Neelamegham, S., Hellums, J. D., Smith, C. W., and Simon, S. I., 1996, “Molecular Dynamics of the Transition From L-Selectin- to Beta 2-Integrin-Dependent Neutrophil Adhesion Under Defined Hydrodynamic Shear,” Biophys. J., 71(6), pp. 3488–3500. [CrossRef] [PubMed]
Merrill, E. W., 1969, “Rheology of Blood,” Physiol. Rev., 49(4), pp. 863–888. [PubMed]
Besarab, A., Bolton, W. K., Browne, J. K., Egrie, J. C., Nissenson, A. R., Okamoto, D. M., Schwab, S. J., and Goodkin, D. A., 1998, “The Effects of Normal as Compared With Low Hematocrit Values in Patients With Cardiac Disease Who Are Receiving Hemodialysis and Epoetin,” N. Engl. J. Med., 339(9), pp. 584–590. [CrossRef] [PubMed]
Fang, W. C., Helm, R. E., Krieger, K. H., Rosengart, T. K., DuBois, W. J., Sason, C., Lesser, M. L., Isom, O. W., and Gold, J. P., 1997, “Impact of Minimum Hematocrit During Cardiopulmonary Bypass on Mortality in Patients Undergoing Coronary Artery Surgery,” Circulation, 96(9 Suppl), pp. II-194–199. [PubMed]
Miller, G. E., 2010, Fundamentals of Biomedical Transport Processes, Morgan & Claypool, San Rafael, CA.
Goldsmith, H. L., Lichtarge, O., Tessier-Lavigne, M., and Spain, S., 1981, “Some Model Experiments in Hemodynamics: VI. Two-Body Collisions Between Blood Cells,” Biorheology, 18(3–6), pp. 531–555. [PubMed]
Goldsmith, H. L., Quinn, T. A., Drury, G., Spanos, C., McIntosh, F. A., and Simon, S. I., 2001, “Dynamics of Neutrophil Aggregation in Couette Flow Revealed by Videomicroscopy: Effect of Shear Rate on Two-Body Collision Efficiency and Doublet Lifetime,” Biophys. J., 81(4), pp. 2020–2034. [CrossRef] [PubMed]
Sutton, D. W., and Schmid-Schonbein, G. W., 1992, “Elevation of Organ Resistance Due to Leukocyte Perfusion,” Am. J. Physiol., 262(6 Pt 2), pp. H1646–H1650. [PubMed]
Bagge, U., and Braide, M., 1982, “Leukocyte Plugging of Capillaries In Vivo,” White Blood Cells (Microcirculation Reviews), Vol. 1, U.Bagge, G. V. R.Born, and P.Gaehtgens, eds., Springer, Amsterdam, The Netherlands, pp. 89–98.
Klitzman, B., and Duling, B. R., 1979, “Microvascular Hematocrit and Red Cell Flow in Resting and Contracting Striated Muscle,” Am. J. Physiol., 237(4), pp. H481–H490. [PubMed]
Meisel, S. R., Shapiro, H., Radnay, J., Neuman, Y., Khaskia, A. R., Gruener, N., Pauzner, H., and David, D., 1998, “Increased Expression of Neutrophil and Monocyte Adhesion Molecules LFA-1 and Mac-1 and Their Ligand ICAM-1 and VLA-4 Throughout the Acute Phase of Myocardial Infarction: Possible Implications for Leukocyte Aggregation and Microvascular Plugging,” J. Am. Coll. Cardiol., 31(1), pp. 120–125. [CrossRef] [PubMed]
De Ville, M., Coquet, P., Brunet, P., and Boukherroub, R., 2012, “Simple and Low-Cost Fabrication of PDMS Microfluidic Round Channels by Surface-Wetting Parameters Optimization,” Microfluid. Nanofluid., 12(6), pp. 953–961. [CrossRef]
Faivre, M., Abkarian, M., Bickraj, K., and Stone, H. A., 2006, “Geometrical Focusing of Cells in a Microfluidic Device: An Approach to Separate Blood Plasma,” Biorheology, 43(2), pp. 147–159. [PubMed]
Shevkoplyas, S. S., Gifford, S. C., Yoshida, T., and Bitensky, M. W., 2003, “Prototype of an In Vitro Model of the Microcirculation,” Microvas. Res., 65(2), pp. 132–136. [CrossRef]
Lima, R., Wada, S., Tanaka, S., Takeda, M., Ishikawa, T., Tsubota, K., Imai, Y., and Yamaguchi, T., 2008, “In Vitro Blood Flow in a Rectangular PDMS Microchannel: Experimental Observations Using a Confocal Micro-System,” Biomed. Microdevices, 10(2), pp. 153–167. [CrossRef] [PubMed]
Goyal, M. R., 2013, Biofluid Dynamics of Human Body Systems, Apple Academic Press, Toronto, Canada.
Doring, Y., Drechsler, M., Soehnlein, O., and Weber, C., 2014, “Neutrophils in Atherosclerosis: From Mice to Man,” Arterioscler., Thromb., Vasc. Biol., 35, pp. 485–491. [CrossRef]
Hansen, P. R., 1995, “Role of Neutrophils in Myocardial Ischemia and Reperfusion,” Circulation, 91(6), pp. 1872–1885. [CrossRef] [PubMed]

Figures

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Fig. 1

Schematics of the experimental flow setups used in the present study. (a) A cone-plate viscometer was used to measure the suspension viscosities of neutrophil populations under a uniform shear field. Activated neutrophils with extended pseudopods, in contrast to inactivated neutrophils, were anticipated to increase viscosity due to enhanced numbers of stochastic cell–cell interactions resulting from increased cell tumbling. (b) For flow studies, cells were perfused through three different experimental chambers designed to test the impact of neutrophil pseudopod activity on rheological flow properties. These chambers included (1) an isopore membrane with pore sizes of 10 μm to simulate microcapillary flow; (2) a single-channel microchamber (w: 500 μm; h: 50 μm; and l: 20 mm) to simulate flow through the large microvessels; and (3) a network of twenty 20 × 50 μm microfluidics channels to simulate flow through noncapillary microvasculature. Pressure changes were recorded with a strain gauge pressure transducer and analyzed in labview signals express.

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Fig. 2

DHL60 pseudopod activity influences cell suspension viscosity. (a) and (b) Representative micrographs show the morphological changes associated with dHL60 stimulation. The dHL60 cells were either maintained in an inactivated state (a) or stimulated with 10 nM fMLP (b) for 5 min prior to fixation. Circularity was measured by manually tracing (white contour) the outlines of cells with imagej software. (c) Average circularities of inactivated (white bar) and activated (black bar) dHL60 neutrophil-like cells were assessed by tracing all cells present in 3–7 images per experimental treatment, with at least 50 cells analyzed. Bars are mean ± SEM for n = 3 independent experiments. (d) Viscosity measurements were obtained by cone-plate rheometry of inactivated (open circle) or activated (solid circle) dHL60 suspensions continuously recorded over a 5-min interval at 450 s−1. (e) Cone-plate rheometry was used to determine the average viscosities for suspensions of inactivated (white bar) or activated (black bar) dHL60 neutrophilic cells. Viscosity is expressed as mean ± SEM. *P < 0.05 compared to 0 nM fMLP stimulation (n = 3).

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Fig. 3

Extension of pseudopods by human neutrophils elevates suspension viscosity. (a) and (b) Representative micrographs show the morphological changes associated with human neutrophil stimulation. The human neutrophils were either maintained in an inactivated state (a) or stimulated with 10 nM fMLP (b) for 5 min prior to fixation. Circularity was measured by manually tracing (white contour) the outlines of cells with imagej software. (c) Average circularities of inactivated (white bar) and activated (black bar) neutrophils were assessed by tracing all cells present in 3–7 images per experimental treatment, with at least 50 cells analyzed. Bars are mean ± SEM for n = 3 independent experiments. (d) Viscosity measurements were obtained by cone-plate rheometry of inactivated (open circle) or activated (solid circle) human neutrophil suspensions continuously recorded over a 5-min interval at 450 s−1. (e) Cone-plate rheometry was used to determine the average viscosities for suspensions of inactivated (white bar) or activated (black bar) human neutrophils. Viscosity is expressed as mean ± SEM. (f) Average viscosity measurements were obtained for mixed populations of human neutrophils and erythrocytes exposed to a constant shear rate of 450 s−1. Erythrocytes were present at 10%, 20%, and 40% hematocrit, while human neutrophils were included as either inactivated (0 nM fMLP, white bars) or activated (10 nM fMLP, black bars) populations. Viscosity is expressed as mean ± SEM. n = 3. *P < 0.05 compared to 0 nM fMLP stimulation. n = 3.

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Fig. 4

RBCs exacerbate neutrophil activation-related elevations in perfusion pressure associated with flow of blood-like cell suspensions through micropore capillary mimics. (a) and (b) Plots of perfusion pressure versus time were generated to quantify the effects of cell activation (in the form of pseudopod extension) on the flow of neutrophil suspensions through micropore filter-based capillary mimics. Neutrophil populations that had either been maintained in an inactivated state (open circle) or activated with 10 nM fMLP for 5 min (solid circle) were perfused across micropore filters (10 μm pore diameters) in the absence (a) or presence (b) of 10% hematocrit. Average perfusion pressure determined from n = 3 experiments was expressed as mean ± SEM. (c) The percent increase in perfusion pressure due to cell activation was quantified by normalizing the perfusion pressures for activated neutrophil suspensions with that for inactivated cell suspensions. This normalization was carried out and plotted for suspensions lacking (open circle) or containing (solid circle) 10% hematocrit. Each data point is expressed as mean ± SEM. *P < 0.05 compared to inactivated controls. #P < 0.05 compared to populations without RBCs. n = 3. (d) Perfusion of purified suspensions of inactivated (right image) and activated (left image) neutrophils (stained with nuclear label 4', 6-diamidino-2-phenylindole (DAPI); black staining) through micropore filters (gray color) was associated with cell plugging.

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Fig. 5

In the absence of RBCs, neutrophil activation does not significantly impact flow resistance of microvascular mimics. (a) and (b) Plots of perfusion pressure versus time for microfluidics chambers comprised of either a single 500 × 50 μm channel (a) or a network of twenty 20 × 50 μm microchannels (b) were generated at various flow rates (0, 1, 1.5, 2, and 5 ml/hr) for purified neutrophil suspensions that had either been maintained in an inactivated state (solid-black line) or activated with 10 nM fMLP (dotted-gray line). (c) Based on these raw data, pressure versus flow curves were generated to determine effects of neutrophil activation on microchannel flow resistance. For each experiment, pressure values were obtained by averaging the measured pressure gradient over the last 2 min at each flow rate. From these curves, resistance values corresponding to single channel (inactivated—open triangles and activated—solid triangle) or microvascular network (inactivated—open circles and activated—solid circles) were found by computing the slope of each line. Points are mean ± SEM. n = 3.

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Fig. 6

In the presence of RBCs, neutrophil activation and pseudopod extension significantly elevate flow resistance of microvascular mimics. (a) and (b) Perfusion pressure versus time plots for microfluidics chambers comprised of either a single 500 × 50 μm channel (a) or a network of twenty 20 × 50 μm microchannels (b) were generated at various flow rates (0, 1, 1.5, 2, and 5 ml/hr) for purified neutrophil suspensions that had either been maintained in an inactivated state (solid-black line) or activated with 10 nM fMLP (dotted-gray line), fixed, and subsequently supplemented with 10% hematocrit. (c) Based on these raw data, pressure versus flow curves were generated to determine effects of neutrophil activation on microchannel flow resistance in the presence of hematocrit. For each experiment, pressure values were obtained by averaging the measured pressure gradient over the last 2 min at each flow rate. From these curves, resistance values corresponding to single channel (inactivated—open triangles and activated—solid triangle) or microvascular network (inactivated—open circles and activated—solid circles) were found by computing the slope of each line. Points are mean ± SEM. n = 3.

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