Nexinhib20 Inhibits Neutrophil Adhesion and β₂ Integrin Activation by Antagonizing Rac-1–Guanosine 5′-Triphosphate Interaction

Neutrophils are the most abundant leukocytes in humans and serve as the first responders to inflammation and infection (1). An intrinsic neutrophil defect leads to pathologies, such as leukocyte adhesion deficiency syndromes (2, 3). In contrast, abnormal activation of neutrophils is critically involved in most inflammatory diseases, such as ischemia-reperfusion (I/R) injury (4–6) and autoimmune diseases (7–10).

β₂ integrin activation is critical for neutrophil recruitment (1, 11–13). Therefore, regulating β₂ integrin signaling is a potential path to reduce inflammatory injury. Several GTPases are involved in β₂ integrin signaling, such as ras homolog (Rho) gene family GTPases (14–17) and Ras-related protein 1 (Rap1) GTPases (18–22). Several neutrophil exocytosis inhibitors (Nexinhibs) were identified by Förster resonance energy transfer–based screens that targeted the interaction of the small GTPase Rab27a and its effector JFC1 (23). The small GTPase Rab27a is an essential regulator of neutrophil exocytosis (24). Molecular docking analysis showed that Nexinhib20 might interact with an epitope formed by I10, K11, R90, D91, M93, Y122, S123, I181, R184, M185, and S188 of Rab27a (23). Nexinhibs did not interact with another small GTPase, Rab11 (23). Whether Nexinhibs affect the function of other GTPases, especially those involved in the integrin activation signaling pathway, is unknown. Identifying a Nexinhib that inhibits both integrin activation and exocytosis may serve as a dual-functional drug for treating inflammatory diseases.

Myocardial I/R injury exacerbates cardiomyocyte death after primary percutaneous coronary intervention in acute myocardial infarction. Neutrophils are recruited to cardiac tissue during myocardial I/R injury (5, 6) where they worsen injury (25, 26). They mediate cardiomyocyte death by causing vascular plugging, releasing degradative enzymes, and generating reactive oxygen species (ROS) (25). Neutrophil depletion in mice (27) and dogs (28) with myocardial I/R injury showed significant benefits in reducing infarct size. Inhibiting or deleting myeloperoxidase, which is mainly expressed by neutrophils (29), improves myocardial function after I/R injury (30, 31). The neutrophil recruitment cascade includes rolling, slow rolling, arrest, spreading, intravascular crawling, transendothelial migration, and migration to the site of inflammation (1, 32, 33); β₂ integrins play critical roles in most steps of the neutrophil recruitment cascade (1, 32, 33). Blocking neutrophil recruitment in mouse knockouts of β₂ integrin (CD18) or its ligand, ICAM-1, significantly reduced infarct size after myocardial I/R injury (34). Similar results were observed in β₂ integrin Ab blocking experiments in primate (35), pig (36), dog (37–39), rabbit (40, 41), and rat hearts (42, 43). Thus, targeting β₂ integrin activation might be a potential path to reduce myocardial I/R injury.

In this study, we tested whether Nexinhib20 could inhibit human neutrophil adhesion and β₂ integrin activation by targeting Ras-related C3 botulinum toxin substrate 1 (Rac-1) GTPase, and whether Nexinhib20 could limit neutrophil recruitment and decrease infarct size after mouse myocardial I/R injury.

Recombinant human P-selectin–Fc, ICAM-1–Fc, and IL-8 were purchased from R&D Systems. The Alexa Fluor (AF)488–conjugated and unconjugated conformation-specific mAbs mAb24 to human β₂-I–like or a domain (which reports the headpiece opening), unconjugated mouse anti-human CD18 mAb (blocking, clone TS1/18), AF594-conjugated rat anti-mouse CD31 mAb, allophycocyanin-conjugated rat anti-mouse CD115 mAb, PE-conjugated rat anti-mouse Ly6G mAb, AF700-conjugated rat anti-mouse CD45 mAb, unconjugated mouse IgG1 isotype control, allophycocyanin-conjugated rat anti-mouse IgG1 secondary mAb, and a Zombie Yellow fixable viability kit were purchased from BioLegend. The KIM127 mAb to the human β₂-I-EGF2 domain, which reports the ectodomain extension, was purified at the Lymphocyte Culture Center at the University of Virginia from hybridoma supernatant (American Type Culture Collection). KIM127 was directly labeled by DyLight 550 (DL550) using DyLight Ab labeling kits from Thermo Fisher Scientific. Nexinhib20 was purchased from Tocris Bioscience. Casein blocking buffer, Fluo-4 AM, and Pierce protease inhibitor mini tablets were purchased from Thermo Fisher Scientific. Ghost Dye Blue 516 was purchased from Tonbo Biosciences. Polymorphprep was purchased from Accurate Chemical. RPMI 1640 without phenol red and PBS were purchased from Life Technologies. Human serum albumin (HSA) and FBS were purchased from Gemini Bio Products. Formalin and nonfat milk were purchased from Fisher Scientific. A Rac-1 activation assay biochem kit, which contains p21-activated kinase 1–p21 binding domain (PAK-PBD) protein beads, purified His-tagged Rac-1 protein, GTPγS (nonhydrolyzable GTP analog), GDP, and several buffers, were purchased from Cytoskeleton. A bulk custom order of purified His-tagged Rac-1 protein was purchased from Quintarabio. fMLF, triphenyl tetrazolium chloride (TTC), PMA, Polybrene, paraformaldehyde (PFA), and DMSO were purchased from Sigma-Aldrich. Laemmli sample buffer (2×) and Mini-PROTEAN TGX precast gels were purchased from Bio-Rad. Mouse monoclonal anti–Rac-1 Ab was purchased from BD Biosciences. HRP-conjugated horse anti-mouse Ab was purchased from Cell Signaling Technology. Trappsol (2-hydroxypropyl-β-cyclodextrin) was purchased from Cyclodextrins Technology Development. ECL Ultra was purchased from Lumigen. Penicillin, streptomycin, and amphotericin B solutions were purchased from HyClone. The total ROS assay kit was purchased from Invitrogen.

Heparinized whole-blood samples were obtained from healthy human donors after informed consent, as approved by the Institutional Review Board of the La Jolla Institute for Immunology and UConn Health in accordance with the Declaration of Helsinki. Informed consent was obtained from all donors. Neutrophils were isolated using a Polymorphprep (a mixture of sodium metrizoate and Dextran 500) density gradient. Briefly, human blood was applied to Polymorphprep and centrifuged at 500 × g for 35 min at 20°C, resulting in neutrophils concentrated in a layer between PBMCs and erythrocytes. After washing with PBS twice, the neutrophils (>95% purity by flow cytometry, no visible activation by microscopy) were resuspended in RPMI 1640 without phenol red plus 2% HSA and used within 4 h. Neutrophils were incubated with FcR blocking reagents for 10 min at room temperature (RT) before all the experiments.

The assembly of the microfluidic devices used in this study and the coating of coverslips with recombinant human P-selectin–Fc and ICAM-1–Fc with or without IL-8 have been described previously (44). Briefly, coverslips were coated with P-selectin–Fc (2 μg/ml) and ICAM-1–Fc (10 μg/ml) without or with IL-8 (10 μg/ml) for 2 h and then blocked for 1 h with casein (1%) at RT. After coating, coverslips were sealed with polydimethylsiloxane chips by magnetic clamps to create flow chamber channels ∼29 μm high and ∼300 μm wide. By modulating the pressure between the inlet well and the outlet reservoir, 6 dyn/cm² wall shear stress was applied in all experiments.

To study the rolling and arrest of neutrophils, isolated human neutrophils (5 × 10⁶ cells/ml) were perfused in the microfluidic device over a substrate of recombinant human P-selectin–Fc and recombinant human ICAM-1–Fc with or without IL-8 under a shear stress of 6 dyn/cm². Neutrophils were incubated with Nexinhib20 (10 µM) or vehicle (DMSO) for 1 h at RT before being perfused into the microfluidic devices. Time-lapse images (one frame per second) were taken by an IX71 inverted research microscope (Olympus America) with a ×40 numerical aperture 0.9 air objective during the perfusion to quantify rolling velocity. The quantification was done using the “Manual tracking” plugin in FIJI-ImageJ v2.0. Cell tracks (Fig. 1A) and rolling velocity were obtained (Fig. 1B, 1C). After perfusion with neutrophils for 10 min, the microfluidic device was washed with RPMI 1640 without phenol red plus 2% HSA for 5 min. Then, the arrested neutrophils were counted in nine fields of view per group (Fig. 1D).

C57BL/6J wild-type mice were originally obtained from The Jackson Laboratory (00664). LysM–enhanced GFP (EGFP) or Lyz2-EGFP mice (45) were originally obtained from Albert Einstein College of Medicine through a material transfer agreement. Mice were fed a standard rodent chow diet and were housed in microisolator cages in a pathogen-free facility in the Center for Comparative Medicine at UConn Health. All experiments followed the UConn Health Institutional Animal Care and Use Committee guidelines, and approval for the use of rodents was obtained from the UConn Health Institutional Animal Care and Use Committee according to criteria outlined in the Guide for the Care and Use of Laboratory Animals from the National Institutes of Health. Both male and female mice aged from 12 to 16 wk were used in the experiments.

Mice were subjected to 35 min of myocardial ischemia and 1 h (for multiphoton microscopy and flow cytometry) or 22–26 h (for TTC/Phthalo Blue staining) of reperfusion. The reason we use two time points to harvest is because neutrophil recruitment happens 1 h after the reperfusion (5), and the infarct size can be significantly quantified by TTC/Phthalo Blue staining after ∼24 h reperfusion (46). Briefly, anesthesia was induced with an i.p. injection of ketamine hydrochloride (125 mg/kg) and xylazine (12.5 mg/kg). Mice were intubated with a 24G × ¾-inch Surflo i.v. catheter and ventilated using a MiniVent 845 (Harvard Apparatus).

Surgeries were performed under an SMZ168 stereo zoom microscope (Motic). Ischemia was achieved by ligating the left anterior descending coronary artery using a 6-0 silk suture with a section of PE-10 tubing placed over this artery, 1 mm from the tip of the normally positioned left atrium. One critical problem in drug administration is water solubility, which greatly affects drug absorption and bioavailability (47). In our study, we used Trappsol as a cosolvent for in vivo administration to increase Nexinhib20 solubility. In clinics, the primary percutaneous coronary intervention is intended to be performed <90 min (within 60 min is preferable) after the patient arrives. To mimic a prevention treatment of reperfusion injury before the primary percutaneous coronary intervention, which is feasible in the clinics, Nexinhib20 (100 mM, 10 µl in DMSO mixed with 190 µl of 10% Trappsol per mouse) or vehicle control was administered i.p. 30 min before the reperfusion. After occlusion for 35 min, reperfusion was initiated by releasing the ligature and removing the PE-10 tubing. The chest wall was closed, the animal extubated, and body temperature was maintained by use of a 37°C warm pad. Hearts were harvested 1 or 22–26 h later. The loosened suture was left in place and then retied for the purpose of evaluating the ischemic area. Sham control and no-drug-administered control were performed as well.

Isolated human neutrophils (2 × 10⁶ cells/ml in RPMI 1640 without phenol red plus 2% HSA) were incubated with Nexinhib20 (10 µM) or vehicle (DMSO) for 1 h at RT before being assayed. To monitor the dynamics of β₂ integrin activation, 400 µl of 2.5 × 10⁵ cells/ml neutrophils was assessed by an LSR II analyzer (BD Biosciences, San Jose, CA) for 10 s. After adding 0.5 µg/ml of AF488-conjugated mAb24 and DL550-conjugated KIM127 (final concentration), cells were put back into the analyzer for another 5 min. Then, after adding 1 µg/ml IL-8, cells were put back into the analyzer for another 10 min. The curves showing the dynamics of integrin activation (Fig. 2A, 2B) were generated by FlowJo software (version 10.6). The Ab specificities were validated in our previous study using β₂ integrin knockout cells and β₂ integrin activation-deficient talin-1 knockout cells (48). Compensations were performed before all experiments.

To quantify the percentage of mAb24 and KIM127 epitopes and assess inhibition of β₂ integrin exocytosis, pan-CD18 mAb24 TS1/18, which has the same isotype (mouse IgG1) as mAb24 and KIM127, was used. Isolated human neutrophils (5 × 10⁵ cells/ml in RPMI 1640 without phenol red plus 2% HSA) were incubated with Nexinhib20 (10 µM) or vehicle (DMSO) for 1 h at RT before being assayed. Neutrophils were mixed with unconjugated mAb24 (1 µg/ml), KIM127 (1 µg/ml), TS1/8 (1 µg/ml), or mouse IgG1 isotype control (1 µg/ml) and incubated with 1 µg/ml IL-8 at RT for 10 min. After incubation, neutrophils were fixed by 1% PFA at 4°C for 10 min. After two washes with PBS, cells were incubated with allophycocyanin-conjugated rat anti-mouse IgG1 secondary mAb (1 µg/ml) at RT for 10 min. After two washes with PBS, cell fluorescence was assessed with an LSR II (BD Biosciences, San Jose, CA). The quantifications of mAb24, KIM127, TS1/18, and isotype mean fluorescence intensities (MFIs) (Fig. 2C–E) were analyzed by FlowJo software (version 10.6) and obtained from six replicates. MFI of isotype controls was subtracted as background signal. Because mAb24, KIM127, and TS1/18 are all IgG1 isotypes and we used the same secondary Ab, the percentage of high-affinity and extended β₂ integrins (Fig. 2F, 2G) can be calculated by dividing the MFI of mAb24 and KIM127 by the MFI of TS1/18.

To monitor the dynamics of intracellular calcium (Ca²⁺) flux, neutrophils (2 × 10⁶ cells/ml in RPMI 1640 without phenol red plus 2% HSA) were incubated with Fluo-4 (4 µg/ml) for 1 h at RT. After washes, neutrophils were resuspended in RPMI 1640 without phenol red plus 2% HSA and assessed by an LSR II analyzer (BD Biosciences, San Jose, CA). One minute after analyzing, 1 µg/ml IL-8 was added to the cells. Cells were put back into the analyzer for another 9 min. The curves showing the dynamics of intracellular Ca²⁺ flux (Fig. 2H) were generated by FlowJo software (version 10.6). The quantification of Fluo-4 MFI (Fig. 2I) was analyzed by FlowJo software (version 10.6) and obtained from three individual experiments.

To assess the viability of neutrophils (Supplemental Fig. 1B), neutrophils (2 × 10⁶ cells/ml in RPMI 1640 without phenol red plus 2% HSA) were incubated with different concentrations (0, 10, 20, 50, and 100 µM) of Nexinhib20 at RT for 1 h. After washes, neutrophils were incubated with Ghost Dye Blue 516 at RT for 15 min. After washes, cell fluorescence was assessed with an LSR II (BD Biosciences, San Jose, CA) and analyzed with FlowJo software (version 10.6).

To assess the neutrophil recruitment in myocardial I/R injury, LysM-EGFP mice underwent 35 min of ischemia and 1 h of reperfusion. To mimic a prevention treatment of reperfusion injury before the primary percutaneous coronary intervention, which is feasible in the clinics, Nexinhib20 (100 mM, 10 µl per mouse) or vehicle control was administered i.p. 30 min prior to the reperfusion. After the 1-h reperfusion, the mouse heart was harvested and perfused with ice-cold PBS to remove residual blood and unbound leukocytes, transferred into an ice-cold gentleMACS C tube, cut into ∼1-mm³ pieces, suspended with 5 ml of PBS plus 2% FBS, 2 mM EDTA, and 0.08 µg/ml allophycocyanin-conjugated anti-CD115 mAb, and homogenized five times by the m_Heart_01 program of the gentleMACS dissociator (Miltenyi Biotec). The cell suspension was filtered by 70-µm nylon mesh strainer (Fisher Scientific), centrifuged at 500 × g, 4°C for 5 min, resuspended in 200 µl of 1:300 diluted Zombie Yellow fixable viability dye, and incubated on ice for 15 min. After centrifuging at 500 × g, 4°C for 5 min, cells were resuspended in 200 µl of ice-cold PBS containing 1.25 µg/ml AF700-conjugated anti-CD45 mAb and 1 µg/ml PE-conjugated anti-Ly6G mAb and incubated on ice for 10 min. After being fixed with 1% PFA and washes with ice-cold PBS, cell fluorescence was assessed with an LSR II (BD Biosciences, San Jose, CA) and analyzed with FlowJo software (version 10.6).

Peripheral blood of the above mice was also collected. One hundred microliters was mixed with 200 µl of 1:300 diluted Zombie Yellow fixable viability dye and incubated on ice for 15 min. After centrifuging at 500 × g, 4°C for 5 min, cells were resuspended in 200 µl of ice-cold PBS containing 1.25 µg/ml AF700-conjugated anti-CD45 mAb, 1 µg/ml PE-conjugated anti-Ly6G mAb, and 2 µg/ml allophycocyanin-conjugated anti-CD115 mAb, and incubated on ice for 10 min. After being fixed with 1% PFA, RBCs were lysed with deionized water for 30 s (stopped by adding 10× PBS). Leukocyte fluorescence was assessed with an LSR II (BD Biosciences, San Jose, CA) and analyzed with FlowJo software (version 10.6).

ROS production of isolated human neutrophils was quantified by using a total ROS assay kit from Invitrogen. A black, clear-bottom, nontreated 96-well plate was used in this assay. Before the assay, the 96-well plate was coated with 10 µg/ml human ICAM-1–Fc at RT for 2 h and washed twice with PBS. Isolated human neutrophils (2 × 10⁶ cells/ml) were incubated with Nexinhib20 (10 µM) or vehicle (DMSO) for 1 h at RT before being assayed. After centrifuging at 300 × g, RT for 2 min, cells were resuspended at 10⁶ cells/ml in ROS assay stain solution from the kit and incubated with 2 µg/ml mouse anti-human CD18 blocking mAb (TS1/18 to block neutrophil adhesion) (49) or isotype control at RT for 10 min. The 100 µl/well neutrophils (three replicates per group) were seeded into the ICAM-1–coated 96-well plate. The background ROS before stimulation were measured by a Cytation 1 cell imaging multi-mode reader (filter set: green, excitation 485/20 nm, emission 528/20 nm, BioTek, Santa Clara, CA). Then, 100 nM PMA was added to each well, and ROS production was measured by a Cytation 1 cell imaging multi-mode reader every 5 min.

The HL60 cells and CXCR2-expressing HL60 (HL60-CXCR2) cells (50) were gifts from Dr. Orion D. Weiner at the University of California San Francisco and Dr. Ann Richmond at the Vanderbilt University School of Medicine, respectively. HL60-CXCR2 cells were selected with G418 (0.5 µg/ml) to maintain CXCR2 expression. Cells were maintained in culture medium (RPMI 1640, 10% FBS, 100 μg/ml penicillin, 100 μg/ml streptomycin, and 250 ng/ml amphotericin B) at 37°C and 5% CO₂. In most experiments, cells were differentiated with 1.3% DMSO for 7 d before assays. Cells were checked monthly for mycoplasma infection using the e-Myco plus Mycoplasma PCR detection kit.

Differentiated HL60 or HL60-CXCR2 cells or isolated human neutrophils (2 × 10⁶ cells/ml in RPMI 1640 without phenol red plus 2% HSA) were incubated with Nexinhib20 (10 µM) or vehicle (DMSO) for 1 h at RT. After washes, cells were resuspended in RPMI 1640 without phenol red (10⁷ cells/ml) and incubated with or without stimulators (100 nM fMLF for HL60, 1 µg/ml IL-8 for HL60-CXCR2) at RT for 1 min. Cells were lysed by 1:1 addition of 2× Triton X-100 lysis buffer (final concentration: 1% Triton X-100, 50 mM HEPES [pH 7.0], 150 mM NaCl, 10% glycerol, 1.5 mM MgCl₂, 1 mM EGTA, 1 mM sodium orthovanadate, 1 mM PMSF, plus protease inhibitor mixture/one Pierce protease inhibitor mini tablet per 5 ml 2× buffer) on ice for 5 min. After centrifuging at 16,000 × g, 4°C for 8 min, supernatants were saved as protein samples.

For the in vitro Rac-1/GTP binding competition assays, purified His-tagged Rac-1 (0.08 µg/ml in 1× Triton X-100 lysis buffer) were mixed with loading buffer (from the Rac-1 Activation Assay Biochem Kit, 1:10) and Nexinhib20 (1, 3, 10, 30, 100, 300, and 1000 µM) or vehicle (DMSO). Then, samples were incubated with GTPγS (0.4 µM) or GDP (0.8 mM) at RT for 15 min. The reaction was stopped by transferring samples to 4°C and adding the stop buffer (from the Rac-1 Activation Assay Biochem Kit, 1:10).

The Rac-1–GTP pull-down was performed using Rac-1 Activation Assay Biochem Kit following the manufacturer’s instructions. Briefly, protein samples were immediately incubated with PAK-PBD beads (10 µl per 1 ml of sample) for 1 h at 4°C. Then, beads were pelleted by centrifugation at 5000 × g, 4°C for 8 min. After removal of most of the supernatant, beads were washed twice with 500 µl of washing buffer from the kit. Beads were resuspended with 2× Laemmli sample buffer and boiled for 2 min.

Protein samples (before and after the Rac-1–GTP pull-down) were separated using SDS-PAGE and transferred onto nitrocellulose membranes. Membranes were blocked for ∼30 min in Tris-buffered saline with 0.1% Tween 20 (TBST) plus 5% nonfat milk. After blocking, membranes were incubated overnight with mouse monoclonal anti-Rac-1 Ab diluted 1:2000 in TBST at 4°C and HRP-conjugated horse anti-mouse Ab diluted 1:5000 in TBST plus 5% non-fat milk at RT for 1 h. ImageQuant LAS 4000 (GE Healthcare) was used to image membranes after adding ECL Ultra (Fig. 3).

Mice underwent 35 min of ischemia and 1 h of reperfusion. To mimic a prevention treatment of reperfusion injury before the primary percutaneous coronary intervention, which is feasible in the clinics, Nexinhib20 (100 mM, 10 µl per mouse) or vehicle control was administered i.p. 30 min prior to the reperfusion. After the reperfusion, the mouse heart was harvested and perfused with PBS to remove residual blood and unbound leukocytes and was perfused with anti–CD31-AF594 mAb (10 µg/ml, 250 µl per heart) to label the coronary artery sequentially. The explanted heart was immersed in PBS and imaged by a multiphoton microscope immediately. The Bruker’s upright multiphoton microscope (no. 4269) was equipped with a Mai Tai high-performance Ti:sapphire femtosecond pulsed laser (tuning range 690–1020 nm, set to 780 nm excitation in this assay) and a ×20 numerical aperture 0.95 water immersion objective. The bandpass filters in front of the corresponding four different photomultiplier tube detectors are 660/40, 595/50, 525/50, and 460/50 nm. The 525/50 nm channel and 595/50 nm channel were used for EGFP and AF594 imaging, respectively. Three-dimensional z stack series (5-µm interval, 10–20 stacks) images of the coronary artery were acquired (Fig. 4A). The MFI of EGFP within the coronary artery was quantified by FIJI-ImageJ v2.0.

To assess the ischemic area at risk after 22–26 h of reperfusion, hearts were excised, infused with PBS and freshly prepared 10% Phthalo Blue (PBS with 0.75% Tween 20) through the aorta and coronary arteries in a retrograde fashion, frozen at −20°C for 10 min, and sliced into five to six 1-mm cross-sections with the aid of a prefreeze acrylic matrix (Zivic Labs). The heart sections were incubated with freshly prepared 1% TTC solution (Sigma-Aldrich) at 37°C for 10 min and fixed with formalin. Viable myocardium stained red, and infarcted tissue appeared white. Images (Fig. 5A) were acquired by an MU130 color complementary metal oxide semiconductor (CMOS) camera (AmScope) equipped on an SMZ168 stereo zoom microscope (Motic). The infarct area (white), the area at risk (red and white), and the total left ventricle area from each section were measured using ZEN v3.1 (Zeiss). Ratios of infarct area/area at risk (Fig. 5B) and of area at risk/left ventricle (Fig. 5C) were calculated and expressed as percentages.

To quantify left ventricle function, we performed echocardiograms on mice before and 7 d after myocardial I/R injury. Mice were anesthetized with 2% isoflurane intranasally and placed on a heating pad. Chest hair was removed using an electric shaver and animals were fixated on their backs. Echocardiography loops were recorded in B and two dimensional–targeted M modes in longitudinal and short-axis views on a Vevo 3100 high-resolution imaging system equipped with an MX550D transducer (VisualSonics, Toronto, ON, Canada). Heart rate was monitored during the procedure. Systole and diastole were defined based on concomitant electrocardiography recordings. The end-systolic time point for left ventricle diameter measurement was defined as the maximum ventricle contraction just before the complete closure of the aortic valve. End-diastole was defined as the maximum left ventricle dilation and filling just before mitral valve closing (when visible) and aortic valve opening. The left ventricular ejection fraction was determined by left ventricle tracing relating the end-systolic left ventricle area as the minimal left ventricle cross-sectional area to the end-diastolic left ventricle area as the maximum left ventricle cross-sectional area in long-axis views. Fractional shortening was assessed by using Vevo LAB software (Visualsonics).

Pharmacokinetics was performed through a service provided by the Shanghai Institute of Materia Medica. Nexinhib20 (160 mM, 5 µl in DMSO mixed with 95 µl of 10% Trappsol per mouse) was administered i.p. to three 18 to 19 g male mice. Blood samples (20 µl) were collected at 3 min, 15 min, 45 min, 2 h, 4 h, 8 h, and 24 h through femoral vein phlebotomy. Two hundred microliters of methanol/acetonitrile (1:1, v/v) with internal standard was added to 20 μl of plasma and vortexed thoroughly. After centrifuging at 11,000 rpm, RT for 5 min, 20 μl of the supernatant was collected and mixed with 20 μl of water for analysis. Samples were analyzed by a TQ-S triple quadrupole mass spectrometer (Waters, Milford, MA). An ACQUITY UPLC BEH C18 column (1.7 μm, 2.0 × 50 mm, Waters) was used for the analysis. Gradient elution was applied consisting of 5 mM aluminum ammonium sulfate dodecahydrate containing 0.1% formic acid and methanol/acetonitrile (1:9, v/v) containing 0.1% formic acid.

Statistical analysis was performed using Prism software (version 8.30, GraphPad Software). Data analysis was performed using a Student’s t test, one-way ANOVA followed by a Tukey’s multiple comparison test, or two-way ANOVA followed by a Tukey’s multiple comparison test, as indicated in the figure legends. A p value <0.05 was considered significant.

Nexinhib20 inhibits exocytosis without inducing apoptosis or cell death (23). In this study, to further analyze whether Nexinhib20 could be toxic to neutrophils, we tested the viability of neutrophils after Nexinhib20 treatment using flow cytometry. We showed that neutrophil viability remained close to 100% even when incubated with 100 µM Nexinhib20 for 1 h at RT (Supplemental Fig. 1A). This is consistent with the previous study that Nexinhib20 did not induce a significant increase in cell death after 1 and 4 h of incubation compared with DMSO vehicle controls (23). We incubated neutrophils with 10 µM Nexinhib20 for 1 h at RT in most of our cellular experiments unless stated otherwise.

To assess the impact of Nexinhib20 on neutrophil adhesion, we performed microfluidic assays as described previously (44, 51). As expected, neutrophils rolled on the substrate of human P-selectin and ICAM-1 under a shear stress of 6 dyn/cm² (Fig. 1A, upper left), which is a typical shear stress in postcapillary venules that commonly show neutrophil recruitment during inflammation (52). Upon addition of IL-8 to the substrate, neutrophils stopped rolling (arrest) and reduced the 100-s rolling distance from ∼200 to ∼40 µm (Fig. 1a, bottom left). When neutrophils were incubated with Nexinhib20 before the perfusion, they failed to arrest (Fig. 1a, bottom right). After quantifying the rolling velocity of these neutrophils (Fig. 1B, 1C), we found that IL-8 stimulation did not slow down the rolling velocity of Nexinhib20-treated neutrophils. After 10 min of rolling and 5 min of washing, we quantified the arrested neutrophils and found that Nexinhib20 significantly decreased the number of arrested neutrophils from ∼200 cells per field of view to ∼20 cells per field of view (∼90%, (Fig. 1D). Thus, Nexinhib20 inhibited adhesion of human neutrophils to P-selectin and ICAM-1 in a microfluidic model of physiological flow conditions.

Because β₂ integrins are critical for neutrophil adhesion (33, 53, 54), Nexinhib20 was tested for its potential to inhibit β₂ integrin expression and activation on neutrophils. Nexinhib20 was developed as a neutrophil exocytosis inhibitor that decreases exocytosis of the integrin α_Mβ₂ (Mac-1, CD11b/CD18) α subunit CD11b (23). It has been shown that after 30 min of stimulation with GM-CSF and fMLF, CD11b on human neutrophils was upregulated to ∼2-fold compared with unstimulated cells. Pretreatment with Nexinhib20 diminished this CD11b upregulation. In this study, we assessed the effect of Nexinhib20 on total β₂ subunit (CD18) surface expression (Fig. 2E). We found that total CD18 expression was upregulated by ∼40% after 10 min with 1 µg/ml IL-8 stimulation. As expected, this β₂ integrin exocytosis was inhibited significantly by Nexinhib20 treatment (Fig. 2E). Second, we tested β₂ integrin activation, which has two major conformational changes of the β₂ integrin extracellular domain-headpiece opening to acquire high-affinity (H⁺) binding to ligands and extension (E⁺) that allow binding ligands in trans (13, 33). H⁺ and E⁺ β₂ integrins can be monitored by using the conformation-specific Abs mAb24 (55–57) and KIM127 (58, 59), respectively. Please note that both lymphocyte function-associated Ag 1 (LFA-1, CD11a/CD18, α_Lβ₂) and Mac-1 were detected. Time-resolved flow cytometry showed that IL-8 induced dramatic increases of both mAb24 (Fig. 2A) and KIM127 (Fig. 2B) binding, and that Nexinhib20 treatment inhibited these effects. Because the time-resolved flow cytometry cannot remove free mAb in the cell suspension that generates background noise, we also used standard flow cytometry with fixation and washing to remove free mAb and get more accurate quantification (Fig. 2C–G). Isotype control (mouse IgG1) was used to determine background noise in the standard flow cytometry assay. After 10 min of 1 µg/ml IL-8 stimulation, we found that mAb24 staining increased to ∼10-fold, and KIM127 staining increased to ∼2-fold. Nexinhib20 inhibited the IL-8–induced elevation of mAb24 (Fig. 2C) by ∼75% and KIM127 (Fig. 2D) by ∼20%. Because TS1/18 (Fig. 2E) is the same isotype (mouse IgG1) as mAb24 and KIM127 and the same secondary Ab was used, the percentage of high-affinity and extended β₂ integrins can be calculated by dividing the MFI of mAb24 and KIM127, respectively, by the MFI of TS1/18. We found that Nexinhib20 reduced the percentage of high-affinity (mAb24, (Fig. 2F) but not extended β₂ integrins (KIM127, (Fig. 2G), suggesting that Nexinihib20 inhibits β₂ integrin high-affinity activation but not extension activation. These results demonstrated that Nexinhib20 significantly limited both the exocytosis and activation of β₂ integrins on human neutrophils, which are critical events for neutrophil adhesion.

Intracellular Ca²⁺ transients are involved in the chemokine-triggered integrin inside-out activation signaling pathway (60). Ca²⁺ and diacylglycerol activate Rap1 GTPases, which are critical for integrin inside-out activation (18, 22), through calcium and diacylglycerol-regulated guanine nucleotide exchange factors (22). The short inside-out Ca²⁺ signal can be triggered by IL-8 through its receptor CXCR2 (61, 62). The disassociated Gβγ activates Ras-related C3 botulinum toxin substrate 1 (Rac-1) and phospholipase C β (PLCβ) sequentially and induces intracellular Ca²⁺ flux (16). Using the intracellular Ca²⁺ dye Fluo-4 and time-resolved flow cytometry, we evaluated transient elevation of Fluo-4 fluorescence in neutrophils upon IL-8 stimulation (Fig. 2H, red trace). Nexinhib20 treatment potently blocked the IL-8–induced Ca²⁺ signal (Fig. 2H, blue trace, 2I).

In the integrin inside-out activation signaling pathway, Rac-1 is an upstream signaling molecule of intracellular Ca²⁺ flux. In this pathway, the activation of G protein–coupled receptors dissociates G protein to Gα and Gβγ subunits (63). Gβγ activates Rac-1 through P-Rex1 and Vav1 (64), then activates PLCβ2 and PLCβ3 and induces intracellular Ca²⁺ flux and downstream signaling molecules mentioned above to activate β₂ integrins (16). Rac-1 knockout neutrophils showed defects in inside-out signaling-triggered adhesion (16). Thus, we tested whether Nexinhib20 can inhibit Rac-1. The neutrophil-like cell line HL60 and HL60-CXCR2 cells were used in the Rac-1 activity assays. Using the PAK-PBD bead pull-down assay, which enriches for the active GTP form of Rac-1 (Rac-1–GTP), followed by anti–Rac-1 Western blots, we found that Nexinhib20 significantly inhibited the IL-8–induced (Fig. 3A, 3B) or fMLF-induced (Fig. 3C, 3D) Rac-1 activation in HL60-CXCR2 or HL60 cells, respectively. Quantification showed that IL-8 (Fig. 3B) and fMLF (Fig. 3D) stimulation increased the amount of Rac-1–GTP by ∼80% and ∼50%, respectively, and Nexinhib20 treatment eliminated these increases (Fig. 3B, 3D). To further confirm our findings in human neutrophils, we performed the Rac-1 pull-down assay using Nexinhib20 or vehicle-treated human neutrophils (Fig. 3E, 3F). Similar to HL60 data, we showed that IL-8 stimulation increases the amount of Rac-1–GTP by ∼60%. Nexinhib20 treatment eliminated these increases. Thus, Nexinhib20 inhibited Rac-1 activity in cells, which was consistent with the inhibition of intracellular Ca²⁺ flux and integrin activation in neutrophils.

Nexinhib20 was discovered by a screen for inhibitors of Ras-related protein Rab27a–synaptotagmin-like 1 (SYTL1 or JFC1) interaction and was expected to directly bind Rab27a by molecular docking analysis (23). Rac-1 was reported to interact with JFC1 as well (65). Thus, we hypothesized that Nexinhib20 directly binds Rac-1 or competes for Rac-1–GTP binding. To test this, we performed in vitro binding assays using purified His-tagged Rac-1 protein (Fig. 3G, 3H). Incubating with the nonhydrolyzable GTP analog GTPγS produced active Rac-1–GTP that was enriched by PAK-PBD beads (Fig. 3G, second column, vehicle control). His-tagged Rac-1 protein incubated with GDP was used as a negative control (Fig. 3G, first column). In the presence of Nexinhib20, the binding of His-tagged Rac-1 and GTPγS was significantly inhibited in a dose-dependent manner (Fig. 3G). After calculating the fitting curve of the inhibition percentage (Fig. 3H), we found that the IC₅₀ of Nexinhib20 to Rac-1–GTP binding was ∼29.3 µM. These data suggested that Nexinhib20 could antagonize the Rac-1–GTP interaction and may directly bind to Rac-1. This direct inhibition indicated that the upstream P-Rex1 and Vav1 for Rac-1 activation might not be relevant in the Nexinhib20 inhibition of neutrophil integrin activation.

Nexinhib20 was shown to inhibit neutrophil extracellular superoxide anion production by ∼50% (23). In this study, we tested the effect of Nexinhib20 on neutrophil total ROS production and its adhesion dependency (Supplemental Fig. 1B, 1C). After 50 min of 100 nM PMA stimulation, Nexinhib20-treated neutrophils showed significantly reduced total ROS production compared with vehicle controls (Supplemental Fig. 1B). Interestingly, this inhibition is not adhesion-dependent because CD18 blockade, which was shown to reduce adhesion (51) and spreading (49) of neutrophils, did not inhibit ROS production in both vehicle control neutrophils (Supplemental Fig. 1C) and Nexinhib20-treated neutrophils (data not shown).

Neutrophils are critically involved in myocardial I/R injury (25, 26). Intravital imaging has shown that neutrophils are recruited abundantly to the coronary artery 60 min after reperfusion (5). To test whether Nexinhib20 inhibits neutrophil recruitment in myocardial I/R injury in vivo, we performed multiphoton imaging on explanted hearts after 35 min of ischemia and 60 min of reperfusion (Fig. 4). To test this possibility, LysM-EGFP mice were used in our study and were also used to monitor neutrophil recruitment in hearts (5, 66, 67). We observed profound accumulation of LysM-EGFP⁺ cells in the coronary artery in vehicle controls (Fig. 4A, left panel). To mimic a prevention treatment of reperfusion injury before the primary percutaneous coronary intervention, which is feasible in clinics, Nexinhib20 was administered 30 min before the reperfusion. Nexinhib20 treatment significantly reduced the number of LysM-EGFP⁺ leukocytes (Fig. 4A, right panel). Quantification of EGFP fluorescence in coronary arteries confirmed that Nexinhib20 significantly limited LysM-EGFP⁺ leukocyte recruitment to the coronary artery during reperfusion (Fig. 4B).

Although macrophages and monocytes may also be highlighted by EGFP in LysM-EGFP mice, it has been shown that ∼90% of EGFP⁺ cells in I/R heart are Ly6G⁺ neutrophils (66). To further explore the components of LysM-EGFP⁺ leukocytes in our experimental setting, we used flow cytometry to quantify the percentage of Ly6G⁺ neutrophils in the I/R heart and blood circulation in mice treated with Nexinhib20 or not (Fig. 4C–E). Consistent with the multiphoton imaging data (Fig. 4A, 4B), there were fewer LysM-EGFP⁺ leukocytes recruited to the heart and more LysM-EGFP⁺ leukocytes retained in the blood circulation of Nexinhib20-treated mice compared with vehicle controls (Fig. 4C, 4F, 4G). In heart LysM-EGFP⁺ leukocytes, ∼73–95% of them were Ly6G⁺ neutrophils, regardless of Nexinhib20 treatment (Fig. 4C, 4D). In blood LysM-EGFP⁺ leukocytes, ∼88–98% of them were Ly6G⁺ neutrophils, regardless of Nexinhib20 treatment (Fig. 4C, 4E). These results are consistent with a previous report (66).

If we define CD45⁺Ly6G⁺ cells as neutrophils, we find that ∼6000 neutrophils were recruited to the heart after 35 min of ischemia and 60 min of reperfusion in mice administered vehicle control (Fig. 4F). Nexinhib20 administration significantly reduced heart neutrophil counts to ∼2000 (Fig. 4F). The percentage of neutrophils in heart leukocytes was also reduced from ∼50% to ∼30% by Nexinhib20 administration (Fig. 4F). Because Nexinhib20 limited neutrophil recruitment, neutrophils retained in the blood circulation were doubled in Nexinhib20-treated myocardial I/R mice compared with vehicle controls (Fig. 4G). These data suggested that Nexinhib20 inhibited neutrophil recruitment in vivo in this mouse preclinical model of myocardial I/R injury.

Neutrophils mediate cardiomyocyte death by causing vascular plugging, releasing degradative enzymes, and generating ROS (25, 26). Because we showed that Nexinhib20 limits neutrophil recruitment to the coronary artery and Nexinhib20 was discovered as a neutrophil exocytosis inhibitor that inhibits degradative enzyme release (23) and ROS production (Supplemental Fig. 1B), we reasoned that it might be useful as a dual-functioning drug to treat myocardial I/R injury. As expected, we found that Nexinhib20 administration significantly decreased infarct size (white area in the TTC/Phthalo Blue staining) after myocardial I/R injury compared with no-drug and vehicle controls (Fig. 5A). Sham controls with little to no infarction were shown as well. Quantification of infarct area/area at risk ratios showed that Nexinhib20 significantly reduced the infarct area percentage from ∼50%, which was shown in mice administered with vehicle, to ∼40% (Fig. 5B). Quantifications of the area-of-risk percentage confirmed the stability and reproducibility of our surgical procedure (Fig. 5C). These data suggested that Nexinhib20 has the potential to treat myocardial I/R injury.

Then, we quantified heart function using a left ventricle echocardiogram. We performed the echocardiogram before and 7 d after myocardial I/R injury. Ejection fraction and fractional shortening were measured to quantify left ventricle function (Fig. 5D, 5E). Ejection fraction is a measurement, expressed as a percentage, of how much blood the left ventricle pumps out with each contraction. Fractional shortening shows the percentage of size differences of the left ventricle as a parameter of how well the left ventricle is contracting, that is, reducing its size during systole. We found that in the vehicle-treated mice, the ejection fraction (Fig. 5D) and fractional shortening (Fig. 5E) were reduced by ∼25% and ∼30%, respectively, after myocardial I/R injury, indicating a loss of left ventricle function. In Nexinhib20-treated mice, there is no significant reduction of either ejection fraction (Fig. 5D) or fractional shortening (Fig. 5E) after myocardial I/R injury. Compared to vehicle-treated mice, Nexinhib20-treated mice have significant improvement in left ventricle function 7 d after myocardial I/R injury (Fig. 5D, 5E).

Nexinhib20 was discovered as a neutrophil exocytosis inhibitor, and we confirmed this by testing β₂ integrin exocytosis after IL-8 stimulation (Fig. 2E). Importantly, we determined that Nexinhib20 also inhibited neutrophil adhesion (Fig. 1) and β₂ integrin activation (Fig. 2) without any effect on cell viability (Supplemental Fig. 1A). Thus, Nexinhib20 was confirmed as a dual-functional neutrophil inhibitor. We then found that Rac-1 is a target of Nexinhib20 (Fig. 3). Nexinhib20 inhibited Rac-1 activation in cells by antagonizing the Rac-1–GTP interaction with an IC₅₀ of 29.3 µM. Because Nexinhib20 was also reported to specifically inhibit the interaction between the small GTPase Rab27a and its effector JFC1 and neutrophil exocytosis with an IC₅₀ of 0.33 µM, it is likely that Nexinhib20 exerts a sequential and concentration-dependent inhibition. Rab27a is critical for neutrophil exocytosis (23, 68–70), adhesion molecule presentation (23, 71), migration (72), and ROS production (70). Rac-1 is important for neutrophil integrin activation (16, 73), adhesion (16, 73), migration (16, 65, 73–76), and phagocytosis (77, 78). Nexinhib20 has the potential to work as an anti-inflammatory drug by blocking neutrophil function. Whether Rab27a or Rac-1 is more important and whether they crosstalk during Nexinhib20 inhibition of neutrophil function remain to be further investigated.

Because it has been shown that Nexinhib20 inhibits both Rab27a and Rac-1, this raises concern about whether Nexinhib20 has poor specificity and may inhibit many other GTPases. All small Rab GTPases share a common mechanism of GTP-dependent binding to their respective effectors. However, each pair is characterized by highly specific binding properties, and therefore it is unlikely that Nexinhib20 would have a high affinity for other GTPases. In fact, the binding affinity of Nexinhib20 to Rac-1 is much lower than Rab27a, as we showed an ∼90-fold IC₅₀ for Rac-1 compared with Rab27a. It has also been shown that another GTPase, Rab11, was not inhibited by Nexinhib20 (23). Nevertheless, we cannot exclude the possibility that Nexinhib20 may also inhibit other GTPases, especially other Rho family GTPases, such as Rac-2, Cdc42, and RhoA.

We have shown that Nexinhb20 inhibited the interaction of recombinant Rac-1 protein and GTPγS in a dose-dependent manner. This suggests that Nexinhib20 may directly bind Rac-1 and interact with key amino acids of the GTP-binding site. The crystal structure of Rac-1 complexed with a GTP analog, guanosine-5′-(βγ-imino)triphosphate (GMPPNP), has been determined (79). The GTP binding site includes the phosphate-binding loop residues 10–17 and residues 57–61, the guanine base recognition motif residues 116–119 and 158–160, and the effector loop, residues 28–38, which interacts with the ribose and the magnesium ion. Whether Nexinhib20 directly interacts with these residues remains to be further investigated.

Nexinhib20 has been reported to inhibit recruitment of neutrophils to the liver and kidney in a LPS-induced systemic inflammation model (23) and to lung lumen and parenchyma in an acute LPS-induced lung injury mouse model (80). Our study was, to our knowledge, the first to show that Nexinhib20 reduced neutrophil recruitment to the coronary artery during myocardial I/R injury (Fig. 4), which was accomplished with multiphoton microscopy. This method could directly visualize neutrophil recruitment in a very accurate manner. The multiphoton microscopy assay also provided information in a sample with ∼100-µm thickness, which was more expansive than quantification using ∼8- to 10-µm histology sections. By combining optical clearing and light-sheet microscopy, neutrophil recruitment in the entire area at risk can be visualized and quantified (6). The attempt to use this method will be limited by instruments and experience with whole-tissue optical clearing and staining. Another method to quantify neutrophil recruitment is flow cytometry (Fig. 4C–(G) of heart single-cell suspensions (81).

Nexinhib20 showed both anti-exocytosis and anti-adhesion activities, suggesting that it might be a dual-functional drug for myocardial I/R injury. We showed that Nexinhib20 improves myocardial I/R injury in mice by reducing infarct size by ∼20% (Fig. 5A, 5B) and almost completely restoring the left ventricle function (Fig. 5D, 5E). Although Abs against β₂ integrins showed benefits in myocardial I/R injury in multiple species (35–43), the clinical trial using a β₂ integrin Ab to treat myocardial I/R injury failed (82). This might be due to the long half-life of Abs in patient circulation that also inhibits the resolution of inflammation after myocardial I/R injury. It has been shown that accurate clearance of dead cells is a prerequisite for favorable myocardial infarction healing, whereas failed resolution promotes unfavorable cardiac remodeling, which may ultimately result in heart failure (83). The clearance of dead cardiomyocytes and inflammatory neutrophils is orchestrated by macrophages, which are thought to derive from recruited Ly6C^hi monocytes (84–86), and β₂ integrin Ab can block the recruitment of monocytes (87). Meanwhile, during the clearance of dead cells, macrophages or monocytes must migrate to dead cells and perform phagocytosis. β₂ integrins are critical for both cell migration (88, 89) and phagocytosis (33, 90). Thus, a small molecule drug such as Nexinhib20 that inhibits β₂ integrin function for a shorter period (several hours) compared with Abs (several weeks to months [91]) might be an advantage in treating acute inflammatory diseases, such as myocardial I/R injury. This is because administering a small-molecule drug can alleviate the proinflammatory responses during acute inflammation and then degrade after several hours, so it will not block the later resolution of inflammation (92). We have studied the pharmacokinetics of Nexinhib20 after i.p. injection in mice (Supplemental Fig. 2) and have shown that Nexinhib20 was degraded quickly within 2 h, which is during the acute inflammation phase. Four hours after administration, the Nexinhib20 concentration is lower than ∼28.8 µg/ml (∼96 nM), which may not block the recruitment of monocytes/macrophages and their mediation of inflammation resolution and healing. This needs to be validated in future investigations. Furthermore, Nexinhib20-mediated inhibition of neutrophil exocytosis (23) and ROS production (Supplemental Fig. 1B) would also contribute to the attenuation of the I/R injury. Because we demonstrated that Nexinhib20 could prevent myocardial I/R injury, our work highlights its possible use for other acute inflammatory diseases involving neutrophils, such as noninfectious acute lung injury (93), I/R injury after transplantation (94, 95), ischemic stroke (96), systemic inflammatory response to severe injury (97), and multiple organ dysfunction syndrome (97).

We have identified Nexinhib20 as an antagonist of the Rac-1–GTP interaction. Because Rac-1 is critically involved in the functions of many cells, Nexinhib20 may be used for treatments targeting other cells. Because our study focused on myocardial I/R injury, it is important to discuss the role of Rac-1 in cardiomyocytes. Rac-1 is not only important for leukocyte activation and ROS production but it is also essential for ROS production by cardiomyocytes (98). Cardiomyocyte-specific overexpression of an active Rac mutation aggravated myocardial I/R injury (99), and myocardial I/R-induced ventricular arrhythmia was significantly decreased in cardiac-specific Rac-1 knockdown mice (100). Another Rac-1 inhibitor, NSC23766, decreased I/R-induced ventricular arrhythmia (100). Active Rac-1 was upregulated in failing myocardium of patients with ischemic cardiomyopathy and dilated cardiomyopathy (101). Statin treatment decreased myocardial Rac-1-GTPase activity (101). Rac-1 activation was involved in the hypertrophic response of cardiomyocytes (102–104), hyperglycemia-induced apoptosis of cardiomyocytes in diabetes (105), and doxorubicin-induced cardiotoxicity (106). Besides cardiomyocytes, shear stress–induced Rac-1 activation in endothelial cells is responsible for ICAM-1 expression, which is critical for the recruitment of neutrophils and inflammatory responses (107). Inhibition of Rac-1 GTPase decreases vascular oxidative stress, improves endothelial function, and attenuates atherosclerosis development in mice (108). Overall, most studies supported that inhibition of Rac-1 was beneficial to most cardiomyopathies; therefore, Rac-1–specific inhibitors, such as NSC23766 or statin, may help as well. Chemical modulation may also help to increase the affinity of Nexinhib20 to Rac-1 that increases inhibition efficiency.

Neutrophil-mediated tissue damage after I/R injury is a multifactorial process that depends on β₂ integrin–dependent neutrophil adhesion, recruitment, and secretion. In this study, we show that Nexinhib20, in addition to exocytosis, also inhibits human neutrophil adhesion and β₂ integrin activation by targeting Rac-1 GTPase. Importantly, Nexinhib20 decreased neutrophil recruitment in vivo and decreased infarct size after mouse myocardial I/R injury, further validating that inhibition of neutrophil recruitment and activation increases the likelihood of a favorable outcome during myocardial tissue damage.

The clonal HL60 cell line was generated by Alba Diz-Munoz (EMBL Heidelberg) and was a gift from the laboratory of Orion Weiner (University of California San Francisco). The CXCR2-expressing HL60 (HL60-CXCR2) cells were a gift from Dr. Ann Richmond at the Vanderbilt University School of Medicine. We acknowledge Dr. Klaus Ley from the La Jolla Institute for Immunology for supporting reagents and instruments to perform neutrophil rolling and integrin activation assays. We acknowledge Dr. Wolfgang Peti from the Department of Molecular Biology & Biophysics at UConn Health for providing the necessary instruments. We acknowledge Dr. Penghua Wang and Dr. Tingting Geng from the Department of Immunology at UConn Health for providing the Cytation 1 cell imaging multi-mode reader for the ROS production assay. We acknowledge Dr. Lixia Yue and Pengyu Zong from the Pat and Jim Calhoun Cardiology Center at UConn Health for reviewing and providing suggestions for this manuscript. We acknowledge Dr. Feifei Lin from the Shanghai Institute of Materia Medica for the service of Nexinhib20 pharmacokinetics. We acknowledge Dr. Christopher “Kit” Bonin and Dr. Geneva Hargis from UConn School of Medicine for help with scientific writing and editing of this manuscript.

The authors have no financial conflicts of interest.