Human thrombin, prostaglandin E1, citrate-dextrose solution, 2% gelatin type B, 3, 3′,5,5′-tetramethylbenzidine liquid substrate, naphthol AS-D chloroacetate (esterase) kit, N-formyl-methionyl-leucyl-phenylalanine (fMLP), phorbol 12-myristate 13-acetate (PMA), and a protease inhibitor cocktail were purchased from Sigma (St. Louis, MO, USA). The Amplex UltraRed hydrogen peroxide and the EnzChek Gelatinase/Collagenase assay kits and Hank’s balanced salt solution (HBSS) were from Thermo Fisher Scientific (Waltham, MA, USA). Recombinant human MPO, recombinant human and mouse TNF-α, and an antibody against MPO were obtained from R&D systems (Minneapolis, MN, USA). 4-aminobenzoic acid hydrazide (4-ABAH), a specific inhibitor of MPO, was purchased from Cayman Chemical (Ann Arbor, MI, USA). Percoll was from GE Healthcare Bio-Sciences AB (Uppsala, Sweden). Nycoprep 1.077 was purchased from Axis-Shield PoC-AB (Oslo, Norway). RPMI1640 media was obtained from Mediatech (Manassas, VA, USA). d-Phe-Pro-Arg-chloromethyl ketone (PPACK) was from EMD Millipore (Billerica, MA, USA). Heparin (10,000 U/mL) was purchased from Frensinius Kabi (Lake Zurich, IL, USA). Vectashield was obtained from Vector Laboratories (Burlingame, CA, USA). PE-conjugated antibodies against mouse αM (M1/70), mouse PSGL1, total human αMβ2 (ICRF44) or activated human αMβ2 (CBRM1/5), an Alexa Fluor 647-conjugated anti-Ly-6G antibody, and PE-conjugated isotype control IgGs were purchased from BioLegend (San Diego, CA, USA). A DyLight 488-conjugated anti-mouse CD42c antibody was from Emfret Analytics (Eibelstadt, Germany). An anti-β-actin antibody was obtained from Novus Biologicals (Littleton, CO, USA). Polyclonal antibodies against αM and β2 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Human umbilical vein ECs (HUVECs) were obtained from ScienCell (Carlsbad, CA, USA).

Wild-type (WT, C57BL/6) and B6.129 × 1-Mpo^tm1Lus/J (MPO KO) mice with a C57BL/6 background were obtained from Jackson Laboratory (Bar Harbor, ME, USA). Mice were used at an age of 6–10 weeks old in this study. Age-matched mice of both genders were used in all studies. The University of Illinois Institutional Animal Care and Use Committee approved all animal care and experimental procedures.

Mouse platelets were prepared as previously described (20). Washed platelets were suspended in HEPES-Tyrode buffer (20 mM HEPES, pH 7.4, 136 mM NaCl, 2.7 mM KCl, 12 mM NaHCO₃, 2 mM MgCl₂, and 5.5 mM glucose) and adjusted to a density of 2 × 10⁸ cells/mL. Human blood and mouse bone marrow neutrophils were isolated as we described (21). The concentration of neutrophils was adjusted to 1 × 10⁷ cells/mL in HBSS buffer, unless otherwise stated. Approval to collect blood samples was obtained from the University of Illinois-Chicago review board in accordance with the Declaration of Helsinki.

Platelet–neutrophil aggregation assays were performed as previously described (3). Platelets (2 × 10⁷) and neutrophils (1 × 10⁶) isolated from WT and MPO KO mice were labeled with Dylight 488-conjugated anti-CD42c and Alexa Fluor 647-conjugated anti-Ly-6G antibodies, respectively. Platelets were activated with 0.025 U/mL thrombin for 5 min at 37°C, followed by incubation with 50 µM PPACK. Neutrophils were mixed with activated platelets under a stirring condition of 1,000 rpm in an aggregometer. After a 5-min incubation, cells were fixed and analyzed by flow cytometry.

Confluent HUVEC monolayers on glass coverslips coated with 0.2% gelatin were stimulated with 20 ng/mL TNF-α for 6 h and placed into a parallel plate flow chamber (Bioptech) as we described previously (22). Mouse WT and MPO KO neutrophils (1 × 10⁶/mL) in HBSS-HEPES buffer (20 mM HBSS buffer supplemented with 10 mM HEPES pH 7.4, 2 mM CaCl₂, 1 mM MgCl₂, and 0.2% BSA) were perfused for 10 min over the activated HUVECs under venous shear (1 dyn/cm²), followed by real-time image acquisition using a Nikon microscope (ECLIPSE Ti, Melville, NY, USA) equipped with 10×/0.25 NA objective lens and recorded with a camera (CoolSNAP ES2). HBSS-HEPES buffer was then perfused for 5 min to wash out weakly bound neutrophils. The data were analyzed using NIS Elements (AR 3.2). Adherent neutrophils were monitored in a field of 0.15 mm² and counted in 4–5 separate fields.

Isolated neutrophils, 1 × 10⁵/mL (for PMA) and 1 × 10⁶/mL (for fMLP) in HBSS buffer, were treated with different concentrations of an agonist, followed by measurement of extracellular H₂O₂ using the Amplex UltraRed reagent (ThermoFisher) according to the manufacturer’s protocol. The fluorescence signal was read in a FlexStation II benchtop fluorometer (Molecular Devices) with an excitation wavelength of 530 nm and an emission wavelength of 590 nm.

Isolated neutrophils in HBSS buffer or phosphate-buffered saline, pH 7.4 (PBS) were pretreated with vehicle or the indicated concentrations of 4-ABAH, a specific inhibitor of MPO, for 30 min at 37°C. The cells were sonicated in PBS (8 × 10⁶ and 1 × 10⁶/mL for mouse and human neutrophils, respectively) containing protease inhibitor cocktail and 1 mM PMSF. The cell lysate was collected after centrifugation and sample reactions were prepared in duplicates. Each reaction contained the lysates of 4 × 10⁵ mouse neutrophils or 1 × 10⁴ human neutrophils with 0.3 mM H₂O₂ and 10.5% 3,3′,5,5′-Tetramethylbenzidine solution (ThermoFisher). The reaction was quenched with the addition of 0.8 N HCl and the absorbance was read on a PHERAstar plate reader (BMG Labtech) at 450 nm. In some experiments, 1.6 × 10⁸/mL mouse neutrophils were stimulated with or without 10 ng/mL PMA for 10 min at 37°C after pretreatment with vehicle or 500 µM 4-ABAH. The supernatant was collected and used in the MPO activity assay. In other experiments, recombinant human MPO (100 ng) was treated with vehicle or the indicated concentrations of 4-ABAH and used in the assay.

Neutrophils (2 × 10⁷/mL in RPMI1640 media) were stimulated with the indicated concentrations of fMLP or PMA for 10 min at 37°C. After centrifugation, the supernatant was used to measure gelatinase activity using the EnzChek^® Gelatinase/Collagenase Assay Kit (ThermoFisher). The fluorescence intensity was read on a PHERAstar plate reader with an excitation wavelength of 485 nm and emission wavelength of 520 nm for 4 h. The gelatinase activity was calculated at a fixed time point within the linear range of data points.

Neutrophils (2 × 10⁶/mL in RPMI1640 media) were treated with or without different concentrations of fMLP or PMA for 10 min at 37°C. In some experiments, 200 nM H₂O₂ was added immediately prior to agonist stimulation. In other experiments, cells were treated with vehicle or 500 µM 4-ABAH for 30 min at 37°C prior to agonist stimulation. For mouse neutrophils, cells were incubated with phycoerythin (PE)-conjugated antibodies against αMβ2, PSGL-1, or isotype control immunoglobulin (IgG). For human neutrophils, cells were labeled with PE-conjugated antibodies against total αMβ2 (ICRF44), activated αMβ2 (CBRM1/5), or isotype control IgG. The cells were then fixed with 1% paraformaldehyde, followed by flow cytometric analysis using a Cyan-ADP flow cytometer (Beckman Coulter). The mean or median fluorescence intensity value of antibodies were normalized to that of the respective unstimulated control and expressed as a fold increase.

Mouse neutrophils were lysed at a concentration of 5 × 10⁷/mL in a lysis buffer (50 mM Tris–HCl, pH 7.4 containing 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 150 mM NaCl, 2 mM EDTA, protease inhibitor cocktail, and 1 mM PMSF) on ice. For the releasate immunoblot, mouse neutrophils (2 × 10⁷/mL in RPMI1640 media) were stimulated with the indicated concentrations of an agonist for 10 min at 37°C. The supernatant was collected after centrifugation and the cell pellets lysed with the lysis buffer at a volume equal to the volume of supernatant collected.

Intravital microscopy was performed in a mouse model of TNF-α-induced cremaster venular inflammation as we described previously (2). WT and MPO KO male mice (6–8 weeks old) were anesthetized with i.p. injection of ketamine and xylazine, followed by exteriorization of the cremaster muscle. Neutrophils and platelets were visualized by infusion of Alexa Fluor 647-conjugated anti-Ly-6G [0.05 µg/g body weight (BW)] and DyLight 488-conjugated anti-CD42c (0.1 µg/g BW) antibodies, respectively. Images were captured in 10 different cremaster venules with a diameter of 25–40 µm in one mouse and recorded using an Olympus BX61W microscope with a 60 × 1.0 NA water immersion objective and a high-speed camera (ORCA-Flash4.0 V2, C11440, Hamamatsu). The numbers of rolling and adherent neutrophils were determined in an area of 0.02 mm² over 5 min, followed by normalization to vessel length. The kinetics of platelet accumulation was monitored by the integrated median fluorescence intensity of the anti-CD42c antibody. The fluorescence signal was normalized to the number of adherent neutrophils and the vessel length and plotted as a function of time. Time “0” was set to when image capture began on each vessel. Data were analyzed using Slidebook (version 6.0, Intelligent Imaging Innovations).

Hepatic I/R injury was induced as we previously described (3). WT and MPO KO mice were anesthetized with ketamine and xylazine and placed on a temperature-controller to maintain the body temperature at 37°C. The liver was exposed with a midline incision. A vascular clamp was placed on the portal vein and hepatic artery caudal to the porta hepatis for 30 min. The clamp was then removed, and the incised abdomen was covered with gauze soaked in sterile saline. Three hours after reperfusion, blood was drawn from the inferior vena cava, and the median and left lobes of the liver were taken out. The damage of hepatocytes was determined by measuring serum levels of aspartate aminotransferase. The median lobe was fixed in 10% formalin for the paraffin-embedded sections. The liver sections were stained with naphthol ASD-chloroacetate esterase kit (Sigma). Images were taken using a Zeiss Axioplan 2 microscope (Carl Zeiss, Oberkochen, Germany) with a 20 × 0.5 NA objective and an Axiocam MRc5 camera. The number of transmigrated neutrophils was counted outside the hepatic vessels.

Data were analyzed with the GraphPad Prism 7 software (v7.04). Statistical significance was assessed by Student’s t-test or one- or two-way analysis of variance (ANOVA) and Tukey’s test. A P value less than 0.05 was considered significant.

Our previous work in NOX2 KO mice demonstrated a crucial role for NOX2-generated ROS in mediating neutrophil–platelet interactions during TNF-α-induced vascular inflammation. To examine the role of MPO in neutrophil recruitment and platelet–neutrophil interactions, we assessed MPO KO mice in the same TNF-α-induced venular inflammation model. Compared to WT mice, MPO KO mice did not exhibit any defect in neutrophil rolling over the inflamed endothelium (Figures 1A,B; Videos S1 and S2 in Supplementary Material). Deletion of MPO significantly reduced the number of neutrophils adherent to the inflamed endothelium but increased the number of transmigrated neutrophils across the endothelial barrier (Figures 1C,D). When normalized to the number of adherent neutrophils, the platelet signal was decreased in MPO KO mice as compared with WT mice (Figure 1E). In a control experiment, MPO deletion did not affect blood counts (Table 1). Since deletion of hematopoietic cell NOX2 results in a remarkable decrease in platelet–neutrophil interactions without affecting neutrophil adhesion to activated ECs (3), these results indicate that neutrophil MPO plays a distinct role in neutrophil recruitment and platelet–neutrophil interactions during vascular inflammation.

To directly examine whether MPO affects the interaction between neutrophils and platelets, we performed the in vitro neutrophil–platelet aggregation assay under stirring conditions mimicking blood shear. We previously reported that activated platelets and neutrophils aggregate with each other under shear conditions, creating a new population in the R1 gate (21). As quantified by the fluorescence intensity of anti-CD42c antibodies in the R1 gate, deletion of neutrophil MPO did not affect neutrophil–platelet aggregation (Figures 2A,B). These results suggest that the decreased platelet–neutrophil interaction seen in intravital microscopy was likely derived from a significant decrease in the number of adherent neutrophils remaining in the vessel due to increased emigration.

We then investigated the contribution of MPO to neutrophil adhesion to activated ECs, which is the step prior to neutrophil transendothelial migration. To study this, we perfused WT and MPO KO neutrophils over a TNF-α-stimulated HUVEC monolayer under venous shear. Compared to WT neutrophils, MPO KO neutrophils exhibited an increased adhesion to the inflamed EC monolayer (Figures 2C,D). Altogether, our data show that MPO negatively regulates neutrophil–EC interactions.

We and others demonstrated that αMβ2 integrin is required for neutrophil–EC interactions, neutrophil transmigration, and neutrophil–platelet interactions (3, 22–24). Unlike other integrins which are mostly present on the plasma membrane, αMβ2 integrin is also stored in secondary (specific) and tertiary (gelatinase) granules and secretory vesicles (25). When neutrophils are stimulated, the integrin is translocated to the plasma membrane and activated, promoting neutrophil adhesive function. We observed that compared to WT neutrophils, MPO KO neutrophils exhibited increased surface levels of αMβ2 integrin after stimulation with different concentrations of fMLP or PMA (Figures 3A–C). There was no statistical difference of the integrin level between unstimulated WT and KO neutrophils (Figure 3A). As assessed by immunoblotting, the increased surface level of αMβ2 integrin in MPO KO neutrophils was not due to an increased expression of αMβ2 integrin protein (Figure S1A in Supplementary Material). In another control experiment, this increased surface expression is not derived from a general increased reactivity in MPO KO neutrophils since PSGL-1 shedding, another surface marker of neutrophil activation, was not increased by MPO deletion (Figure S1B in Supplementary Material). To further explore the role of MPO in degranulation, we assessed the release of gelatinase from tertiary granules. We found that treatment of neutrophils with fMLP or PMA released gelatinase (Figure S2A in Supplementary Material) and that there was no difference in the degranulation of gelatinase containing granules in WT and MPO KO neutrophils (Figures S2B,C in Supplementary Material). As a control experiment, both agonists concentration-dependently increased secretion of MPO (Figure S2D in Supplementary Material) from primary granules. These results suggest that the pro-adhesive and pro-migratory phenotype in MPO KO neutrophils may result from increased αMβ2 degranulation from granules which are different from ones containing gelatinase.

In the neutrophil respiratory burst, MPO generates the cytotoxic HOCl from H₂O₂. Recently, we demonstrated that loss of H₂O₂ generation in NOX2 KO neutrophils impairs the ligand-binding activity of αMβ2 integrin without affecting the membrane translocation during cell activation and that such a defect was completely rescued by the addition of 1 µM H₂O₂ to NOX2 KO cells (3). Since MPO deletion enhanced the surface level of αMβ2 integrin following agonist stimulation (Figures 3A–C), we hypothesized that loss of MPO accumulates H₂O₂ in the extracellular space. To examine this, we used the Amplex Red assay which detects extracellular H₂O₂ (26). Compared to WT neutrophils, MPO KO neutrophils had increased levels of extracellular H₂O₂ in response to fMLP or PMA (Figures 3D,E). To test whether this increase in H₂O₂ levels facilitates the membrane translocation of αMβ2 integrin, we added exogenous H₂O₂ prior to stimulation with fMLP or PMA to WT neutrophils and examined the surface levels of αMβ2 integrin. With the addition of 200 nM H₂O₂, WT neutrophils increased the surface level of αMβ2 integrin (Figures 3F,G). These results suggest that loss of MPO results in increased extracellular H₂O₂ levels upon cell activation, which facilitates the membrane translocation of αMβ2 integrin.

Since the enzyme activity of MPO is critical for the production of HOCl from H₂O₂ (8), we utilized the specific MPO inhibitor, 4-ABAH (27). While 4-ABAH (1–100 µM) inhibited the activity of recombinant human MPO in a concentration-dependent manner (Figure S3A in Supplementary Material), much higher concentrations (100–500 µM) of 4-ABAH were required to inhibit MPO in mouse neutrophils (Figure S3B in Supplementary Material), which may result from poor cell permeability of the inhibitor. We observed that pretreatment with 500 µM 4-ABAH inhibited MPO activity by >80% in neutrophils. MPO KO neutrophils had no activity, demonstrating specificity of this assay in neutrophils. With pretreatment of the inhibitor, we observed >75% decrease in the activity of MPO released from PMA-stimulated neutrophils (Figure S3C in Supplementary Material), indicating that the released MPO bound to 4-ABAH did not have any activity. As a control, we confirmed that treatment with the inhibitor did not impair MPO secretion after agonist stimulation (data not shown).

Pretreatment of WT neutrophils with 500 µM 4-ABAH increased the surface level of αMβ2 integrin in response to fMLP or PMA (Figures 4A,B). In a control experiment, treatment of MPO KO neutrophils with 500 µM 4-ABAH did not further increase the surface level of αMβ2 integrin following agonist stimulation (Figure S4 in Supplementary Material), suggesting that 4-ABAH is specific for MPO at the concentration used in this assay. In human neutrophils pretreated with 4-ABAH, we observed a significant increase in the surface level and activation of αMβ2 integrin following fMLP or PMA stimulation as determined by the conformation-specific antibodies against total (ICRF44) or activated αMβ2 (CBRM1/5) (Figures 4C–F). In line with our findings in MPO KO neutrophils, these results suggest that inhibition of MPO activity enhances the membrane translocation and activation of αMβ2 integrin.

We recently reported that NOX2-generated ROS positively modulate neutrophil infiltration into damaged liver tissue, contributing to the pathogenesis of I/R-induced hepatic injury in mice (3). Thus, we sought to examine the pathophysiological role of MPO in the same disease model. We found that compared to WT mice, MPO KO mice exhibited a remarkable increase in the number of transmigrated neutrophils into the interstitial space following I/R injury (Figures 5A,B). Nevertheless, MPO deletion significantly reduced the tissue damage as assessed by serum levels of aspartate aminotransferase (Figure 5C). These results suggest that neutrophil MPO-generated oxidants, such as HOCl, are the major players in tissue damage under inflammatory conditions and support our findings that loss of MPO increases the surface level of αMβ2 integrin upon activation, which results in a pro-adhesive and pro-migratory phenotype in sterile inflammation.