α-1 Antitrypsin regulates human neutrophil chemotaxis induced by soluble immune complexes and IL-8 (original) (raw)
AAT activity regulates IL-8 and immune complex–induced neutrophil chemotaxis. The chemotactic index of ZZ neutrophils obtained from 6 clinically stable AATD individuals was compared with that of control neutrophils (MM) from 6 healthy donors. Results revealed that in response to IL-8 (10 ng), ZZ neutrophils exhibited a 2- or 8-fold increase in chemotactic activity when experiments were performed in PBS or serum (50% v/v), respectively (Figure 1A). In addition, in response to IL-8 the chemotactic index of MM neutrophils suspended in PBS was superior to that of MM cells suspended in serum (P = 0.02). Moreover, the chemotactic index of ZZ neutrophils in response to soluble immune complex (sIC) (10% v/v) was 25% lower than that of MM control cells when performed in PBS (P = 0.001), yet 3.6-fold higher when experiments were repeated in the presence of respective ZZ and MM sera (Figure 1B).
Chemotactic analysis of neutrophils in response to AAT. (A) Increased mean chemotactic index of ZZ-AATD neutrophils (n = 6) compared with healthy control (MM) cells (n = 6) in response to IL-8 (10 ng/2 × 107 cells). Assays were preformed in PBS or serum (50% v/v) from respective ZZ or MM individuals. (B) Decreased mean chemotactic index of ZZ-AATD neutrophils (n = 6) in PBS in response to sIC (10% v/v), yet increased mean chemotactic index of ZZ-AATD neutrophils in serum (50% v/v) compared with healthy control cells (n = 6). (C) An increase in chemotactic inhibition efficiency of increasing concentration of AAT (0.2–27.5 μM). Black bar, positive IL-8 (10 ng) control; white bar, negative HSA (27.5 μM) control. *P < 0.05 versus IL-8 control. (D) Neutrophil chemotaxis in a linear gradient of IL-8 (1, 10, 20, or 40 ng) or complex gradient in the presence of inhibitory AAT (27.5, 6.9, 0.9, 0.2 μM) (*P < 0.05 compared with AAT-untreated cells). (E) Increased chemotactic inhibition efficiency of increasing concentrations of AAT (0.4–27.5 μM). Black bar, positive sIC (10% v/v) control; white bar, negative HSA (27.5 μM) control. *P < 0.05 versus sIC control. Experiments illustrated in A–E were performed in triplicate on 3 consecutive days, and for comparative analysis the PBS control was set at a chemotactic index of 1. Each measurement is the mean ± SEM.
To investigate whether the increased chemotactic activity of ZZ neutrophils in response to IL-8 was a result of AATD, we explored the effect of exogenous AAT on neutrophil chemotaxis. As illustrated in Figure 1C, the chemotactic index of MM neutrophils in response to IL-8 (10 ng) was 6.05 ± 0.66. AAT inhibited MM neutrophil chemotaxis in a dose-dependent manner, with an IC50 of 0.5 μM. HSA (control plasma protein, 27.5 μM) did not affect neutrophil chemotaxis. At a higher concentration, AAT (6.9 μM) reduced the MM neutrophil chemotactic index produced by 10 ng or 20 ng IL-8 by 91.5% (P = 0.002) and 46.8% (P = 0.07), respectively (Figure 1D). Biologically relevant serum concentrations of AAT (27.5 μM) abolished neutrophil chemotaxis in response to 1 ng or 10 ng IL-8 and reduced the cellular chemotaxis index to 20 ng and 40 ng IL-8 by approximately 72% and 52% (P = 0.009 and P = 0.09, respectively). To investigate the inhibitory effect of AAT on IL-8’s chemotactic activity (versus chemokinesis), we also used checkerboard analysis with serial dilutions of IL-8 above and below the filter. Neutrophil migration depended on the presence of an IL-8 gradient, implying that AAT inhibits directed cell migration rather than chemokinesis (Supplemental Figure 1, A and B; supplemental material available online with this article; doi:10.1172/JCI41196DS1). The effect of exogenous AAT (0.4–27.5 μM) on the chemotactic index of MM neutrophils in response to sIC (10% v/v) was explored next. As illustrated in Figure 1E, AAT inhibited MM neutrophil chemotaxis in a dose-dependent manner, with an IC50 of 8.2 μM, while HSA (27.5 μM) had no effect on sIC-mediated chemotaxis.
To understand (a) how AAT inhibited both IL-8 and sIC mediated MM neutrophil chemotaxis, (b) why ZZ cells demonstrated an increased chemotactic response to IL-8, and (c) why ZZ neutrophils illustrated conflicting chemotactic responses to sIC in PBS and serum was the goal of subsequent experiments.
AAT is localized to the neutrophil membrane. To establish how AAT impacted on neutrophil chemotaxis, it was necessary to pinpoint its location within the cell. By subcellular fractionation of neutrophils, AAT was defined as a component of the cell membrane (Figure 2A). Successful separation of plasma membranes from secretory vesicles was validated by Western blotting for HLA class I and HSA, and 83% ± 1.5% of AAT was localized within the plasma membrane fraction (Figure 2B). Confocal laser scanning microscopy was employed to confirm localization of AAT to the membrane and showed bright foci of anti-AAT labeling in the membrane region of isolated cells (Figure 2C). In addition, AAT was found in association with detergent-free low-density sucrose fractions (15% w/w) containing membrane lipid rafts (Figure 2D). A similar distribution of flotillin-1 was observed, and treatment with the cholesterol-sequestering agent methyl-β-cyclodextrin (MβCD) diminished AAT and reduced the level of flotillin-1 in the low-density sucrose fractions. 2D electrophoresis of isolated MM neutrophil membranes (Supplemental Figure 2A) and purified membrane lipid rafts (Figure 2E) revealed 4 isoforms of AAT of similar molecular mass (~52 kDa). By employing FACS analysis of nonpermeabilized cells, we detected AAT on the plasma membrane of unstimulated resting MM healthy neutrophils (Figure 2F). In contrast, neutrophils isolated from ZZ-AATD individuals illustrated significantly lower levels of AAT (mean fluorescence: control MM cells, 31.41; ZZ-AATD cells, 17.52; P = 0.001). Fractionation of ZZ neutrophil cytosol, membrane, and granule compartments localized Z-AAT to neutrophil membranes, with reduced levels (~50%) confirmed by Western blotting (Supplemental Figure 2B). Collectively, these findings support localization of AAT to the neutrophil membrane and reveal that ZZ-AATD cells express significantly lower levels of AAT.
Localization of AAT in peripheral blood neutrophils. (A) Coomassie blue–stained gel of normal MM neutrophils subjected to subcellular fractionation yielding a cytosol fraction (lane 1), combined secretory vesicle/plasma membrane fraction (lane 2), and primary and secondary granule fraction (lanes 3 and 4, respectively). Western blotting employed antibodies to MPO and PR3 as markers of primary granules and antibodies against p22phox of cytochrome _b_558 as a marker of secondary granules and combined secretory vesicle/membrane fractions. AAT was localized to the secretory vesicle/membrane fraction at the interface between the 17.5/35% (w/w) sucrose. (B) By Percoll gradient fractionation, approximately 83% of AAT was localized to the plasma membrane (PM) compared with secretory vesicles (SV). As controls, HLA served as a marker of PM and HSA as a marker of SV. (C) By confocal microscopy, the distribution of AAT in non-permeabilized MM neutrophils was predominantly localized to the membrane margin of the cell. Cell nuclei are stained red. Scale bar: 10 μm. (D) Detection of AAT and flotillin-1 in the low-density lipid raft fractions (15% w/w sucrose) of human MM neutrophil lysates: untreated (–) or treated with MβCD (10 mM) (+). Immunoblots were probed with polyclonal anti-AAT and monoclonal anti–flotillin-1. (E) Coomassie blue–stained 2D SDS-PAGE gel of isolated MM neutrophil membrane lipid rafts (top panel). A Western blot probed with polyclonal anti-AAT revealed 4 associated AAT isoforms (lower panel). (F) Flow cytometry analysis of membrane-bound AAT in isolated resting MM (green) and ZZ-AATD (red) neutrophils. The level of AAT was found to be significantly higher on resting MM neutrophils. The isotype control antibody is illustrated in black (filled). Experiments in A, D, and E are each representative gels and blots of 3 separate experiments. Results in B represent experiments performed in triplicate on 3 consecutive days, and each bar is the mean ± SEM. Confocal analysis in C and FACS analysis in F are one illustrative result from 3 independent experiments.
Origin of membrane-associated AAT. To clarify whether AAT was being produced by the neutrophil, as has been described by others (15), or whether it was being absorbed from the serum, we compared the phenotype of MM and ZZ-AATD neutrophil membrane-associated AAT with that of a ZZ-AATD individual after liver transplantation. Figure 3A shows the different patterns of phenotypes frequently observed for serum of MM and ZZ-AATD individuals (16). The phenotype of the ZZ-AATD serum sample after liver transplantation was entirely MM, a result confirmed by the use of an ELISA specific for the ZZ protein (17, 18) (Figure 3B). In contrast, neutrophil membranes of the ZZ-AATD liver transplant recipient contained detectable levels of Z-AAT protein (Figure 3B). Additionally, although these cells contained lower levels of membrane Z-AAT compared with a non-transplant patient, greater levels of total AAT (Z and M isoforms) were detected by ELISA (Figure 3C) and immunoblotting (Figure 3D), most likely due to absorption of MM AAT from the serum. Collectively, these experiments suggest that both serum- and neutrophil-derived AAT binds the cell membrane.
Serum- and neutrophil-derived AAT binds the neutrophil membrane. (A) Isoelectrofocusing patterns for ZZ, heterozygous MZ, or MM serum compared with serum of a ZZ-AATD individual after liver transplantation (ZZ+LT). The isoform numbers for the M and Z variants are indicated. The AAT pattern of the ZZ+LT sample was of the M variant. (B) ELISA for detection of the Z-form of AAT in serum and neutrophil membranes. Results in arbitrary fluorescence units (FU) indicate positive detection of Z-AAT in serum and neutrophil membranes of a ZZ-AATD patient but only in neutrophil membranes after LT (P = 0.2 between ZZ and ZZ+LT neutrophil membranes). An ELISA for detection of total AAT (M and Z variants) (C) and Western blots probed with goat (Gt) polyclonal anti-AAT (D) revealed significantly higher levels of total AAT on ZZ-AATD neutrophil membranes after liver transplantation (ZZ+LT) compared with a non-transplant ZZ-AATD patient (*P = 0.01, **P = 0.001). Results represent experiments performed in triplicate, and data in B and C are the mean ± SEM.
Proinflammatory cytokine exposure causes release of AAT within a protein complex. The effect of proinflammatory stimuli (IL-8 or TNF-α) on release/shedding of AAT from the neutrophil membrane was investigated. MM neutrophils were treated with TNF-α (10 ng), a known potent inducer of L-selectin (CD62-L) shedding (Figure 4A). After 10 minutes of treatment, a significant decrease in cell surface AAT was detected by flow cytometry (mean fluorescence: control untreated cells, 20.7; TNF-α–treated cells, 10.28; P < 0.001) (Figure 4A). A corresponding increase in released AAT in the surrounding medium was detected by Western blot analysis, with a maximum increase in extracellular released AAT at 40 ng TNF-α and 20 ng IL-8 (Figure 4B). To assess the kinetics of shedding, we treated neutrophils with 10 ng IL-8 or TNF-α and assessed extracellular released AAT at 2, 5, and 10 minutes. As depicted in Figure 4C, an immunoreactive band corresponding to released AAT was detected at 5 and 10 minute after TNF-α treatment and at an earlier time point of 2 minutes in response to IL-8. Maximum release of AAT from neutrophil membranes exposed to either IL-8 or TNF-α occurred at 10 minutes (Supplemental Figure 3, A–D). ELISA was performed to quantify levels of released AAT, and at the 2- and 5-minute time points, neutrophils released 4.46 ± 1.13 and 5.25 ± 0.44 ng AAT per 1 × 107 neutrophils in response to IL-8 (10 ng) (Figure 4D). In contrast, and at the same time points, neutrophils released significantly lower levels of AAT in response to TNF-α treatment (P = 0.02 and P = 0.014 for 2 and 5 minutes, respectively). To investigate whether neutrophils synthesized AAT to substitute for membrane-released AAT, we exposed cells to IL-8 (10 ng) and quantified levels of extracellular and membrane-associated AAT for up to 16 hours. Results revealed no increase in either membrane-associated or newly released levels of AAT (Supplemental Figure 3E), suggesting that de novo protein synthesis of AAT did not occur.
Release of biologically active AAT from TNF-α– or IL-8–treated MM neutrophils. (A) Membrane expression of L-selectin or AAT on neutrophils at baseline (green) or in response to TNF-α (red). (B) Western blot showing extracellular released AAT (52 kDa) following treatment with TNF-α or IL-8 in a dose- (B) and time-dependent manner (C). Serum AAT (52 kDa) was loaded as a positive antibody control, and untreated cells are in lanes labeled as control (Con). (D) Time course analysis of the release of AAT (*P < 0.05). (E) Gel filtration chromatography of neutrophil-released AAT. The start material (St) was released AAT from IL-8–treated neutrophils (10 ng). Inset: Western blot analysis illustrating altered migration of purified serum AAT (arrow) compared with neutrophil-released AAT (arrowheads). By Western blot analysis, fraction 28 contained neutrophil-released AAT and chromatographed at a molecular mass of 100–110 kDa. Fractions 30–33 were run on separate gels. (F) Formation of AAT-protease complexes. Samples were immunoblotted employing a rabbit anti-human AAT antibody. Lanes 1 and 3 show serum purified AAT and AAT released from neutrophils, respectively. Lanes 2 and 4 illustrate serum and neutrophil-released AAT incubated with NE. FACS analysis illustrated in A is a representative result of 3 independent experiments. B, C, and F are each representative Western blots of 3 separate experiments. Results in D represent experiments performed in triplicate. Each measurement is the mean ± SEM. FPLC analyses (E) were performed in triplicate on 2 consecutive days.
Native gel electrophoresis was employed to examine potential conformational differences between serum purified AAT and IL-8–induced neutrophil-released AAT. Serum AAT migrated faster and ran as a predominant single band (Figure 4E, inset). In contrast, neutrophil-released AAT migrated differently from serum AAT, showing a smeared, retarded mobility suggestive of a significant degree of conformational heterogeneity and/or protein-protein complexation. For assessment of the complexed state of the AAT species present, neutrophil-released AAT was chromatographed by gel filtration on a Superose 6 column. Protein elution was monitored by Western blot analysis, and as can be seen in Figure 4E, neutrophil AAT protein eluted at a position corresponding to a complex molecular mass of approximately 100–110 kDa. To exclude the possibility that neutrophil-released AAT eluted at increased molecular mass due to a protease-inhibitor complex, we confirmed the affinity of neutrophil-released AAT for neutrophil elastase (Figure 4F).
AAT interacts with the neutrophil membrane via the GPI-anchored lipid raft membrane protein FcγRIIIb. AAT was immunoprecipitated from IL-8–induced neutrophil-shedded material by rabbit polyclonal anti-AAT and the immunoprecipitates resolved by denaturing and reducing SDS-PAGE. A Coomassie blue stained gel is illustrated in Figure 5A, and a putative 45- to 50-kDa AAT binding partner was identified by LC-MS/MS as FcγRIIIb (CD16b). FcγRIIIb is a GPI-linked low-affinity IgG receptor, and to exclude the possibility of experimental binding artefact, we performed reciprocal immunoprecipitation of FcγRIIIb and subsequent identification of AAT by LC-MS/MS (Figure 5B). This finding was further supported by the presence of FcγRIIIb and AAT on Western blots of an immunoprecipitation for AAT (Figure 5C, top panel) and for FcγRIIIb, respectively (Figure 5C, lower panel). By expanding the gel filtration result illustrated in Figure 4E by Western blot analysis for FcγRIIIb, we observed the coelution of AAT and FcγRIIIb in fraction 28 (Supplemental Figure 4A). Moreover, native gel electrophoresis and Western blot analysis of fraction 28 supported the possibility of an AAT-FcγRIIIb interaction, while excluding formation of AAT polymers or neutrophil elastase (NE) and PR3 as possible binding partners within this complex (Supplemental Figure 4B). The exposure of AAT-coated surfaces to FcγRIIIb revealed that FcγRIIIb bound to AAT in a concentration-dependent manner but not to a second non-GPI-linked IgG receptor, FcγRIIa (Figure 5D). In addition, an ELISA utilizing mouse capture antibody to FcγRIIIb and goat detection antibody for AAT quantified significant levels of cell-free serum FcγRIIIb-AAT complex in serum of CF patients (Figure 5E). Reciprocal ELISA utilizing goat capture antibody to AAT and mouse detection antibody for FcγRIIIb quantified considerably higher levels of AAT-FcγRIIIb complex in CF serum, compared with MM and ZZ-AATD serum (Supplemental Figure 4C). Since AAT and FcγRIIIb coimmunoprecipitated, it also appeared logical that AAT and FcγRIIIb would colocalize on the neutrophil membrane. Analysis by confocal microscopy indicated that, as expected, AAT (red) and FcγRIIIb (green) stained the plasma membrane (ring-like staining at the edge of the cells) (Figure 5F), and superimposition of the two images clearly identified colocalization (yellow) of the two signals. We next tested the hypothesis that enzymatic cleavage of the GPI-anchored FcγRIIIb with phosphatidylinositol-specific phospholipase C (PI-PLC) would also displace the associated AAT binding partner (Figure 5G). Western blot analysis of PI-PLC–treated (1 U/1 × 106 cells) neutrophil membranes resulted in release of AAT and FcγRIIIb, but not p22phox, a control non-GPI membrane component. Further treatment of membranes with NaCl (350 mM) caused release of AAT but not FcγRIIIb (Figure 5G). In addition, in comparison to MM donors, neutrophils of ZZ-AATD patients showed a significant decrease in the level of FcγRIIIb protein (Figure 5H and Supplemental Figure 4D). The observation that levels of FcγRIIIb were significantly elevated in serum of ZZ-AATD patients compared with normal MM controls (P = 0.01) (Figure 5I) contrasts results illustrating similar levels of FcγRIIIb-AAT complex (Figure 5E), possibly indicating that a proportion of FcγRIIIb in ZZ-AATD serum is AAT unbound. Collectively, these experiments further suggest FcγRIIIb as a binding partner that is necessary for AAT membrane presentation and suggest that AAT may functionally impact on maintaining FcγRIIIb membrane expression.
AAT binds FcγRIIIb on the neutrophil membrane. (A) Immunoprecipitate (IP) employing rabbit (Rb) antibody to AAT. The AAT binding partner (labeled 1) was identified as FcγRIIIb (40–60 kDa). (B) Reciprocal immunoprecipitate with FcγRIIIb goat (Gt) antibody. Coimmunoprecipitated proteins were identified as FcγRIIIb (labeled 1; accession number AA128563) and as AAT (labeled 2; accession number CAJ15161). (C) Western blots of immunoprecipitate reactions were performed with goat antibody against AAT (top panel) and mouse (Mo) antibody against FcγRIIIb (bottom panel). Serum purified AAT and recombinant human (Rh) FcγRIIIb are included as positive controls. (D) Binding of Rh FcγRIIIb or Rh FcγRIIa to AAT (*P = 0.02 compared with control containing no FcγRIIIb). (E) FcγRIIIb-AAT complex in serum of CF (n = 4) individuals was significantly higher than that of normal MM (Con; n = 6) or AATD (n = 6) subjects (*P < 0.05, CF versus control or ZZ). (F) Colocalization (merged image in yellow) of rhodamine labeling for AAT and FITC for FcγRIIIb, at the membrane margin of cells (×64 magnification, ×4 zoom). (G) NaCl and PI-PLC treatment of neutrophil membranes. Immunoblots included untreated membranes (Con) and antibodies against p22phox. (H) Flow cytometry analysis of membrane-bound FcγRIIIb (mean fluorescence: MM, 41.93; ZZ, 15.44). The isotype control antibody is in black (filled). (I) Levels of FcγRIIIb in serum of ZZ-AATD individuals and MM controls (Con) (*P = 0.01). In A–C and G, images are representative results from 1 of 3 separate experiments. Lanes in A and C were run on the same gel but were noncontiguous. D, E, H, and I represent results performed in triplicate. Images in F are representative results of 2 independent experiments.
AAT regulates release of FcγRIIIb by inhibiting metalloprotease ADAM-17 activity. Evidence exists that neutrophil chemotaxis requires activity of surface sheddases, including ADAM-17, which has been implicated in the shedding of FcγRIIIb (19), a prerequisite for neutrophil chemotaxis (20–23). This was confirmed when cells were exposed to sIC (10% v/v) and the extracellular supernatants analyzed by Western blot for the presence of released FcγRIIIb (Figure 6A). Release of FcγRIIIb was observed at 5 and 10 minutes after exposure, an effect greatly reduced by the specific ADAM-17 inhibitor TAPI-1 (10 μM) (24). To investigate the possibility that AAT regulated sIC-induced chemotaxis by modulating shedding of FcγRIIIb, we exposed cells to sIC (10% v/v) in the presence or absence of AAT and monitored the kinetics of FcγRIIIb shedding by immunoblotting. As illustrated in Figure 6B, AAT inhibited the release of FcγRIIIb elicited by sIC at 5 and 10 minutes by 58% ± 1.2% and 69% ± 5.9%, respectively (P < 0.05). The ability of AAT to directly inhibit ADAM-17 activity was analyzed fluorometrically, and the results indicated that AAT inhibited ADAM-17 enzymatic activity with a Ki value of 5.7 μM (Figure 6C). In addition, an equimolar concentration of AAT competitively inhibited ADAM-17, with an association rate constant of 2.23 × 104 ± 0.37 × 104 M–1s–1, although this may underestimate the physiological inhibitory capacity, as AAT is one of the most abundant proteins in the circulation. As polymerization and oxidation have previously been shown to alter enzymatic activity of AAT against NE, we tested whether an intact reactive center loop was also essential for inhibiting ADAM-17 activity. As illustrated in Figure 6D, a loss of anti–ADAM-17 activity was observed following oxidation of AAT with N-chlorosuccinimide, heat-induced polymerization, or by the AAT cleavage product C-36. Additionally, results revealed a significant 68% reduction in ADAM-17 activity in the presence of 27.5 μM AAT (P = 0.001), a value comparable to inhibition of ADAM-17 activity by TAPI-1 (10 μM) (Figure 6D). Moreover, the exposure of AAT-coated surfaces to ADAM-17 (250 ng) together with native AAT (250 ng) reduced the level of detectable bound ADAM-17 by approximately 36% (P = 0.005), a competitive effect not observed with either polymerized AAT or the C-36 cleaved fragment of AAT (Figure 6E). From these experiments, we conclude that neither polymers nor the C-terminal fragment of AAT can inhibit binding of native AAT to ADAM-17.
AAT prevents neutrophil chemotaxis in response to sIC through inhibition of ADAM-17 activity. (A) Western blots showing time course of sIC-induced release of FcγRIIIb from MM neutrophils with or without TAPI-1 (10 mM). (B) Quantification of FcγRIIIb release from MM neutrophils treated with sIC (10% v/v) with or without AAT (27.5 mM) (*P < 0.05). Inset: Western blot illustrating FcγRIIIb release at the 10-minute time point. (C) Competitive inhibition of ADAM-17 by AAT. (D) The effect of oxidation (Ox), polymerization (Poly), or the cleaved AAT peptide (C-36 fragment) on TACE inhibition (percent inhibition of maximum activity) was compared with native AAT. *P < 0.05, native AAT or TAPI-1 versus maximal activity; §P = 0.008, polymerized AAT versus native AAT (27.5 mM). (E) Binding of ADAM-17 (250 ng) to immobilized AAT detected in the presence or absence of equimolar concentration of polymerized, C-36 fragment, or native AAT (*P = 0.005). Levels of membrane-bound AAT (F) and FcγRIIIb (G) on isolated resting control MM (green) and ZZ-AATD neutrophils before (red) and 2 days after augmentation therapy (blue). (H) The mean chemotactic index of ZZ-AATD neutrophils (black bars) toward sIC (10% v/v) in serum (50% v/v) was significantly reduced on day 2 when compared with day 0 and with day 7 after augmentation therapy. For comparative analysis, MM neutrophils treated with sIC (white bar) were set at a chemotactic index of 1. Immunoblots in A and B are representative results from 1 of 3 separate experiments. B–E and H are the mean ± SEM of triplicate experiments. F and G are representative results from 1 of 3 separate experiments.
We next explored the effect of intravenous augmentation therapy on FcγRIIIb expression in vivo. Two days after treatment, ZZ-AATD (n = 4) patients possessed serum AAT concentrations slightly elevated above MM control serum levels (30.51 ± 1.9 μM and 24.8 ± 0.68 μM AAT, respectively) (Supplemental Figure 5) but also significantly greater than on day 0 (pretreatment) (6.3 μM AAT, P < 0.0001) and 7 days after treatment (13.72 μM AAT, P < 0.0001). Results revealed that infused AAT bound to circulating neutrophils (Figure 6F), as a significant increase in the level of membrane-bound AAT was detected on day 2 after therapy (mean fluorescence, 43) compared with day 0 (mean fluorescence, 17.23; P < 0.001). In addition, downregulation of FcγRIIIb was observed on ZZ compared with MM control cells before therapy (FcγRIIIb mean fluorescence: ZZ-AATD day 0, 16.94; control MM, 29.35; P < 0.001) (Figure 6G). However, on day 2 after treatment, an increase in the level of FcγRIIIb-expressing ZZ neutrophils was observed, comparable to that in MM controls and significantly greater than on day 0 (FcγRIIIb mean fluorescence: ZZ-AATD day 2, 30.41; P < 0.001). A chemotaxis assay toward sIC (10% v/v) in the presence of respective ZZ and MM sera revealed that ZZ neutrophils on day 2 after augmentation therapy had a chemotactic index similar to that of the MM neutrophils and which was significantly reduced compared with day 0 and with day 7 after treatment (P = 0.001 and P = 0.03, respectively) (Figure 6H).
Collectively, these experiments provide evidence for the mechanism by which native AAT modulates neutrophil chemotaxis in response to sIC and illustrate the ability of AAT to bind the circulating neutrophil in vivo and prevent release of FcγRIIIb from the neutrophil membrane, via inhibitory activity against ADAM-17.
AAT regulates IL-8–induced chemotaxis by modulating interaction with CXCR1. To understand the possible mechanism by which AAT inhibits IL-8–induced chemotaxis, we first explored the ability of AAT to modulate ligand binding to CXCR1. The exposure of CXCR1-coated surfaces to IL-8 in the presence or absence of AAT (27.5 μM) revealed that AAT blocked the ability of IL-8 (10 ng) to engage with CXCR1 (Figure 7A). This result implied direct binding of AAT to IL-8, and as IL-8 interacts with glycosaminoglycans with high affinity (24–26), we hypothesized that binding of IL-8 to glycosylated AAT may modulate IL-8–induced chemotaxis. To challenge this hypothesis, we exposed surfaces coated with AAT at a concentration of 27.5 μM to increasing concentrations of IL-8, and the plateau level of maximum binding observed was approximately 20 ng IL-8/cm2 (Figure 7B). In contrast, binding of IL-8 to HSA (control protein) remained linear. The importance of glycosylated moieties of AAT for successful IL-8 binding was confirmed, as nonglycosylated recombinant AAT produced in E. coli failed to bind IL-8 (Figure 7C). The impact of IL-8 binding to AAT or HSA on its ability to stimulate Akt Ser473 phosphorylation, a key step in IL-8–mediated neutrophil chemotaxis (27), was explored. Figure 7D shows that IL-8 (1 ng) stimulated adequate Akt phosphorylation after just 10 minutes and that this effect was unaffected by pre-binding of IL-8 to HSA (27.5 μM). In contrast, pre-binding of IL-8 to AAT (27.5 μM) inhibited IL-8–mediated Akt Ser473 phosphorylation to the level observed after the inclusion of the phosphatidylinositol 3-kinase inhibitor wortmannin. Additionally, binding of IL-8 to AAT (27.5 μM) significantly reduced IL-8 neutrophil Akt phosphorylation in response to both 10 and 20 ng IL-8 by approximately 81% and 62%, respectively (P < 0.05) (Figure 7E). Conversely, however, AAT had no effect on Akt Ser473 phosphorylation stimulated by 40 ng IL-8, illustrating that the described inhibitory effect can be overcome. In addition, an increase in cytoplasmic Ca2+ (calcium flux) is central to the transduction of neutrophil response to IL-8 (28), and changes in average Fluo-4 NW emission in IL-8–stimulated (10 ng) cells was measured over the course of 2 minutes (Figure 7F). IL-8 caused a robust increase in intracellular calcium that reached a peak after 50 seconds, and in comparison the level of intracellular calcium release in the presence of AAT was greatly diminished.
AAT binds IL-8 and modulates neutrophil chemotaxis by controlling CXCR1 binding. (A) Binding of IL-8 (10 ng) to immobilized CXCR1 was inhibited in the presence of AAT (27.5 mM) (*P = 0.002). (B) Comparative binding of IL-8 to AAT and HSA. AAT (27.5 mM) bound approximately 20 ng IL-8. (C) Comparative binding of IL-8 to serum purified AAT and recombinant non-glycosylated (Rec non-glycos) AAT (*P = 0.001). (D) Expression levels of phospho-Akt Ser473 after IL-8 (1 ng/1 × 107 cells) treatment. Wortmannin (Wort, 100 nM) and AAT (27.5 mM), but not HSA, inhibited Akt activity efficiently (*P < 0.05 versus IL-8 control). (E) AAT (27.5 mM) significantly inhibited phosphorylation of Akt induced by 10 ng and 20 ng IL-8 (*P < 0.05 versus IL-8 control). (F) Time course of changes in Ca2+ intensity in response to IL-8 (10 ng) with or without AAT (27.5 mM). (G) AAT suppression of IL-8–induced actin cytoskeletal rearrangements. Immunoblot with anti-actin antibody for the distribution of G-actin (in the supernatant fraction [S]) and F-actin (in the pellet fraction [P]) in untreated or AAT-treated (27.5 mM) cells (top panel). Lower panel: IL-8–treated (10 ng) MM neutrophils with or without AAT or control Wort (100 nM). Distribution ratios are shown. (H) Confocal images of neutrophils (MM) undergoing chemotaxis in response to IL-8 (10 ng). F-actin at the leading edge of neutrophils (arrows) was not apparent in resting cells (Un) or cells treated with IL-8 plus AAT (27.5 mM) (×40 magnification, ×10 zoom). (I) IL-8–induced (10 ng) mean chemotactic index of ZZ-AATD neutrophils (black bars) on day 2 after therapy compared with day 0 and with day 7 after augmentation therapy. All experiments were performed in triplicate on 3 consecutive days, and each measurement is the mean ± SEM. Images in D-H are representative results of 1 of 3 separate experiments.
Remodeling of the actin cytoskeleton is a prerequisite for the chemotactic process, and to analyze the effect of AAT on the redistribution of F- versus G-actin after IL-8 exposure, we employed a biochemical assay to analyze in situ F-actin levels. While AAT (27.5 μM) on its own had no effect, IL-8 (10 ng) induced a substantial change in the ratio of G-actin (supernatant fraction) versus F-actin (pellet fraction) (Figure 7G). The presence of AAT or wortmannin (100 nM) as a control suppressed IL-8–induced F-actin formation. Since coronin is an F-actin–interacting protein solubilized upon neutrophil stimulation (29), we analyzed whether AAT could impact on the solubilization of coronin-1 in response to IL-8. As illustrated in Supplemental Figure 6, AAT (27.5 μM) significantly reduced the level of soluble coronin-1 by 40% (P = 0.01). In addition, polarization of F-actin to the leading edge during neutrophil chemotaxis was greatly diminished in the presence of a physiological concentration of AAT (Figure 7H). Confocal microscopy images illustrated that resting neutrophils had punctuate F-actin staining intensity similar to that of cells treated with IL-8 in the presence of AAT, indicative of sites of contact between cell and substratum (30), rather than polarization. Results also revealed that in response to IL-8 (10 ng), in the presence of respective ZZ and MM sera, ZZ neutrophils on day 2 after augmentation therapy showed a chemotactic index that was similar to the MM controls and was also significantly reduced compared with ZZ neutrophils on day 0 and on day 7 after treatment (P = 0.02 and P = 0.003, respectively) (Figure 7I). Taken together, these data suggest that glycosylated AAT–IL-8 complex formation determines CXCR1 engagement, and in this process AAT functions as a negative regulator of cell chemotaxis.