The IRE1α Stress Signaling Axis Is a Key Regulator of Neutrophil Antimicrobial Effector Function

Neutrophils are critical first responders to infection, poised to rapidly kill invading pathogens with a dense array of secretory granules containing a diverse repertoire of antimicrobial molecules (1, 2). Neutrophils can kill intracellular microbes through fusion of secretory granules with phagosomes and generation of reactive oxygen species (ROS). These polymorphonuclear (PMN) leukocytes can also kill extracellular microbes by degranulation, releasing antimicrobial molecules into the extracellular space (3). Upon stimulation, neutrophils can release sticky web-like structures composed of genomic DNA and associated proteins, termed neutrophil extracellular traps (NETs) (4). NET formation is induced by a wide range of microbes, including bacteria, fungi, parasites, and viruses (5) and likely limits spread from the primary site of infection (6, 7). NETs are enriched with neutrophil-derived granule enzymes like neutrophil elastase (NE), myeloperoxidase (MPO), defensins and other antimicrobial peptides (8). NETs are also linked to inflammatory diseases such as systemic lupus erythematosus, psoriasis, or diabetes and exaggerated NET formation or defects in NET clearance can exacerbate these diseases (9, 10). Thus, NET formation must be highly regulated to promote host defense without resulting in pathological inflammatory consequences.

Neutrophils secrete inflammatory mediators, which rely on optimal function of the endoplasmic reticulum (ER). In general, perturbation of ER homeostasis triggers an adaptive stress response, termed the unfolded protein response, and is controlled by three resident ER sensors, of which the inositol-requiring enzyme 1-α (IRE1α) is the most evolutionarily conserved (11). During ER stress, oligomerization of IRE1α triggers autophosphorylation and activation of its cytoplasmic endonuclease domain (12, 13). The endonuclease removes a 26-nucleotide intron from the cytoplasmic Xbp1 mRNA, licensing translation of the transcription factor X-box binding protein-1 (XBP1) (14). XBP1 then induces expression of many genes that improve ER protein folding and degradation to restore ER homeostasis (15, 16). Activation of IRE1α in immune cells stimulates ROS production (17) and enhances production of many proinflammatory cytokines, such as IL-1β, IL-6, and TNF-α (18–20). IRE1α is implicated in inflammation during infection and in many autoimmune diseases such as rheumatoid arthritis and systemic lupus erythematosus, in which neutrophils and NET formation contribute to the progression of these diseases (9, 21).

NET release requires generation of ROS and histone modification to promote decondensation of nuclear chromatin (22, 23). Notably, ROS that contribute to NET formation can be generated by multiple sources including the major phagocyte oxidase, NADPH oxidase 2 (NOX2) and mitochondria (24–26). Concomitantly, histone citrullination by peptidylarginine deiminase-4 (PAD4) reduces net positive charge and decreases histone binding affinity for nuclear DNA, allowing DNA to be expelled from the cell (23, 27–29). PAD4 requires calcium as a cofactor to become active and localizes to the neutrophil nucleus upon stimulation (30, 31). Although IRE1α is known to regulate ROS production in many cell types, how this global stress regulator impacts neutrophil function specifically is poorly understood.

We recently demonstrated that IRE1α aids in clearance of methicillin-resistant Staphylococcus aureus (MRSA) in an s.c. abscess model that is characterized by robust neutrophil recruitment and neutrophil-derived IL-1β (32, 33). In this study, we investigate the contribution of IRE1α to neutrophil antimicrobial function in the context of MRSA infection. We show that MRSA infection triggers neutrophil IRE1α activation and find that this activation is required for efficient neutrophil killing of MRSA, production of IL1-β and NET formation.

All animals were housed and treated in accordance with an Institutional Animal Care and Use Committee–approved protocol (PRO00008690) in Unit for Laboratory Animal Medicine facilities at the University of Michigan Medical School. Blood samples were obtained from healthy adult donors according to the protocol approved by the University of Michigan Medical School (HUM00044257). Written consent was obtained from all donors.

Blood derived from healthy human volunteers was collected into citrate tubes (Becton Dickinson) and layered into Ficoll-Paque Plus (Sigma-Aldrich). Samples were centrifuged at 1440 rpm in a swinging bucket centrifuge for 20 min at 23°C without braking. Pellets containing RBCs were allowed to sediment in a 3% Dextran/saline solution (40 min, room temperature). Supernatants were collected and centrifuged (1600 rpm, 10 min, 4°C). The remaining RBCs in pellets were then lysed with 9 ml of sterile water for 20 s and isotonicity restored by adding 1 ml of 10× HBSS. Samples were centrifuged (1600 rpm, 10 min, 4°C), and pellets containing neutrophils were resuspended in PBS. Neutrophils were counted using an Invitrogen automated cell counter and the purity assessed by flow cytometry using FITC anti-CD16 and allophycocyanin anti-CD15 Abs (Miltenyi Biotec) as markers characteristic of human neutrophils.

Mice bearing Ire1α-floxed alleles (Ire1α^flox/flox) (34, 35) were crossed with S100A8 promoter-driven Cre recombinase (MRP8-Cre) mice (36). The resulting MRP8-Cre⁺Ire1α^WT/flox mice were then backcrossed with Ire1α^flox/flox to generate mice deficient in neutrophil IRE1α (MRP8-Cre⁺Ire1α^flox/flox) and control wild-type (WT) littermates (MRP8-Cre⁻Ire1α^flox/flox). Bone marrow was extracted from mouse femurs and tibias and mechanically dissociated into single-cell suspensions using a 70-μm strainer (Thermo Fisher Scientific). Cells were collected by centrifugation and resuspended in 2 ml of HBSS. Neutrophils were isolated by Percoll gradient as previously described (37). Briefly, bone marrow cells were overlaid on three layers of Percoll (78%, 69%, and 52%) and centrifuged at 1500 × g for 30 min without braking. Cells from the interface of 78% and 69% were collected, washed with PBS, and centrifuged (1600 rpm, 5 min, 4°C). Pellets containing neutrophils were resuspended in medium (RPMI + 10% FBS) and counted using an Invitrogen automated cell counter. Neutrophil purity was assessed by flow cytometry using anti–Ly-6G Ab (BioLegend).

USA300 LAC, a community-associated MRSA strain and its isogenic strain harboring pSarA-sGFP plasmid (MRSA-GFP) (38) were stored at −80°C in LB medium containing 20% glycerol. Bacteria were streaked onto tryptic soy agar (Becton Dickinson) plates and selected colonies grown overnight at 37°C with shaking (240 rpm) in liquid tryptic soy broth (Becton Dickinson). Bacteria were pelleted, washed, and resuspended in PBS. The bacterial inoculum was estimated based on OD₆₀₀ and verified by plating serial dilutions on agar plates to determine CFU. Neutrophils were infected with a multiplicity of infection (MOI) of 10 in RPMI culture medium containing 1% BSA for human neutrophils or 10% FBS for mouse neutrophils for 4 h. For cytokine analysis, samples were treated with lysostaphin (10 U/ml) to kill extracellular bacteria. Supernatants were collected 24 h postinfection (pi) and cytokines quantified by ELISA at the University of Michigan ELISA core.

Neutrophils and cells from dissociated abscess tissue were collected by centrifugation and resuspended in TRIzol reagent (Qiagen). Total RNA was extracted by Direct-zol RNA Kit according to manufacture protocol (Zymo Research) and quantified by NanoDrop. cDNA synthesis was performed using 500 ng of total RNA, murine leukemia virus reverse transcriptase (Invitrogen), and random hexamers (Applied Biosystems). XBP1 transcripts (spliced and unspliced) were amplified by RT-PCR using PCR master mix (Thermo Fisher Scientific) and the following primers: human XBP1 forward (5′-AAACAGAGTAGCAGCTCAGACTGC-3′) or mouse Xbp1 forward (5′-AAACAGAGTAGCAGCGCAGACTGC-3′) and a common reverse (5′-TCCTTCTGGGTAGACCTCTGGGAG-3′). PCR conditions were as follows: step 1, 95°C 2 min; step 2, 35 amplification cycles (95°C 30 s, 60°C 30 s, 72°C 30 s); and step 3, 72°C 10 min. The PCR product was purified and digested with PstI to discriminate between unspliced (290 bp and 183 bp) and spliced (473 bp) for human XBP1. Mouse Xbp1 PCR product is 3 bp longer than human. A hybrid band of spliced and unspliced, especially for mouse Xbp1, is also observed and labeled as Xbp1_H, as previously described (39). Spliced and unspliced DNA fragments were resolved by electrophoresis on a 2.5% agarose gel. Band intensities were measured using ImageJ software, and the percent spliced was calculated using the following formula: [XBP1_S/(XBP1_S + XBP1_U)], which represents the band intensity of the spliced XBP1 relative to total spliced and unspliced band density. Real-time PCR quantification of Xbp1 was performed as previously described using the following primers: spliced Xbp1 forward (5′-GCTGAGTCCGCAGCAGGT-3′), spliced Xbp1 reverse (5′-CAGGGTCCAACTTGTCCAGAAT-3′), GAPDH forward (5′-ACCACAGTCCATGCCATCAC-3′), and GAPDH reverse (5′-TCCACCACCCTGTTGCTGTA-3′) (40).

Neutrophils were seeded in 24-well plates at 1 × 10⁶ cells per well in 500 μl of media per well and infected with MRSA (MOI 10). At 4 h, 100 μl aliquot of cell culture media was mixed with 900 μl of (H₂O + 0.1% NP-40) to lyse neutrophils and release intracellular bacteria. Subsequently, samples were diluted and plated on tryptic soy broth plates. CFUs were enumerated on the next day and used to calculate neutrophils percent killing relative to CFU obtained from control samples where bacteria were inoculated without neutrophils.

Phagocytosis was assessed by flow cytometry. MRSA were fluorescently labeled with biotin-conjugated chicken antiprotein A (Immunology Consultants Laboratory) followed by streptavidin-PE (MRSA-PE). Mouse bone marrow neutrophils were infected with MRSA-PE (MOI 10) for 1 h in the presence or absence of 10% autologous serum. Extracellular bacteria were killed by treating the samples with lysostaphin (10 U/ml). Neutrophils were subjected to flow cytometry, and data were analyzed by FlowJo. Percent of phagocytosis was determined by gating against mock-infected neutrophils.

The initial oxidative burst in human neutrophils was monitored by the reduction of ferricytochrome C, as previously described (41). Neutrophils were incubated in HBSS without phenol red in the presence of 80 μM of ferricytochrome C (equine heart; MilliporeSigma), treated with inhibitors (4μ8C, 25 μM; DPI, 10 μM) or control DMSO and infected with MRSA (MOI 10) for 15 min. Cells were centrifuged, supernatants were collected, and absorbance at 550 nm was measured using a Cytation 5 plate reader. Infection-induced superoxide was calculated by subtracting the absorbance from supernatant collected from untreated cells. The change in absorbance was used to calculate the amount of reduced cytochrome C using the Beer-Lambert law equation with a molecular extinction coefficient of 21.1 mM⁻¹ cm⁻¹.

Neutrophils were first adhered onto polylysine coated no. 1.5 coverslips in six-well plates, treated with inhibitors (4μ8C; 25 μM, NecroX-5; 10 μM, Z-VDVAD-FMK; 10 μM) or control DMSO and infected with MRSA-GFP (MOI 10). Cells were fixed at 4 h pi with 3.7% paraformaldehyde (overnight, 4°C) and permeabilized with PBS+ 0.1% Triton X-100 for 15 min. NE was visualized using rabbit anti-NE (ab68672; Abcam) in a staining buffer (PBS, 0.1% Triton X-100, 5% BSA, and 10% normal goat serum). Goat anti-rabbit secondary Ab conjugated to Alexa-594 was used according to the manufacturer’s procedure (Thermo Fisher Scientific). Coverslips were mounted on microscope slides using Prolong Diamond (Thermo Fisher Scientific). Cells were imaged using an Olympus BX60 microscope. For histology sections, immunofluorescence was performed using rabbit anti-MPO (A0398; Dako) followed by a secondary goat anti-rabbit Ab conjugated to Alexa 488 (Thermo Fisher Scientific). DAPI was used to stain DNA. Images were taken on the Nikon A1 confocal microscope. All fluorescence images were processed and analyzed using ImageJ.

Neutrophils (2 × 10⁵ cells per well) were seeded onto 96-well plates in the presence of cell-impermeable SYTOX Green DNA-Binding Dye (500 nM) (Thermo Fisher Scientific). Cells were left untreated or treated with control solvent, DMSO, or inhibitors (25 μM 4μ8C, 10 μM NecroX-5, 10 μM Z-VDVAD-FMK, and 10 μM KIRA6). Cells were either left uninfected (mock) or infected with MRSA (MOI 10). Fluorescence intensity was measured by the Biotek microplate reader with excitation/emission (485:520), at 4 h pi. NET formation was normalized to SYTOX Green fluorescence intensity obtained from untreated cell control samples.

Neutrophils (2 × 10⁵ cells per well) were seeded onto 96-well and infected with MRSA (MOI 10) in the presence and absence of inhibitors or stimulated with 100 nM PMA for 4 h. Cells were pelleted by centrifugation and culture supernatants were analyzed by MPO/DNA complex ELISA using anti-MPO clone 4A4 (Bio-Rad Laboratories) and anti-DNA POD (Cell Death Detection ELISA; MilliporeSigma) as previously described (42). Levels of MPO/DNA complex were normalized relative to untreated cells culture supernatant media. For measurement of MPO/DNA complex in vivo, s.c. abscesses were mechanically dissociated into single-cell suspension in PBS using 70-μm strainer and supernatants were subjected to ELISA. Levels of MPO/DNA complex in abscess were normalized relative to total MPO.

Flow cytometry analysis was performed using FACSCanto and LSRFortessa Cell Analyzers (BD Biosciences). Total ROS and mitochondrial ROS (mROS; hydrogen peroxide), calcium mobilization, mitochondria membrane potential, CASPASE-2 activity, and CASPASE-3/7 activity were measured using flow cytometry. For ROS measurements, cells were treated with 10 μM CM-H₂DCFDA (Thermo Fisher Scientific), 5 μM CellROX Deep Red (Thermo Fisher Scientific), or 10 μM MitoPY1 (TOCRIS) for 1 h. Cells were washed twice with media and, where indicated, treated for 30 min with inhibitors or control solvent prior to infection with MRSA (MOI 10) or stimulated with 1 μM PMA or 100 μM H₂O₂. For calcium mobilization, neutrophils were loaded with 5 μM Fluo-4 AM (Thermo Fisher Scientific) and incubated at 37°C for 30 min. Cells were treated with inhibitors or control DMSO and the baseline fluorescence is acquired prior to addition of calcium ionophore (5 μM A23187) or infected with MRSA (MOI 10). Fluorescence intensity was recorded over time for 300 s. Data were analyzed by FlowJo software for maximum fluorescence intensity relative to baseline. For other measurements, neutrophils were first treated for 30 min with inhibitors or control solvent and then infected with MRSA (MOI 10) for 4 h. Neutrophils were stained with 2 μM JC1 dye (Thermo Fisher Scientific) for 20 min at 37°C, CASPASE-2 FAM-VDVAD-FMK FLICA substrate (ImmunoChemistry Technologies) for 1 h at 37°C or 2 μM CellEvant CASPASE-3/7 Green detection reagent (Thermo Fisher Scientific) for 30 min at 37°C. Cells stained with CASPASE-2 substrate and JC1 dye were washed twice with PBS prior to flow cytometry analysis. Data were further analyzed with FlowJo software and mean fluorescence intensity (MFI) for each condition was determined as the geometric mean. Ratiometric analysis of red fluorescence (FL2) to green fluorescence (FL1) was used to reflect mitochondrial membrane potential. Percent CASPASE-2⁺ or CASPASE-3/7⁺ cells was determined by gating against mock unstained cells.

Cells were lysed with 1% NP-40 lysis buffer and analyzed by SDS PAGE. Proteins were transferred onto nitrocellulose membrane and blocked with 5% nonfat milk or BSA prior to incubation overnight at 4°C with anti-IRE1α (clone 14C10; Cell Signaling), anti–phospho-p40phox (Thr154) (4311S; Cell Signaling), anti-MPO (A0398; Dako), anticitrullinated histone H3 (ab5103; Abcam), anti-GAPDH (DSHB-hGAPDH-2G7), or anti-GAPDH (clone 0411; Santa Cruz Biotechnology) Abs. Membranes were then incubated with the appropriate secondary Abs for detection.

The s.c. MRSA infections were performed as previously described (43). Male and female WT C57BL/6, Casp2^−/−, MRP8-Cre⁺Ire1α^flox/flox, or MRP8-Cre⁻Ire1α^flox/flox mice were shaved on the right flank. Mice were inoculated with 10⁷ bacteria in 100 µl of PBS s.c. in the shaved area of the skin using a 27-gauge needle. Mice were sacrificed on day 3, and abscesses were excised, weighed, and homogenized in PBS. Total CFU per mouse abscess was enumerated by serial dilution and plating on tryptic soy agar. Cytokines were quantified by ELISA at the University of Michigan ELISA core and converted to picogram per milligram of tissue weight. For histology samples, abscesses were excised on day 2 and fixed in neutral formalin solution. Histology samples were further processed to obtain 5 μm paraffin sections by the Research Histology Core at the University of Michigan Medical School.

Abscesses were excised on day 2 pi and mechanically dissociated into a single-cell suspension using a 70-μM strainer. For blood leukocyte staining, citrated whole blood was collected from mice and RBCs were lysed with RBC lysis buffer (BioLegend). Cells were treated with TruStain FcX (anti-mouse CD16/32) Ab (BioLegend), according to the manufacturer’s instructions, to block Fc receptor binding. Subsequently, cells were stained with the following fluorescent dye conjugated Abs: anti–Ly-6C clone HK1.4, anti–Ly-6G clone 1A8, anti-CD45 clone 30-F11, anti-CD11c clone N418, anti-F4/80 clone BM8, anti-CD3 clone 17A2, and anti-CD45R/B220 clone RA3-6B2 (BioLegend) for 30 minutes on ice. Cells were washed and analyzed on an LSRFortessa Cell Analyzer (BD Biosciences). Data were analyzed with FlowJo software.

Supernatants from abscess-derived single-cell suspensions were collected by centrifugation. The volumes of supernatants were normalized based on total protein concentration. Total DNA was isolated from 200 μl of normalized supernatants using DNeasy Blood and Tissue Kit (Qiagen). Nuclear and mitochondrial DNA in the samples were quantified by real-time PCR using the following primers: 18S forward (5′-TAGAGGGACAAGTGGCGTTC-3′); 18S reverse (5′-CGCTGAGCCAGTCAGTGT-3′); cytochrome C oxidase 1 (MT-CO1) forward (5′-GCCCCAGATATAGCATTCCC-3′); MT-CO1 reverse (5′-GTTCATCCTGTTCCTGCTCC-3′). Nuclear (18S) and mitochondrial (MT-CO1) DNA levels were calculated based on the standard curve that was generated using known DNA concentrations isolated from mouse splenocytes.

Data were statistically analyzed and graphed using GraphPad Prism 7. Significant differences between the two groups were tested using the Mann–Whitney U test. Significant differences between three or more groups were tested using one-way ANOVA and followed up by Tukey multiple comparisons test. Significant differences between two groups with two or more independent variables were tested using two-way ANOVA and followed up by Sidak multiple comparisons test. The mean of at least three independent experiments was presented with error bars showing SD as indicated in the figure legends. The p values <0.05 were considered significant and designated by *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001. All statistically significant comparisons within experimental groups are marked.

To define the role of IRE1α in neutrophil host defense against bacterial infection, we first determined whether infection activates neutrophil IRE1α, using the clinically derived MRSA strain USA300 LAC. Human PMN cells were purified from blood of healthy volunteers as previously described (44), yielding cell populations > 95% CD15⁺CD16⁺, characteristic of neutrophils (Supplemental Fig. 1A). Neutrophils incubated with MRSA for 4 h, increased levels of the spliced variant of XBP1, indicating that IRE1α was activated (Fig. 1A, 1B). The 4 h time point was chosen to allow for accumulation of detectable spliced XBP1. Because it is not possible to acquire or generate primary human neutrophils deficient in IRE1α, we used small molecule inhibitors of the IRE1α endonuclease, 4μ8C or KIRA6 (45, 46), to test the effect of reduced IRE1α activity on neutrophil bactericidal function and IL-1β production. In inhibitor-treated human neutrophils, XBP1 splicing was substantially reduced (Fig. 1B), and these cells were less capable of killing MRSA (Fig. 1C). In addition, inhibitor-treated neutrophils produced lower levels of IL-1β (Fig. 1D) compared with controls. To determine whether these outcomes were due to drug toxicity on MRSA and/or neutrophils, we monitored effects of 4μ8C or KIRA6 on MRSA growth and neutrophil survival. Treatment with 4μ8C or KIRA6 did not affect either axenic growth of MRSA (Supplemental Fig. 1B, 1C) or neutrophil cell viability (Supplemental Fig. 1D, 1E). These data suggest a requirement for IRE1α in neutrophil killing and IL-1β production upon MRSA infection.

Neutrophils form bactericidal NETs during Staphylococcus aureus infection (4, 7). Therefore, we investigated the role of IRE1α in NET formation stimulated by MRSA infection. Neutrophils were cultured on poly-l-lysine–coated coverslips and infected with MRSA-GFP at an MOI of 10 for 4 h in the presence or absence of IRE1α inhibitors. NET formation was analyzed using immunofluorescence microscopy to measure the extracellular colocalization of NE and DNA (Fig. 1E). Uninfected neutrophils predominantly had a multilobed nucleus and intracellular staining of NE, indicating that these cells had not formed NETs. During MRSA infection, NE and DNA were observed external to many cells, of which some had lost the multilobed nuclear morphology, suggesting that these cells had undergone NET formation. When neutrophils were treated with IRE1α inhibitor 4μ8C during MRSA infection, we observed decreased extracellular NE and DNA association compared with DMSO control, indicating that IRE1α contributes to MRSA-induced NET formation. As one approach to quantify NET formation, we measured accumulation of extracellular DNA with the cell-impermeant dye SYTOX Green, which fluoresces upon binding to nucleic acids. MRSA infection stimulated neutrophils to release extracellular DNA, as indicated by increased SYTOX fluorescence intensity, compared with untreated neutrophils (Fig. 1F). Similar to the fluorescence microscopy data, treatment with IRE1α inhibitors reduced SYTOX fluorescence intensity during MRSA infection to a level similar to untreated neutrophils. Although SYTOX Green has been used extensively to monitor NET formation, this assay cannot distinguish between DNA released as NETs and DNA released by other types of cell death. To more specifically evaluate NET formation, we measured the level of extracellular MPO/DNA complexes (Fig. 1G). MRSA infection induced MPO/DNA complex release, which was significantly decreased upon inhibition of IRE1α.

To evaluate the requirement of IRE1α in neutrophil host defense, using a genetic approach, we generated neutrophil-specific IRE1α knockout mice by crossing Ire1α^flox/flox mice (34) with MRP8-Cre-IRES/GFP (MRP8-Cre) mice that express bicistronic Cre recombinase and GFP specifically in neutrophils (36). Analysis of peripheral blood leukocytes for GFP expression showed that MRP8 promoter restricts Cre-GFP expression to neutrophils and to a small percentage of monocytes (Supplemental Fig. 2A). Approximately 40% of neutrophils were GFP negative, suggesting either that the Cre-GFP cassette might only lead to partial deletion in neutrophils or that Cre-GFP might have been expressed and then downregulated. To investigate whether the activity of IRE1α was impaired in MRP8-Cre⁺Ire1α^flox/flox neutrophils, we monitored the level of spliced Xbp1 during MRSA infection or in response to stimulation with the ER stressor, thapsigargin. Neutrophils isolated from mouse bone marrow yielded >80% of Ly-6G⁺ cells regardless of mouse genetic background (Supplemental Fig. 2B). Neutrophils isolated from MRP8-Cre⁺Ire1α^flox/flox mice showed a reduction in spliced Xbp1 in response to MRSA or thapsigargin treatment when compared with WT neutrophils isolated from control littermates (MRP8-Cre⁻Ire1α^flox/flox) (Fig. 1H and Supplemental Fig. 2C, 2D). Consistent with our inhibitor data, IRE1α-deficient neutrophils released significantly less MPO/DNA complex into the extracellular space and killed MRSA with less efficiency but were capable of phagocytosing MRSA similarly when compared with WT neutrophils (Fig. 1I, 1J and Supplemental Fig. 2E). Taken together, our data suggest that IRE1α activation promotes neutrophil NET formation and bactericidal function in response to MRSA infection.

NET formation occurs in an oxidant-dependent manner. The NOX2 complex is a major generator of ROS and is required for induction of NET formation by some triggers (24). However, mROS is also important for NET formation during sterile inflammation and has been associated with NOX2-independent NET formation (25, 26). We recently showed that macrophage IRE1α induces mROS in response to MRSA infection, which is required for effective MRSA killing in phagosomes (17). Thus, we hypothesized that neutrophil IRE1α controls NET formation during MRSA infection via production of mROS. To test this hypothesis, we first monitored production of total ROS and mROS by neutrophils during MRSA infection. Total ROS was measured by flow cytometry using the CM-H₂DCFDA indicator probe, and mitochondrial hydrogen peroxide (mH₂O₂), measured by the MitoPY1 probe (17, 47, 48). MRSA infection increased both total ROS and mH₂O₂ at 4 h pi (Fig. 2A, 2B). When neutrophils were treated with IRE1α inhibitors or the mROS specific scavenger NecroX-5 (49–52), total ROS and mH₂O₂ production decreased. Notably, mH₂O₂ production was profoundly decreased by IRE1α inhibition when compared with total ROS, suggesting that a decrease in total ROS was likely due to decreased mH₂O₂ production. Correspondingly, IRE1α inhibition did not interfere with MRSA-induced early superoxide production, which is initiated in neutrophils via NOX2, nor did IRE1α inhibition interfere with NOX2 activation when assessed by phosphorylation of the NOX2 p40phox subunit (Fig. 2C, 2D) (53). In addition to increased mH₂O₂ production, MRSA infection caused a reduction in mitochondrial membrane potential, which was rescued by pretreatment with IRE1α inhibitors or the mROS scavenger, NecroX-5 (Fig. 2E). We next investigated the contribution of mROS to neutrophil bactericidal activity and NET formation. Pretreating neutrophils with NecroX-5 decreased MRSA killing (Fig. 2F) and suppressed NET formation (Fig. 2G, 2H). It was recently reported that the IRE1α inhibitor 4μ8C possesses antioxidant properties independent of its inhibitory effect on IRE1α (54), which could explain the phenotype that we observed. However, whether 4μ8C reduces cellular ROS accumulation in the absence of IRE1α has not been directly tested. We therefore monitored total ROS and mROS in stimulated IRE1α-deficient macrophages (17) (Supplemental Fig. 3A) in the presence or absence of 4μ8C or N-acetyl-cysteine (NAC; a general ROS scavenger). In contrast to NAC, treatment with 4μ8C did not reduce total ROS (Supplemental Fig. 3B) or mROS (Supplemental Fig. 3C) in IRE1α-deficient cells stimulated with PMA or with H₂O₂, leading us to conclude that 4μ8C does not have antioxidant properties in this context. Thus, our data point to a role for IRE1α activation in promoting neutrophil mH₂O₂ production, which is required for MRSA-induced NET formation and bactericidal function.

Mitochondria-derived molecules such as ATP, mROS, cardiolipin, cytochrome C and oxidized DNA are potent modulators of innate immune pathways (55). We recently found that the IRE1α–CASPASE-2 axis mediates NLRP3 inflammasome activation in Brucella-infected macrophages (19). CASPASE-2 acted upstream of mitochondrial damage, coordinating release of mitochondrial components into the cytosol to initiate activation of the inflammasome. We therefore investigated whether CASPASE-2 played a role in mitochondrial damage in human neutrophils and NET formation during MRSA infection. We monitored CASPASE-2 activation in neutrophils upon MRSA infection using the fluorescent probe FLICA FAM-VDVAD that covalently binds to active CASPASE-2. MRSA infection activated CASPASE-2 in 64% of cells (Fig. 3A). Pretreatment of cells with CASPASE-2 inhibitor (z-VDVAD), 4μ8C or NecroX-5 prior to infection prevented CASPASE-2 activation, suggesting that activity of IRE1α and mROS production is necessary for CASPASE-2 activation in neutrophils and indicates that activation of the IRE1α–CASPASE-2 axis is conserved between neutrophils and macrophages in response to microbial infection. Importantly, CASPASE-2 inhibition alleviated the loss of mitochondrial membrane potential (Fig. 3B), decreased NET formation (Fig. 3C, 3D) and reduced IL-1β production (Fig. 3E) during MRSA infection. CASPASE-2 has previously been implicated in promoting apoptosis through activation of CASPASE-3 (56, 57). A recent study showed that CASPASE-3–mediated apoptosis and NET formation occur concurrently in neutrophils under certain conditions, such as high UV radiation (58). To determine whether CASPASE-3–dependent apoptosis occurs during MRSA infection and whether the IRE1α–CASPASE-2 axis plays a role in this process, we monitored CASPASE-3 activity during MRSA infection with or without inhibitors of IRE1α or CASPASE-2 (Fig. 3F). Neutrophils infected with MRSA exhibited CASPASE-3/7 activity. However, treatment with CASPASE-2 or IRE1α inhibitors did not interfere with CASPASE-3 activation, indicating that IRE1α and CASPASE-2 are not involved in MRSA-induced CASPASE-3 activation. These data suggest that the IRE1α–CASPASE-2 axis mediates NET formation independently of CASPASE-3–mediated apoptosis during MRSA infection.

Histone modification by citrullination is vital for NET formation (29). In neutrophils, histones are citrullinated by PAD4, a process that requires intracellular calcium mobilization (59). To test whether IRE1α and mROS contribute to MRSA-induced histone citrullination and calcium influx, we measured induction of histone citrullination during MRSA infection by immunoblot analysis using an Ab specific for citrullinated histone H3 (Cit-H3). MRSA infection triggered an increase in histone H3 citrullination, which was blocked by pretreatment with IRE1α inhibitors (Fig. 4A). In contrast, treatment with the mROS scavenger, NecroX-5, did not affect histone H3 citrullination triggered by MRSA infection (Fig. 4B). We then monitored calcium flux in neutrophils during MRSA infection using the calcium fluorescent indicator dye, Fluo-4 AM. Kinetic analysis by flow cytometry revealed a sharp, transient increase in Fluo-4 AM fluorescence intensity during MRSA infection, indicating a spike in intracellular calcium (Fig. 4C). This increase was followed by a decline in fluorescence intensity remained above baseline value observed in nonstimulated PMN. Treatment with an IRE1α inhibitor, but not an mROS scavenger, suppressed calcium flux in MRSA-infected cells. Stimulation with a calcium ionophore, A23187, caused a rapid increase in calcium similar to MRSA infection (Fig. 4D). However, the ionophore-induced increase in calcium influx did not decline over time and was independent of IRE1α and mROS. These data indicate that IRE1α is critical for infection-induced calcium mobilization and histone modification and suggest that IRE1α controls NET formation via mROS-dependent and independent mechanisms.

CASPASE-2 has been shown to promote cell death in epithelial cells during S. aureus infection (60) and in macrophages during Salmonella infection (61), which may impact host defenses. But the contribution of CASPASE-2 to antimicrobial host defenses has not been tested. To evaluate the role of CASPASE-2 during MRSA infection in vivo, we used a mouse model of s.c. MRSA infection and monitored inflammation, NET formation and bacterial killing. WT and CASPASE-2–deficient mice (Casp2^−/−) were infected with MRSA (10⁷ CFU) s.c. on the right flank. Bacterial burden and cytokine levels in the abscess were measured at 3 d pi. Compared with WT mice, there were significantly higher bacterial counts (Fig. 5A) and higher TNF-α levels in the abscesses of Casp2^−/− mice (Fig. 5B). This increased level of TNF-α in Casp2^−/− abscesses likely reflect higher bacterial burden. In contrast to WT mice, Casp2^−/− mice had a decreased level of abscess IL-1β (Fig. 5C), which is critical for clearance of S. aureus in a skin and soft-tissue infection model (32). Thus, CASPASE-2 contributes to IL-1β production and is necessary for S. aureus clearance in a mouse s.c. abscess model.

During S. aureus skin infection, NETs are rapidly released by neutrophils and thus prevent the systemic spread of bacteria (7). We next investigated whether NETs could be observed in abscesses from WT and Casp2^−/− mice during MRSA infection, using immunofluorescence microscopy. NET formation was assessed by staining abscess histological sections with DAPI to visualize DNA and for the neutrophil marker MPO (Fig. 5D). In WT mice, we observed extracellular diffuse DNA associated with MPO, consistent with our observation of NET release in WT neutrophils ex vivo. In contrast, DNA in abscesses from Casp2^−/− mice delineated a multilobed morphology more consistent with intact neutrophils. MPO appeared localized predominantly within neutrophil cytoplasm, suggesting that CASPASE-2-deficient neutrophils released fewer NETs during MRSA s.c. infection. To further quantify NET formation in vivo, we used an ELISA to measure the level of MPO/DNA complexes, indicative of NET formation, in dissociated s.c. tissues of infected mice (Fig. 5E) (42, 62, 63). MPO/DNA complex levels were significantly higher in abscesses from WT mice compared with CASPASE-2–deficient mice, suggesting that CASPASE-2 is required for NET formation in vivo during MRSA infection.

To determine the requirement of neutrophil IRE1α in host immunity against MRSA and NET formation in vivo, we infected neutrophil IRE1α-deficient (MRP8-Cre⁺Ire1α^flox/flox) and control littermate (MRP8-Cre⁻Ire1α^flox/flox) mice with MRSA s.c. and measured leukocyte infiltration, levels of spliced Xbp1, bacterial burden, and extracellular DNA and MPO/DNA complexes in infected abscesses (Fig. 6). MRSA infection induced similar levels of neutrophil infiltration in abscesses of both neutrophil IRE1α-deficient and control littermate mice (Fig. 6A). However, the level of spliced Xbp1 in cellular abscesses was reduced in neutrophil IRE1α-deficient mice, indicating that IRE1α activity in neutrophil abscesses was impaired (Fig. 6B). Notably, neutrophil IRE1α-deficient mice were more susceptible to MRSA infection, exhibiting significantly higher bacterial burden in abscesses compared with control littermate mice (Fig. 6C). Furthermore, we observed reduced NET formation in abscesses from the neutrophil IRE1α-deficient mice, indicated by lower levels of extracellular DNA and MPO/DNA complex formation (Fig. 6D, 6E). Collectively, these data support a role for IRE1α and CASPASE-2 signaling in promoting bacterial clearance through controlling neutrophil NET release in vivo.

The IRE1α arm of the ER stress response is integral to development or function of some immune cell types and has been implicated in many inflammatory diseases. Notably, neutrophils can promote inflammatory disease progression and persistence (21, 64, 65), but the role of IRE1α in neutrophils during inflammatory challenge had not been well studied. Our previous work identified IRE1α as being required for innate immunity in an s.c. model of MRSA infection, where neutrophils predominate (32, 33). Our findings in this study clearly demonstrate that IRE1α is a positive regulator of neutrophil effector function during infection. Activation of IRE1α in neutrophils enhanced production of IL-1β and was required for generation of NETs. We found that inhibition of IRE1α impaired mROS generation, which was needed for CASPASE-2 to drive NET formation and IL-1β production ex vivo and in the s.c. abscess. Notably, mROS generation was required for NET formation during sterile inflammation, suggesting a common step regulating NET production in response to both infectious and noninfectious stimuli. Lastly, IRE1α mediated MRSA-induced calcium flux to activate histone citrullination, a key step in the chromatin decondensation that precedes NET formation (23). Our work reveals a central role for the ER stress sensor IRE1α in coordinating neutrophil effector functions.

The ER governs many fundamental processes such as protein synthesis, calcium homeostasis, and lipid metabolism that occur in all cells. Our data highlight IRE1α as a key regulator of cell type-specific processes as well. For example, IRE1α enhances the ability of dendritic cells to process and present Ag to stimulate T cells (66). In the liver, IRE1α signaling enhances hepatic lipogenesis, the process by which dietary carbohydrates are converted into fatty acid–forming triglycerides, independently of its role in the ER stress response (67). In macrophages, IRE1α becomes activated via TLR stimulation augmenting macrophage antimicrobial capacity and production of proinflammatory cytokines (18, 33). In this study, we establish a new role for IRE1α in innate immunity in controlling NET formation, a process that occurs uniquely in neutrophils. These studies emphasize that IRE1α function in particular cell types may encompass both its role in ameliorating ER stress and as an amplifier of signaling, which can impact metabolism and inflammation.

Neutrophils possess lower mitochondrial mass than other cells and mainly rely on glycolysis for ATP production (68), thus the contribution of mitochondria to neutrophil function was not historically appreciated. More recent work has demonstrated that neutrophils contain a complex network of mitochondria and that the activity of this organelle augments many essential neutrophil functions, including chemotaxis, sustained oxidative stress, NF_KB signaling, degranulation, and apoptosis (69–72). Additionally, mitochondrial products such as mROS and NAD⁺ produced through the activity of complex I are required for NET formation during sterile inflammation and in response to other immune signals (26, 73). Our data show an essential role for mitochondrial function in the response of neutrophils to infectious challenge. Collectively, our work, together with recent studies, establish mitochondria as a central regulator of neutrophil function.

Calcium release from ER stores serves as a potent stimulatory signal driving immune activation. Neutrophil stimulation with a calcium ionophore increases cytosolic calcium concentration to induce PAD4-dependent histone citrullination and NET formation (25, 27). The upstream signals that initiate calcium flux and PAD4 activation in neutrophils have not been well defined; however, our findings now point to neutrophil IRE1α as a mechanism to control calcium influx and histone citrullination in response to bacterial infection. Of note, although IRE1α mediated calcium flux specifically in response to MRSA infection, IRE1α activation was not required for ionophore-dependent calcium flux. Calcium can be released from the ER into the cytosol by the activity of the 1,4,5-triphosphate receptor (InsP3R) channels (74), and recent work has implicated InsP3R channels as a mechanism by which IRE1α can mediate flux (75). In that case, IRE1α acted as a scaffold independently of its enzymatic activity to concentrate InsP3R channels at ER-mitochondria contact sites to facilitate calcium release. However, our data suggest that IRE1α endonuclease activity is required for calcium flux during MRSA infection because an inhibitor of the endonuclease domain prevented calcium flux and NET formation. Thus, IRE1α may control calcium flux through multiple mechanisms in a context-dependent manner.

Targeting IRE1α activity with small molecule inhibitors represents a potential therapeutic strategy for treating inflammatory diseases. Our data show that IRE1α augments host defense against bacterial challenge and thus its inhibition could inadvertently increase susceptibility to microbial infection (18, 33). Some evidence in murine models of inflammatory disease, such as rheumatoid arthritis or atherosclerosis, suggests that IRE1α inhibition can counteract the progression of chronic inflammation (20, 76, 77). Because IRE1α contributes to a range of cellular outcomes, such as cell survival, cell death, and inflammation, inhibition of its activity could impact different diseases through diverse mechanisms. In an experimental arthritis mouse model, IRE1α inhibition with 4μ8C attenuated joint inflammation by decreasing production of proinflammatory cytokines such as IL-1β, IL-6, and TNF-α (76). Together, these studies support the investigation of IRE1α inhibitors as possible treatments in human disease with attention to potential susceptibility to infection. Further exploration of the specific mechanisms by which IRE1α controls immunity and inflammation will provide needed molecular context for effective therapeutic strategies.

We thank O’Riordan laboratory members for many helpful discussions. We thank Dr. Karla Passalacqua for helpful comments and reviewing the manuscript. We thank the Microscopy and Image Analysis Laboratory, and the Comprehensive Cancer Center Immunology and Research Histology Cores at the University of Michigan Medical School. We thank Dr. Ling Qi (University of Michigan Medical School) for providing the Ire1α^flox/flox mice.

The authors have no financial conflicts of interest.