Apoptosis is the most common form of neutrophil death under both physiological and inflammatory conditions. However, forms of nonapoptotic neutrophil death have also been observed. In the current study, we report that human neutrophils undergo necroptosis after exposure to GM-CSF followed by the ligation of adhesion receptors such as CD44, CD11b, CD18, or CD15. Using a pharmacological approach, we demonstrate the presence of a receptor-interacting protein kinase-3 (RIPK3)—a mixed lineage kinase–like (MLKL) signaling pathway in neutrophils which, following these treatments, first activates p38 MAPK and PI3K, that finally leads to the production of high levels of reactive oxygen species (ROS). All these steps are required for necroptosis to occur. Moreover, we show that MLKL undergoes phosphorylation in neutrophils in vivo under inflammatory conditions. This newly identified necrosis pathway in neutrophils would imply that targeting adhesion molecules could be beneficial for preventing exacerbation of disease in the neutrophilic inflammatory response.
Neutrophils are the most abundant leukocytes in human blood and are essential for innate immune responses against pathogens (1). Apoptosis is the most common physiological cell death in neutrophils and limits tissue damage by preventing the release of histotoxic contents from dying cells (2). However, other types of neutrophil death have also been reported. For instance, in presence of certain proinflammatory cytokines, ligation of Siglec-9 (3), CD44 (4), or FcαRI (5) can induce a caspase-independent neutrophil death. Moreover, community-associated methicillin-resistant Staphylococcus aureus (CA-MRSA) strain USA300 and the Gram-negative bacterium Shigella flexneri have both been reported to trigger necrosis in human neutrophils (6, 7). Thus, neutrophils can undergo both apoptotic and nonapoptotic types of cell death, of which the latter seems to be the more related to inflammatory conditions.
Necroptosis is a form of regulated necrotic cell death which depends on receptor-interacting protein kinase-3 (RIPK3) and mixed lineage kinase–like (MLKL) activity (8, 9). It can be induced by death receptors, IFNs, TLRs, and intracellular RNA and DNA sensors (8, 9). RIPK3 belongs to the kinase family also including RIPK1. For necroptosis initiated by death receptors, both RIPK1 and RIPK3 are required. TNF-α is the most extensively studied extracellular signal that can lead to RIPK1-RIPK3-MLKL-mediated necroptosis. However, necroptosis can also occur in the absence of RIPK1. For example, TLR4 or TLR3, can induce Toll/IL-1 receptor domain-containing adaptor protein-inducing IFN-β (TRIF)-mediated necroptosis which depends on RIPK3, but not RIPK1 (9–11). Similarly, cytosolic DNA-dependent activator of IFN regulatory factors (DAI) can mediate RIPK3-mediated necroptosis in response to viral dsDNA and do not require RIPK1 (9, 12). RIPK1, TRIF, and DAI interact with RIPK3 with the help of a RIP homotypic interaction motif (RHIM) domain, which is essential for necroptosis induction (9). MLKL is a pseudokinase involved in executing cell death in necroptosis downstream of RIPK3 (13–15). An important pathological role of necroptosis has been suggested by a number of genetic studies in mice, in which RIPK3-mediated necroptosis was associated with embryonic lethality and inflammation (8, 9). In contrast, little is known regarding the role of necroptosis in human diseases.
CD44 is a cell-surface protein and plays a role in neutrophil adhesion and transendothelial migration (16). Both CD11b and CD18 are also adhesion molecules expressed on neutrophils. They belong to the integrin family, which are heterodimeric cell surface proteins involved in neutrophil migration and functional responses (17–19). CD15 is a carbohydrate adhesion molecule that is associated with glycoproteins, glycolipids, and proteoglycans (20, 21). As a penta-saccharide, CD15 is abundant in human milk (22). In human leukocytes, CD15 is expressed preferentially on mature neutrophils, monocytes, and all myeloid cells from the promyelocyte stage onwards, for which it serves as a cell surface marker (23–26). Furthermore, through ligation of CD15, neutrophil adhesion to endothelium can be increased (27) and it is believed to play a role in many neutrophil functions, including cell-cell interactions, phagocytosis, degranulation, and respiratory burst (23, 27–31).
We previously reported that CD44 ligation induces a programmed necrosis in GM-CSF-primed, freshly isolated neutrophils (4). Interestingly, such cell death could also be triggered in the absence of GM-CSF if the neutrophils had been isolated from blood of patients suffering from sepsis or from synovial fluids of patients with rheumatoid arthritis, suggesting that CD44 is able to trigger necrosis under inflammatory conditions (4). In the current study, we demonstrate that, besides CD44, ligation of CD11b, CD18, or CD15 can also induce programmed necrosis, suggesting that adhesion can trigger cell death through multiple neutrophil surface receptors under inflammatory conditions. Pharmacological inactivation of RIPK1, RIPK3, and MLKL was able to block adhesion receptor-triggered cell death, suggesting that the programmed necrosis seen under these conditions can be considered to be a necroptosis.
Materials and Methods
Anti-CD44 mAb (clone A3D8) and dihydrorhodamine 123 (DHR) were from Sigma-Aldrich (Buchs, Switzerland). Anti-CD11b mAb (clone CBRM1/5) was purchased from Santa Cruz Biotechnology (Labforce, Muttenz, Switzerland). Anti-CD18 mAb (clone MEM-48) was obtained from Invitrogen (Camarillo, CA). Anti-CD15 mAb (clone W6D3), anti-RIPK1 mAb, and the caspase inhibitor z-VAD-FMK (zVAD) were purchased from BD Biosciences (Erembodegem, Belgium). Anti-CD84 (clone CD84.1.21) and IgG1 isotype control Ab were from AbD Serotec (Düsseldorf, Germany). GM-CSF was supplied by Novartis Pharma (Nürnberg, Germany). Wortmannin, PMA, and diphenylene iodonium (DPI) chloride were purchased from Calbiochem Novabiochem (La Jolla, CA). TGX221 was from Merck (Zug, Switzerland). IC87114 was a kind gift from Dr. Peter Shepherd (University of Auckland, New Zealand). PI103, AS604850, necrostatin-1 (Nec-1), and mito-Tempo were from Enzo Life Sciences (Lausen, Switzerland). Mitoquinone (MitoQ) was obtained from Biotrend Chemical (Zurich, Switzerland). GSK’843 and GSK’872 were from Glixx Laboratories (Southborough, MA) and GW806742X from Adipogen AG (Liestal, Switzerland). Necrosulfonamide (NSA) and PD169316 were obtained from Merck Millipore (Darmstadt, Deutschland). Anti-FAS agonistic mAb (CH11) was purchased from MBL International (Woburn, MA). Rabbit anti-AKT, rabbit anti-phospho-Ser473-AKT, rabbit anti–phospho-Thr334-MK2 (pMK2), rabbit anti-p38, rabbit anti–phospho-Thr180/Tyr182-p38, and rabbit anti-RIPK3 Abs were all from Cell Signaling Technology (Danvers, MA). Rabbit anti-MLKL and rabbit anti-phospho-MLKL (phospho Ser358) mAb were from Abcam (Cambridge, U.K.), and mouse anti-myeloperoxidase (MPO) mAb from Dako (Baar, Switzerland). Anti-MHC class I mAb (clone W6/32) was purchased from BioLegend (San Diego, CA). Anti-GAPDH mAb was obtained from Chemicon International (Chandlers Ford, U.K.). The F(ab′)2 fragments of the secondary goat anti-mouse (GaM) Ab were purchased from Jackson ImmunoResearch Laboratories (Milan Analytica; Roche Diagnostics, Rotkreuz, Switzerland). HRP-coupled secondary Abs (anti-mouse IgG and anti-rabbit IgG) were from GE Healthcare (VWR, Switzerland). Luminata Forte Western HRP substrate was purchased from Millipore Corporation (Billerica, MA). Anti-TNFR1 (mAb225), anti-TNFR2 (mAb226), and anti-TNF-α mAb (AF-210-NA) were from R&D Systems Europe (Abingdon, U.K.). Control F(ab′)2 fragments (anti-CD16, anti-CD32, and anti-CD64) were from Ancell (Bayport, MN).
Peripheral blood neutrophils were purified from healthy normal individuals or chronic granulomatous disease (CGD) patients using Ficoll-Hypaque centrifugation, as described previously (3, 4, 32). The purity of the isolated human neutrophil populations was always >95%, as assessed by staining with Diff-Quik (Baxter, Düdingen, Switzerland) and light microscopic analysis. Written informed consent was obtained from all blood donors, and the Ethics Committee of the Canton Bern approved this study.
Human blood neutrophils were cultured at 1 × 106/ml. Neutrophils were cultured in RPMI 1640 medium plus GlutaMAX (Invitrogen) supplemented with 5% FCS and antibiotics in the presence and absence of GM-CSF (10 ng/ml), anti-FAS (1 μg/ml), PD169316 (10 μM), wortmannin (100 nM), PI103 (100 nM), TGX221 (100 nM), IC87114 (1 μM), AS604850 (1 μM), Nec-1 (25 μM), GSK’843 (50 μM), GSK’872 (10 μM), GW806742X (5 μM), NSA (5 μM), zVAD (20 μM), MitoQ (100 nM), mito-Tempo (100 μM), or DPI (1 μM). GM-CSF stimulation before ligation of adhesion molecules and preincubation with inhibitors were performed for 30 min. Anti-CD44 (6 μg/ml), anti-CD11b (5 μg/ml), anti-CD18 (5 μg/ml), anti-CD15 (5 μg/ml), anti-CD84 (5 μg/ml), or IgG1 (5 μg/ml) mAbs were added for 15 min prior to addition of GaM (20 μg/ml) for receptor ligation. Cells were cultured for the indicated time periods. To block Fc receptors, we used F(ab′)2 fragments of anti-CD16, anti-CD32, and anti-CD64 mAbs which were added 90 min before GM-CSF priming and subsequent receptor ligation. To test for TNF-α-mediated effects, anti-TNFR1 (10 μg/ml), anti-TNFR2 (10 μg/ml), and anti-TNF-α (5 μg/ml) mAbs were added 90 min before GM-CSF priming and subsequent receptor ligation. In other experiments, anti-TNFR1 (10 μg/ml), anti-TNFR2 (10 μg/ml), and anti-TNF-α (5 μg/ml) mAbs were used in combination with GaM (20 μg/ml) to stimulate TNF receptors.
Determination of cell death
Cell death was assessed by uptake of 25 μM ethidium bromide and flow cytometric analysis (FACSCalibur; BD Biosciences) (32, 33). Redistribution of phosphatidylserine (PS) was measured by flow cytometry (4). For morphologic analysis, an Axiovert 35 microscope equipped with a 63×/1.4 NA oil immersion objective lens was used (Carl Zeiss, Jena, Germany).
Enzymatic caspase-3 assay
Neutrophils were cultured under the indicated conditions for 9 h, washed with cold PBS, and subsequently lysed on ice for 10 min in digitonin lysis buffer (20 mM HEPES/KOH [pH 7.4], 100 mM KCl, 100 mM sucrose, 2.5 mM MgCl2) to which 0.0125% digitonin, 1 mM DTT, protease inhibitor mixture (P8340; Sigma-Aldrich), and 1 mM EDTA had been added just prior to use. The caspase-3 activity assay was performed with 10–25 μg of cell lysate in assay buffer (0.1 M HEPES [pH 7.5], 10% sucrose, 0.1% CHAPS, and 10 mM DTT). Substrate (Ac-DEVD-AMC; Bachem AG, Bubendorf, Switzerland) was added to a final concentration of 50 μM before measurement (excitation 360 nm, emission 465 nm).
Oxidative burst measurements
Neutrophils were cultured as indicated and subsequently incubated with 1 μM DHR at 37°C for 30 min, placed on ice, and analyzed by flow cytometry (4). As a positive control, PMA at a final concentration of 65 nM was used for a 15 min stimulation.
Immunoblotting experiments were performed as previously described (4, 32). Briefly, human neutrophils were lysed in Triton lysis buffer (1% Triton X-100, 150 mM NaCl, 50 mM Tris-HCl [pH 7.4], and 1 mM EDTA) containing both protease (protease inhibitor mixture) and phosphatase inhibitors (1 μM okadaic acid, 1 mM Na3VO4, and 5 mM NaF). In case of RIPK1 and RIPK3 detection, neutrophils were lysed with RIPA buffer (50 mM Tris-HCl [pH 8], 150 mM NaCl, 0.5% sodium deoxycholate, 1% Triton X-100, 1% SDS, and 1 mM EDTA; supplemented with protease inhibitor mixture). The lysates were heated at 95°C for 5 min. Proteins were separated by SDS-PAGE and electroblotted onto polyvinylidene difluoride membranes (Immobilion-P; Millipore, Bedford, MA). The membranes were routinely blocked in TBS containing 0.1% Tween-20 and 5% nonfat dry milk, followed by incubation overnight with the indicated Abs at 4°C. The next morning, samples were further incubated with the appropriate HRP-conjugated secondary Ab for 1 h at room temperature. Filters were developed using an ECL technique (ECL-Kit; GE Healthcare) according to the manufacturer’s instructions.
Cells, either untreated or treated, were immobilized on glass slides by cytospin at 2200 rpm for 2 min. Cells were then fixed with 4% paraformaldehyde at room temperature for 10 min and washed with PBS. Following permeabilization with 0.05% saponin, cells were again washed with PBS. After treatment with ice-cold acetone for 15 min at −20°C, immunocytochemistry was performed using the Dako REAL Detection System, Alkaline Phosphatase/RED kit (Baar, Switzerland) according to the manufacturer’s instructions and as previously described (34). Briefly, cells were incubated with rabbit monoclonal anti-human MLKL Ab (phospho S358; 1:250) (35, 36), followed by a biotin-streptavidin system coupled with alkaline phosphatase. The images were visualized using a light microscope (Carl Zeiss).
Double immunofluorescence staining was carried out on 5-μm-thick paraformaldehyde-fixed paraffin-embedded tissue sections from cutaneous vasculitis, ulcerative colitis, and psoriasis patients as previously described (34). Paraffin embedded tissue sections were deparaffinized and rehydrated with graded ethanol dilutions, after which microwave Ag retrieval in 10 mM sodium citrate buffer (pH 6) was carried out. Slides were incubated in blocking buffer (2.5 mg/ml human Igs, 2.5 mg/ml normal goat serum, 2.5 mg/ml BSA in PBS) for 1 h at room temperature and indirect immunostaining was performed using rabbit anti-human MLKL (phospho S358; 1:250) and mouse anti-human MPO (1:100) primary Abs. Tissues were washed with PBS and incubated with Alexa Fluor 555-conjugated GaM and Alexa Fluor 488-conjugated goat anti-rabbit secondary Abs (Thermo Fisher Scientific; distributed by LuBioScience GmbH, Lucerne, Switzerland; both 1:400) for 1 h in the dark at room temperature. Subsequently, samples were washed with PBS (pH 7.4) and nuclei were stained with 1 μg/ml Hoechst 33342. Samples were mounted in ProLong Gold mounting medium (Thermo Fisher Scientific) and examined by confocal microscopy (LSM 700; Carl Zeiss). The Ethics Committee of the Canton Bern approved the retrospective investigation of these tissue samples.
Data were analyzed using GraphPad Prism 5 software (La Jolla, CA). Figures depict mean levels ± SEM. The one-way ANOVA followed by Tukey’s multiple comparisons test or the Kruskal–Wallis test were applied. All p values <0.05 were considered as statistically significant.
Ligation of CD44, CD11b, CD18, or CD15 induces an oxidative burst and subsequent caspase-independent cell death in GM-CSF-primed neutrophils
Ligation of CD44 had already been shown to induce cell death in GM-CSF-primed neutrophils (4). In this study we investigated whether ligation of other adhesion molecules can also result in cell death in GM-CSF-primed neutrophils. CD44 ligation was used as a positive control. For ligation, we used anti-CD11b, anti-CD18, or anti-CD15 mAbs in conjunction with F(ab′)2 fragments of a polyclonal anti-mouse Ab (GaM). Activation of GM-CSF-primed neutrophils through adhesion molecules resulted in the induction of significant cell death, which appeared to be comparable to that seen following CD44 ligation (Fig. 1A). In these experiments, we also investigated anti-FAS mAb which induced neutrophil death as expected. Interestingly, the death fraction induced by FAS ligation was somewhat smaller compared with adhesion-molecule-triggered death in GM-CSF-primed neutrophils (Fig. 1A). In the absence of prior GM-CSF priming, ligation of adhesion molecules had no detectable effect in this system (Fig. 1A). Furthermore, an isotype-matched IgG1 control mAb or an anti-CD84 mAb was also used in conjunction with GaM. Neither of the mAbs had any effect on neutrophil death either in the presence or absence of GM-CSF (Fig. 1A), thus largely excluding the likelihood of Fc receptor-mediated effects and supporting previously published work (3, 4). To exclude directly any possible Fc receptor-mediated effects, Fc receptors were blocked with F(ab′)2 fragments of anti-CD16, anti-CD32, and anti-CD64 mAbs before stimulation. Such pretreatment did not prevent adhesion-molecule-triggered death in GM-CSF-primed neutrophils (Supplemental Fig. 1).
To determine whether the adhesion-molecule-triggered death in GM-CSF-primed neutrophils is owing to apoptosis, we analyzed PS redistribution, which is a characteristic feature of cells undergoing apoptosis (2). No increased PS redistribution was observed compared with untreated neutrophils within a culture period of 4 h, suggesting that the death induced through adhesion receptors following GM-CSF priming is unlikely to be apoptosis. FAS ligation served as a positive control in these experiments (Fig. 1B).
CD44-mediated neutrophil death is dependent on an active NADPH oxidase, since neutrophils from patients with CGD exhibiting a genetic defect in this enzyme did not show a decline in viability following CD44 ligation (4). Therefore, we measured the generation of reactive oxygen species (ROS) following ligation of adhesion molecules in the presence and absence of GM-CSF. Stimulation of GM-CSF-primed neutrophils resulted in high intracellular ROS concentrations within 15 min (Fig. 1C). An isotype-matched IgG1 control mAb and an anti-CD84 mAb served as control Abs in these experiments. In the absence of GM-CSF priming, adhesion molecules had a very limited capacity to trigger ROS. Thus, we observed an association between high ROS production and cell death induction, suggesting that the cell death mediated by adhesion molecules is ROS-dependent. This assumption was supported by data obtained with neutrophils derived from a CGD patient. Such NADPH oxidase-deficient human neutrophils were unable to produce increased amounts of ROS following GM-CSF priming and CD44 ligation or PMA stimulation (Supplemental Fig. 2A). In contrast to normal neutrophils, following GM-CSF priming, they also did not undergo an accelerated cell death after CD44, CD11b, CD18, or CD15 ligation (Supplemental Fig. 2B).
Pharmacological inactivation of caspases blocks apoptosis. We used the pan-caspase inhibitor zVAD to further exclude the possibility that the death mediated by adhesion molecules is apoptosis. In contrast to DPI, which inhibits the NADPH oxidase, zVAD significantly blocked neither the cell death (Fig. 1D) nor intracellular ROS production (Fig. 1E) of GM-CSF-primed neutrophils following ligation of CD44, CD11b, CD18, or CD15. Moreover, the flavoprotein inhibitor MitoQ (37) and the mitochondria-targeted antioxidant mito-Tempo (38) were also unable to block either cell death (Fig. 1D) or ROS production (Fig. 1E), excluding a requirement for mitochondrial ROS in this system. DPI, zVAD, MitoQ, and mito-Tempo alone (without adhesion receptor stimulation) had no effect on ROS production in either the presence or absence of GM-CSF (Supplemental Fig. 2C).
To more precisely investigate the potential involvement of caspases, we specifically measured the enzymatic activity of caspase-3, a key effector caspase in neutrophil apoptosis (2), at 9 h after triggering neutrophil death. Whereas FAS ligation by anti-FAS mAb resulted in strong caspase-3 activation, ligation of adhesion molecules demonstrated either no effect, or a slightly reduced caspase-3 activity in GM-CSF-primed neutrophils (Supplemental Fig. 3A). Morphologically, we observed multiple necrotic neutrophils as a consequence of combined GM-CSF and CD44, CD11b, CD18, or CD15 activation. In contrast, neutrophils activated by anti-FAS mAb exhibited an apoptotic phenotype (Supplemental Fig. 3B). Taken together, these data suggest that the cell death triggered by the ligation of adhesion molecules in GM-CSF-primed neutrophils is caspase-independent. However, it is possible that not all neutrophils are sufficiently activated under these conditions, undergoing a subsequent constitutive apoptosis, explaining the small antideath effect of zVAD in 24 h neutrophil cultures (Fig. 1D).
Cell death of GM-CSF-primed neutrophils following ligation of CD44, CD11b, CD18, or CD15 depends on sequential activation of p38 MAPK and PI3K
To investigate the proximal signaling events initiated by ligation of adhesion molecules in GM-CSF-primed neutrophils, we again used a pharmacological approach. Inhibition of p38 MAPK by PD169316 and PI3K by wortmannin completely blocked ROS production and cell death in such activated neutrophils (Fig. 2A, 2B). Since PI3Ks are usually linked to multiple survival pathways and different classes of PI3K play distinct roles in different signaling pathways (39, 40), we next investigated which PI3K isoform(s) are required for adhesion-molecule-induced neutrophil death with several PI3K subclass inhibitors. We observed that inhibition of class IA PI3Ks (p110α, p110β, p110δ) by pan-class IA inhibitor PI103, but not the inhibition of class IB PI3K p110γ by AS604850, prevented cell death triggered by ligation of adhesion molecules in GM-CSF-primed neutrophils (Fig. 2C). Moreover, the p110β-specific inhibitor TGX221 and the p110δ-specific inhibitor IC87114 both increased neutrophil survival in our system (Fig. 2C). Thus, our findings suggest that several class IA PI3K isoforms transduce death signals initiated by ligation of adhesion molecules.
We next analyzed activation of p38 MAPK and PI3K following ligation of adhesion molecules in GM-CSF-primed neutrophils by immunoblotting. Upon 15 min activation, phosphorylation of MAPK p38, of MAPKAP kinase-2 (MK2; a direct target of p38) (41), and of AKT (an indirect readout for PI3K activity) were dramatically increased compared with unstimulated cells (Fig. 3). Pharmacological inhibition of p38 MAPK by PD169316 and PI3K by wortmannin significantly blocked phosphorylation of the target proteins MK2 and AKT, respectively, demonstrating the efficacy of these two inhibitors. On the other hand, PD169316 did not inhibit phosphorylation of p38 MAPK itself (Fig. 3). A possible explanation is that PD169316 is unable to block phosphorylation of p38 by an upstream kinase; however, it is known to inhibit p38 catalytic activity for downstream target activation (32, 41). Similarly, it has been reported that SB203580, another blocker of p38 MAPK, inhibits p38 activity by binding to the ATP binding pocket, but does not block its own phosphorylation (42).
We observed, moreover, that phosphorylation of p38 MAPK and MK2 were not affected following PI3K inhibition with wortmannin, whereas AKT phosphorylation was abolished as a consequence of p38 MAPK inhibition with PD169316 (Fig. 3). These results suggest that p38 MAPK is proximal to and required for PI3K activation in the neutrophil death pathway initiated by ligation of adhesion molecules in GM-CSF-primed neutrophils.
Cell death of GM-CSF-primed neutrophils following ligation of CD44, CD11b, CD18, or CD15 depends on a RIPK3-MLKL pathway
Earlier work had suggested that CD44 could trigger a necrotic death pathway in GM-CSF-primed neutrophils (4). To investigate whether the caspase-independent neutrophil death triggered by ligation of adhesion molecules following GM-CSF priming involves the RIPK3-MLKL pathway, we investigated the levels of phospho-MLKL (35, 36). Freshly isolated, GM-CSF-stimulated, or anti-FAS mAb-treated human neutrophils exhibited no evidence of MLKL phosphorylation. In contrast, following the combined stimulation with GM-CSF and either CD44, CD11b, CD18, or CD15, but not following cross-linking of MHC class I molecules, we observed strong phospho-MLKL signals within 15 min (Fig. 4A), suggesting activation of the RIPK3-MLKL pathway. Kinetic experiments revealed that phospho-MLKL can be detected as early as 5 min after CD44 activation in GM-CSF-primed neutrophils, while the optimal stimulation time was ∼15 min (Fig. 4B). These data suggest that MLKL phosphorylation is an early event after activation of adhesion molecules in GM-CSF-primed neutrophils. It should be noted that the phospho-MLKL signal was completely lost in necrotic neutrophils (data not shown). Moreover, we observed strong phospho-MLKL signals in infiltrating tissue neutrophils using biopsies from patients with cutaneous vasculitis, ulcerative colitis, and psoriasis (Fig. 4C), suggesting that this pathway is also activated in inflammatory neutrophils under in vivo conditions.
To test whether the RIPK3-MLKL pathway is functionally active and whether the caspase-independent death observed might represent necroptosis, we blocked essential components of the necroptotic pathway. Inhibition of RIPK1 by Nec-1 (43) or RIPK3 by GSK’872 or GSK’843 (44) significantly reduced ROS production (Fig. 5A) and prevented cell death mediated by adhesion molecules in GM-CSF-primed neutrophils (Fig. 5B). Moreover, treatment with GW806742X (45) for pharmacological inactivation of MLKL, an essential effector protein in the necroptotic cell death pathway downstream of RIPK3 (13, 14), reduced ROS production and caused a block in neutrophil death (Fig. 5). On the other hand, NSA, a potential human MLKL inhibitor (13), had no effect on ROS production and subsequent neutrophil death in our system (Fig. 5). Based on the results with Nec-1, GSK’872, and GSK’843 inhibition, we conclude that enzymatic activation of RIPK1 and RIPK3 seems to be required for the caspase-independent cell death induced by adhesion molecules in GM-CSF-primed neutrophils. Although NSA exhibited no efficacy, it is likely that MLKL is also involved in this death pathway.
Since autocrine TNF-α is required for necroptosis induction in some cell lines (46), we next investigated whether this mechanism also applies to the adhesion-molecule-mediated cell death in GM-CSF-primed neutrophils. Addition of a neutralizing anti-TNF-α mAb to the culture medium had no effect in our system (Supplemental Fig. 4A). Moreover, blocking TNFR1 or TNFR2 by anti-TNFR1 or anti-TNFR2 mAb (32) also failed to prevent the death induction by adhesion molecules in GM-CSF-primed neutrophils (Supplemental Fig. 4A). Furthermore, we also investigated the effect of anti-TNFR1, anti-TNFR2, and anti-TNF-α mAb in conjunction with GaM for stimulating human neutrophils; however, these treatments of GM-CSF-primed neutrophils also failed to influence neutrophil viability (Supplemental Fig. 4B). Thus, we obtained no evidence for a role for autocrine TNF-α in the cell death pathway triggered by adhesion molecules in GM-CSF-primed neutrophils.
The RIPK3-MLKL pathway is required for p38 MAPK activation
To examine the possibility that the RIPK3-MLKL pathway is required for activation of p38 MAPK and PI3K following the ligation of adhesion molecules in GM-CSF-primed neutrophils, we again blocked RIPK1, RIPK3, and MLKL using pharmacological inhibitors. The p38 MAPK and PI3K activation were monitored by immunoblotting. Although phosphorylation of p38 was often unaffected, inactivation of RIPK1 and RIPK3 prevented or reduced phosphorylation of MK2, suggesting that p38 activation had been largely blocked (Fig. 6A). Moreover, inhibition of MLKL by GW806742X completely blocked both p38 phosphorylation and its activation (Fig. 6B). It should be noted, however, that Nec-1 treatment in most experiments resulted in only a partial inhibition (Fig. 6A). Expression analyses by immunoblotting revealed that human neutrophils strongly express MLKL and clearly detectable levels of RIPK3. On the other hand, RIPK1 is difficult to detect, suggesting low expression, particularly when compared with Jurkat cells (Fig. 6C). Taken together, one concludes that following ligation of adhesion molecules in GM-CSF-primed neutrophils, the RIPK3-MLKL pathway is required to activate p38 MAPK, which then further transduces the necroptosis signal, involving PI3K and NADPH oxidase.
We have previously shown that neutrophils can undergo a programmed necrosis which requires functional NADPH oxidase and is characterized by cytoplasmic vacuolization, a phenomenon seen both in vitro and in vivo under inflammatory conditions (4). Here, we demonstrate that several adhesion molecules, including CD44, CD11b, CD18, and CD15, can induce such a caspase-independent neutrophil death if cells have been exposed to GM-CSF. These data thus indicate that adhesion of neutrophils can trigger necrosis under inflammatory conditions, explaining why cytoplasmic vacuolization of neutrophils can be observed in inflamed tissues of patients suffering from cystic fibrosis, vasculitis, folliculitis, and psoriasis (4). In this paper, we provide evidence that adhesion molecules can activate the RIPK3-MLKL pathway under inflammatory conditions, both in vitro and in vivo. Moreover, pharmacological inhibition of RIPK1, RIPK3, or MLKL blocked the death pathway triggered by adhesion molecules, suggesting that the neutrophil necrosis observed under these conditions is necroptosis (8, 9). It should be noted that results in this study are novel in that few other investigations have demonstrated necroptosis in primary human cells, especially in the context of a possible role in human disease pathogenesis. So far, evidence for necroptosis in human diseases has been obtained only for epithelial cells of the liver, intestine, and skin (9). On the other hand, we would like to emphasize that most of the experiments performed in this study represent in vitro work. Therefore, additional experimentation using an in vivo model will be required to confirm our findings. Specifically, since not all infiltrating neutrophils seem to undergo necroptosis, it remains unclear which conditions are required to trigger this pathway in inflamed tissues.
Necroptosis can be induced by death receptors, TLRs, and intracellular RNA and DNA sensors (9). Here we provide evidence that the ligation of adhesion molecules can also result in necroptosis, at least in neutrophils. Pharmacological inactivation of RIPK3 with two different inhibitors completely prevented neutrophil necroptosis in each case. Similarly, inhibition of RIPK1 also blocked cell death, but less effectively. It is possible that the death pathway triggered by adhesion molecules depends only partially on RIPK1. This assumption is supported by the low expression of RIPK1 in neutrophils. It should be noted that necroptosis has already been described in the absence of RIPK1 (9–12). In TNFR1-induced necroptosis, the necrosome, defined as the complex containing RIPK1 and RIPK3, plays a key role in the initiation of necroptosis (9, 47). Using neutralizing and blocking Abs, we obtained no evidence that TNF-α mediates adhesion-molecule-triggered necroptosis in GM-CSF-primed neutrophils. However, we cannot exclude the activation of TNFR1 in the absence of its ligand as a consequence of adhesion molecule ligation. Clearly, the mechanism of RIPK3 activation by adhesion molecules in cytokine-primed neutrophils should be further investigated.
In contrast to RIPK3, pharmacological inactivation of MLKL with two different inhibitors revealed contrasting results. GW806742X, a small molecule that binds to the nucleotide binding site in the MLKL pseudokinase domain (45), completely prevented ROS generation and cell death following ligation of adhesion molecules in GM-CSF-primed neutrophils. On the other hand, NSA had no effect in this system. It has been reported that NSA targets the cysteine 86 residue in the N-terminal CC domain of MLKL, but not its kinase-like domain, where MLKL is phosphorylated by RIPK3 at threonine 357 and serine 358 residues; both phosphorylation events are required for necroptosis execution (13). Thus, the differences between GW806742X and NSA in their binding to MLKL may partially explain the differences in efficacy.
Recent studies have suggested that RIPK3-mediated activation of MLKL leads to binding at cellular membranes, thereby triggering ion fluxes (35, 45, 48). However, in neutrophil necroptosis, direct MLKL-mediated plasma membrane disruption does not seem to occur. Instead, MLKL, perhaps in collaboration with RIPK3, was required to activate p38 MAPK leading to PI3K and NADPH oxidase activation. A connection of MLKL to downstream effectors has been suggested. For instance, it has been shown that, in the absence of MLKL, RIPK3 fails to trigger phosphorylation on the mitochondrial protein phosphatase PGMA5S (49). Moreover, overexpression of MLKL was reported to activate JNK (14). MLKL has also been shown to contribute to inflammasome activation (50), further supporting the idea that it may be involved in signaling pathways. Therefore, MLKL should be seen not only as a necroptosis executor protein, but also as an adaptor protein for necrosis signaling. While the crucial roles of p38 and PI3K in the activation of the NADPH oxidase within neutrophil cell death pathways has been reported earlier (4, 32, 51), it remains unclear how the RIPK3/MLKL complex is able to activate p38 MAPK.
The necroptosis pathway described here involves a RIPK3–MLKL–p38 MAPK–PI3K axis, in which all these molecular components are required for the generation of ROS and subsequent necrosis. The activation of NADPH oxidase is absolutely required, since neutrophils from patients suffering from CGD were unable to undergo CD44-mediated necrosis either in the presence or in the absence of GM-CSF (4). In agreement with this earlier study, activation of adhesion molecules in GM-CSF-primed CGD neutrophils did not result in the induction of cell death. Moreover, pharmacological inhibition of the NADPH oxidase with DPI completely prevented neutrophil death triggered by ligation of CD44, CD11b, CD18, or CD15. Therefore, increased ROS production is a requirement for neutrophil necroptosis and occurs downstream of RIPK3-MLKL signaling. Clearly, such high concentrations of ROS may result in irreversible damage to biomolecules and are therefore severely harmful for cell survival. The execution of necrosis appears to involve the permeabilization of granules (4), but the exact mechanism remains to be investigated.
Taken together, we provide evidence for a RIPK3-MLKL signaling pathway in neutrophils leading to necroptosis. This pathway can be activated in GM-CSF-primed neutrophils through several adhesion molecules. MLKL does not seem to act as an executor protein in this pathway, but rather serves to activate more distal signaling molecules required for the production of high ROS levels and subsequent cell death. Characterizing the precise mechanisms of different forms of regulated neutrophil death will likely provide new candidate molecules for targeting in future anti-inflammatory drug treatments.
We are indebted to Dr. Dagmar Simon (Department of Dermatology, University Hospital Bern, Bern) and Dr. Christian Bussmann (Division of Pathology, Histopathology Viollier, Basel) who provided tissue samples from patients.
This work was supported by the Swiss National Science Foundation (Grant 310030_166473 to H.-U.S.). X.W. is a Ph.D. student of the Graduate School of Cellular and Biomedical Sciences at the University of Bern and is supported by the China Scholarship Council. Images were acquired on equipment supported by the Microscopy Imaging Center at the University of Bern.
The online version of this article contains supplemental material.
Abbreviations used in this article:
chronic granulomatous disease
mixed lineage kinase–like
receptor-interacting protein kinase
reactive oxygen species
caspase inhibitor z-VAD-FMK.
The authors have no financial conflicts of interest.