ABSTRACT Neutrophil extracellular traps (NETs) are associated with diseases linked to aberrant coagulation. The blood clotting cascade involves a series of proteases, some of which induce

NET formation via a yet unknown mechanism. We hypothesized that this formation involves signaling via a factor Xa (FXa) activation of the protease-activated receptor 2 (PAR2). Our findings

revealed that NETs can be triggered in vitro by enzymatically active proteases and PAR2 agonists. Intravital microscopy of the liver vasculature revealed that both FXa infusion and

activation of endogenous FX promoted NET formation, effects that were prevented by the FXa inhibitor, apixaban. Unlike classical NETs, these protease-induced NETs lacked bactericidal

coagulation and other aseptic conditions. SIMILAR CONTENT BEING VIEWED BY OTHERS NEUTROPHIL EXTRACELLULAR TRAPS IN HOMEOSTASIS AND DISEASE Article Open access 20 September 2024 NEUTROPHILS

CAN PROMOTE CLOTTING VIA FXI AND IMPACT CLOT STRUCTURE VIA NEUTROPHIL EXTRACELLULAR TRAPS IN A DISTINCTIVE MANNER IN VITRO Article Open access 18 January 2021 NEUTROPHIL-DERIVED MIGRASOMES

ARE AN ESSENTIAL PART OF THE COAGULATION SYSTEM Article Open access 12 July 2024 INTRODUCTION Neutrophil extracellular traps (NETs) are chromatin structures decorated with histones and other

proteins that are released by neutrophils as an antibacterial mechanism that plays an important role in innate immunity [1]. Many pathogens and virulence factors are NET inducers, such as

bacterial lipopolysaccharides (LPS) and exotoxins [1,2,3]. However, NETs are also produced during non-infectious diseases, including autoimmune disorders such as rheumatoid arthritis,

psoriasis, systemic lupus erythematosus and arteriosclerosis [4]. Many endogenous NET inducers have been identified, including chemokines (IL-8, CXCL1, CXCL2, CXCL3), interferons (IFN-α,

IFN-γ), anaphylatoxins (C5a) and growth factors (GM-CSF) [5,6,7,8]. Furthermore, alarmins such as β-defensin 1 (hBD-1) [9] and the high mobility group box 1 protein (HMGB1) are known to

trigger NETs [10, 11]. Finally, components of the coagulation system, such as von Willebrand factor (VWF) [12], platelet factor 4 (PF4) [12, 13] and coagulation factor XII (FXII) are also

recognized as inducers of NETs [14]. Microbial proteases are implicated in NET formation [2, 15]. Arginine-specific gingipains secreted by the periodontal pathogen _Porphyromonas gingivalis_

trigger NETs via the activation of protease-activated receptor 2 (PAR2) [2]. This implies that endogenous host proteases may also induce NET, especially in the inflammatory environment,

where their activity is enhanced by neutrophil degranulation, proteolytic cascades (coagulation, fibrinolysis, and kinin generation), and the secretion of proteases from immune and

structural cells [16]. No human proteases are yet known to directly induce NETosis, but many activate PAR-dependent signaling pathways in different cell types and may provide insight into

the endogenous triggers of NETs. The human protease-activated receptors (PAR1–4) belong to the G protein-coupled receptor (GPCR) family [17] and are activated by proteolytic cleavage at a

specific peptide bond in their N-terminal domain. This uncovers a new N-terminal motif that acts as a tethered ligand and interacts with an extracellular domain of the receptor. The

canonical pathway induced by PAR activation engages PLC/Ca2+/PKC endocytosis and endosomal ERK signaling [18, 19]. Alternatively, proteolysis at the N-terminus but away from the canonical

site generates a tethered ligand that does not result in endocytosis, but instead activates adenylyl cyclase and ultimately leads to the activation of PKA [18]. Neutrophils predominantly

express PAR2, whose role in inflammatory diseases has been extensively studied but to the best of our knowledge not in the context of NET formation in response to receptor activation by

endogenous proteases. We therefore investigated human proteases known to signal via PARs, focusing on PAR2 to induce NET formation in the sterile environment of chronic inflammation. Here we

show for the first time the formation of a new category of NETs induced by endogenous proteases via a non-classical mechanism. Our results might explain the accumulation of these structures

under aseptic conditions due to excessive blood clotting associated with inflammation. RESULTS HOST PROTEASES PROMOTE THE FORMATION OF NETS VIA PAR2 ACTIVATION We previously showed that the

_P. gingivalis_ protease gingipain R induces NETs in a PAR2-dependent manner [2]. To determine whether the formation of NETs in response to proteases is a general phenomenon, we tested the

host enzymes trypsin and KLK14. These enzymes signal via PAR2 by removing the extracellular N-terminus at a canonical site (Arg36/Ser37), exposing a tethered ligand in the new N-terminal

sequence (Fig. 1A) [20]. We also applied cathepsin G and neutrophil elastase, which cleave PAR2 at non-canonical sites and thus inactivate the receptor (Fig. 1A) [20]. Using neutrophils that

highly express PAR2 (Fig. 1B, Fig. S1) [21], we found that only the first group of enzymes generated NETs in a concentration-dependent manner (Fig. 1C). The process was prevented by

pre-incubation with protease inhibitors (Fig. 1D) and/or a PAR2 antagonist (Fig. 1E), confirming that proteolytic activity and PAR2 activation are necessary to trigger NETs. PAR2 can also be

activated by small-molecule agonists mimicking the insertion of the tether ligand [22]. We, therefore, confirmed the induction of NETs via PAR2 using the synthetic PAR2 agonists AC 264613

and SLIGRL-NH2, whereas the control peptide with a reversed amino acid sequence (LRGILS-NH2) did not stimulate the release of NETs (Fig. 1F). The formation of NETs in response to proteases

and PAR2 agonists was visualized by confocal microscopy, revealing characteristic extracellular DNA fibers colocalized with elastase when neutrophils were treated with trypsin, KLK14 or

either of the PAR2 agonists, but not with cathepsin G or neutrophil elastase (Fig. 1G). Finally, the role of the PAR2 activation was confirmed using neutrophils from the _PAR2_–/– mouse

strain, which did not respond to trypsin or the PAR2 agonists (Fig. 1H, I). These results show decisively that host proteases promote the formation of NETs in both human and murine

neutrophils via the proteolytic activation of PAR2. COAGULATION FACTOR XA ACTIVATES NETS Coagulation factor Xa is one of the physiological activators of PAR2 (Fig. 2A) [23], which, together

with FVIIa, is considered crucial in extensive crosstalk between coagulation and inflammatory responses [24]. We found, that FXa promotes dose-dependent NET formation (Fig. 2B, C). The

observed effect depends on the proteolytic activity of FXa (Fig. 2D) and is limited in the presence of PAR2 antagonist (Fig. 2E). To determine whether the formation of NETs is induced during

coagulation, we applied two potent activators of FX that induce clotting [25, 26]: Russell viper (_Daboia russelli_) venom (RVV-X), and RgpA, a _P. gingivalis_ cysteine protease (Fig. S2A,

B). Clotting made it impossible to image NET formation in the ex vivo setting, so we can only show that FXa generated by the pre-incubation of FX with RVV-X or RgpA stimulated NET formation

(Fig. S2C, D). Notably, we used 10 nM RgpA, the concentration much lower than needed to induce NETs by direct action on PAR2 (50 nM) [2]. NET formation was prevented by the pre-treatment of

RgpA with Kyt-1, a specific inhibitor of Rgp gingipains, but was unaffected if Kyt-1 was added after the treatment of FX with RgpA (Fig. S2D). These data showed for the first time that FXa

is a potent inducer of NETs and that the exogenous activation of FX leads to the generation of NETs in vitro. Since NET formation stimulated by FXa is associated with enhanced clotting (Fig.

S2E), it may trigger a feed-forward loop of pathological outcomes involved in inflammatory diseases. FXA INDUCES NETS IN VIVO IN THE LIVER MICROCIRCULATION The activation of FX is a key

event in the blood clotting pathway, but excessive FXa activity can trigger many signaling pathways via PARs and other receptors. FXa can thus cause several inflammatory diseases [27], each

of which is associated with NETs generation [4, 28]. The formation of NETs via FXa-dependent PAR2 signaling has not been studied in detail, so we assessed the ability of FXa to induce NETs

in vivo. We used multichannel SD-IVM to visualize extracellular DNA colocalized with elastase and histones in the liver microcirculation following the _i.p_. administration of exogenous FXa.

We then compared the neutrophil response to FXa in the liver microcirculation of wild-type and _PAR2_–/– C57BL6/J mice. FXa generated NETs that were localized in the liver vessels of

wild-type but not PAR2-deficient mice (Fig. 3A). Visualization of the 3D structure of NETs in wild-type mice after FXa treatment revealed neutrophils surrounded by extracellular DNA, along

with neutrophil elastase and histone H2A.X (Fig. 3B). We determined the quantity of NET components relative to the number of neutrophils, which confirmed that the activation of PAR2

following treatment with FXa caused NET formation in the liver microcirculation (Fig. 3C-E). These results show that activation of the coagulation cascade in vivo can lead to the formation

of intravenous NETs and that both FXa and PAR2 are required. INDUCTION OF INTRAVASCULAR COAGULATION IS ASSOCIATED WITH NET FORMATION Although NETs induced by FXa strongly suggest a causative

link between coagulation and NET formation, we were determined to show this link more directly. We, therefore, designed experiments in which coagulation in the liver microcirculation was

induced by the infusion of pure RgpA directly into the liver vessel. The application of RVV-X to trigger intravascular NETs was not possible due to the lethal effect of the venom [29]. In

agreement with our in vitro results (Fig. S2), RgpA triggered NET formation in the liver microcirculation (Fig. 4A-D), and the effect was dependent on gingipain activity because it could be

prevented by pre-incubation of the protease with Kyt-1 (Fig. 4A-D). Given that RgpA directly signals through PAR2 [30] and can cause NETs at higher concentrations [2], we confirmed that

formation of NETs observed in vivo was not caused directly by RgpA but by the FXa generated by RgpA. This was achieved by applying apixaban, a selective inhibitor of FXa that does not affect

gingipain activity (Fig. S3). The formation of NETs was abolished in mice treated with apixaban (Fig. 4E-H), thus confirming that intravascular NETs result from the direct action of FXa.

Furthermore, this result argues against the involvement of plasma kallikrein, activated protein C and thrombin in the coagulation pathway, all generated by RgpA in human plasma [31,32,33]

and all known to signal via PAR2 [22, 34]. Finally, we compared the pattern of NETs formed in response to FXa and LPS because the latter promotes NET formation in liver sinusoids during

endotoxemia [35]. We found that, in contrast to the diffuse pattern of NETs induced by LPS (Fig. 4I), NETs triggered by exogenous or endogenous FXa led to the formation of well-defined but

limited clusters of dense NETs (Fig. 4I), apparently due to blood clotting in the liver vasculature. These results show unambiguously that intravascular coagulation is associated with NET

formation promoted by FXa. BIOCHEMICAL AND FUNCTIONAL CHARACTERIZATION OF PROTEASE-DERIVED NETS Our studies conclusively demonstrated the formation of NETs triggered by proteases in vivo,

but did not answer the question regarding the biochemical nature and biological functions of these structures. To address this question, we have first investigated the signaling pathways

activated by proteases during the formation of NETs. Initially, we examined the activation of MEK/ERK and PI3K-AKT a well-known components of the PAR2 signaling pathways [36] by FXa and AC

264613 (Fig. S4). We confirmed intracellular calcium release following the treatment of neutrophils with FXa, trypsin, and AC264613 (Fig. 5A), a hallmark of PAR2 signaling [20]. Moreover, we

found that NET formation was also dependent on the activation of ERK (Fig. 5B). We have, therefore, shown that the canonical signal transduction pathway mediated by the activation of PAR2

is mechanistically responsible for NETs formation induced by proteases. As biological functions of NETs depend on the composition of proteins decorating the chromatin structure [37], we

performed a detailed proteomic analysis of PAR2-dependent NETs (Fig. 5C). To this end, samples were prepared from neutrophils of three healthy blood donors, stimulated with AC 264613 to

stimulate NET formation or left untreated (control) (Fig. 5D), and subjected to quantitative mass spectrometry analysis. In total 768 proteins were found within a 1% false discovery rate

(FDR) and number of unique peptides ≥2 (Table S1). Principal component analysis (PCA) displays the clustering of replicates and separation between experimental groups (control vs AC 264613)

indicating their different expression (Fig. 5E). Venn diagram shows 34 proteins exclusively quantified in at least one of the three donors after NET induction by AC 264613 compared to the

untreated control (Fig. 5F). Among them are those crucial for endosome trafficking, PAR2 signaling (Rab proteins) and cellular stress response (Fig. 5G). Volcano plot analysis shows

different abundance of 46 proteins; among them, 43 were significantly more abundant after NET induction by AC 264613 (Fig. 5G). Among the top Gene Ontology (GO) biological process terms

enriched were the organization of chromatin and interactions with nucleic acids and proteins (Fig. 5I). Among KEGG pathways are the formation of NETs, autoimmune diseses and carcinogenesis

(Fig. 5I) (DAVID database). Proteomic data confirm the role of PAR2 signaling in NET formation. Then, to further explore the biological functions of protease-derived NETs, we probed them for

antimicrobial immune defense function. Initially, we examined their antibacterial potential by comparing them to PMA-induced NETs. Surprisingly, we found reduced bactericidal activity in

NETs induced by proteases (Fig. 5J). We attributed this result to the absence of the antimicrobial peptide LL-37 from the protease-derived NETs (Fig. 5K) and the low activity of major

antibacterial enzymes, including elastase and cathepsin G, in the protease-derived NETs when compared to PMA-induced NETs (Fig. 5L, M). Collectively, we showed the unique biological

properties of PAR-dependent NETs, proposing that protease-induced NETs should be classified as a novel type of NETs with a potentially fundamental role in regulating aseptic inflammation.

DISCUSSION NETs evolved as an innate immunity mechanism to immobilize and inactivate pathogens [1, 38]. However, the excessive induction of NETs and/or slow clearance by nucleases and

phagocytosis is pathological [39]. This can lead to autoimmune diseases that reflect the breakdown of immunological tolerance to self-antigens massively released from netting granulocytes

[40]. The deposition of NETs also promotes tumor growth and progression, metastasis, and cancer-associated thrombosis [41]. The accumulation of NETs is also associated with blood vessel

occlusion and has been identified in atherosclerotic lesions and arterial thrombi [42]. It is, therefore, important to identify factors that trigger NET formation during aseptic

inflammation. NETs were only recently classified as an important component of the coagulation cascade and are proposed to trigger atherosclerotic plaque formation and arterial thrombosis.

The extruded DNA network binds platelets, enabling their aggregation and activating them via histone H4 [43]. Furthermore, once activated by H4, platelets secrete polyP, which induces

coagulation by FXII [35, 44]. NETs also initiate coagulation by binding to and facilitating the activation of FXII [45] and tissue factor, which initiate the extrinsic (FXa) and intrinsic

(FVIIa) coagulation cascades [46, 47]. Interestingly, the role of NETs in coagulation is bidirectional. Platelets induce the formation of NETs on contact with neutrophils [48] but the

molecular mechanism remained unknown until this study, which revealed for the first time that FXa is a potent inducer of NETs in vitro and in vivo. Moreover, other components of the

coagulation cascade (FVIIa, activated protein C, and thrombin) and the fibrinolysis pathway (plasmin) also activate PAR2 signaling [22, 23] and may also promote NETs. This is supported by

the clusters of NETs visible in the liver circulation, apparently limited to intravenous clots, distinguishing these structures from the diffused NETs induced by LPS (Fig. 4). Finally, we

cannot exclude the possibility that other PARs may be involved. However, PAR1 and PAR4 are expressed at much lower levels than PAR2 on neutrophils (Fig. 1A) and the more abundant PAR3 forms

heterodimers with PAR1 [49, 50]. Therefore, unless PAR3 has an unknown function involving crosstalk with PAR2, we assume that PAR2 signaling triggers protease-induced NETs associated with

coagulation. This novel finding mechanistically explains the link between coagulation and NET formation and confirms the bidirectional mechanism in which coagulation and PAR2-induced NETs

form a feedforward loop. In rheumatoid arthritis, NETs are released into the synovial fluid, where they are found with anti-citrullinated protein antibodies (ACPA). Citrullinated and

carbamylated antigens are implicated in the pathogenesis of rheumatoid arthritis, so in-depth studies of the NET proteome are needed, given that PAR2-activating enzymes are also found in the

synovium [51]. These include coagulation factors and mast cell tryptase, which exacerbates synovitis in rheumatoid arthritis and osteoarthritis by signaling via PAR2 [52, 53]. Activation of

the coagulation pathway by gingipains may also be relevant because _P. gingivalis_, the keystone pathogen in periodontitis, is considered the etiological factor that promotes rheumatoid

arthritis [54]. Many of the proteases present in tumors activate PAR signaling, including elevated levels of KLK14 in tumors and inflammatory skin diseases such as the Netherton syndrome

[55,56,57]. Moreover, the tumorigenic role of PAR2 in response to trypsin has been demonstrated in colon cancer, ovarian cancer, breast cancer, and colorectal cancer [58,59,60,61]. FXa

promotes tumor migration and invasion in breast cancer [62]. Importantly, neutrophils are key innate immune effector cells in cancer, explaining the presence of NETs in Ewing’s sarcoma [63],

breast cancer [64], pancreatic cancer [65], colorectal cancer [66] and ovarian cancer [67], but their role is not clearly defined. NETs inhibit the proliferation of colon cancer cells and

have a cytotoxic effect against melanoma cells [68, 69]. However, they also capture and protect circulating cancer cells from NK cells and CD8+ T cells [70, 71]. NETs also promote the

awakening of dormant cancer cells [72]. Furthermore, NET-associated proteases promote the remodeling of the extracellular matrix. Accordingly, our results may lead to new research focusing

on proteases in the cancer microenvironment that act as NETs inducers, revealing their yet undiscovered role in cancer development and progression. Justification for this research is

suggested by the proteome of PAR2-induced NETs. In summary, we have discovered a new category of non-classical, endogenous protease-triggered NETs, providing insight into the etiology of

non-infectious inflammatory diseases, including those related to coagulation, but also autoimmune diseases and carcinogenesis (graphical abstract). Our results provide a broader view of the

undoubtedly bidirectional relationship between NETs and coagulation. Most importantly, the new category of NETs reported here for the first time may be considered therapeutic targets for

protease inhibitors and/or PAR2 antagonists. MATERIALS AND METHODS ISOLATION OF HUMAN NEUTROPHILS A fraction enriched in granulocytes was obtained by density gradient centrifugation in

lymphocyte separation medium (Pan Biotech). Neutrophils were separated from erythrocytes using 1% polyvinyl alcohol (POCH). After 30 min of sedimentation, the upper layer was harvested and

centrifuged (280 × g, 10 min, room temperature) and the residual erythrocytes were lysed in water. Neutrophils were resuspended in serum-free Dulbecco’s modified Eagle’s medium (DMEM)

without phenol red (Gibco/Thermo Fisher Scientific). The purity of the human neutrophils was assessed by flow cytometry analysis. Cells were incubated in 0,5% BSA in PBS 1x with addition of

PerCP-Cy™5.5 Mouse Anti-Human CD14 Clone MφP9 (BD Pharmingen, cat. no. 562692), FITC anti-human CD15 (SSEA-1) Antibody, Clone HI98 (BioLegend, cat. no. 301904), or CD3 Monoclonal Antibody

(UCHT1), APC (eBioscience™, cat. no. 17-0038-42) in 1:100 dilution for 30 minutes in 4 °C. Data was acquired by the BD LSR Fortessa system (Becton Dickinson) and analyzed by BDFACS Diva

sofware. The data are presented in the Fig. S5A. SEX AS A BIOLOGICAL VARIABLE Our study examined male and female animals, and similar findings are reported for both sexes. The animals were

allocated in experimental groups randomly, within a given phenotype. ISOLATION OF MURINE PERITONEAL NEUTROPHILS C57BL6/J wild-type and PAR2 knockout (_PAR2_–/–) mice, 6–8 weeks old, were

injected intraperitoneally (_i.p_.) with 1 ml of 4% sterile thioglycolate (Fluka) to induce peritonitis. After 3 h, peritoneal lavage was performed with 10 ml ice-cold phosphate-buffered

saline (PBS; Gibco/Thermo Fisher Scientific). The resulting cell suspension was centrifuged (280 × g, 5 min, room temperature) and the retained erythrocytes were lysed in water. The

neutrophils were resuspended in serum-free DMEM without phenol red. The purity of the peritoneal neutrophils was assessed by flow cytometry analysis (Fig. S5B). Cells were incubated in 0,5%

BSA in PBS 1x with addition of FITC-conjugated rat anti-mouse Ly-G6 antibody (ThermoFisher Scientific, Invitrogen, cat. no. 1-9668) in 1:200 dilution for 30 minutes in room temperature. Data

was acquired by the BD LSR Fortessa system (Becton Dickinson) and analyzed by FlowJo v10 software. ISOLATION OF MURINE BONE MARROW-DERIVED NEUTROPHILS Bone marrow neutrophils were isolated

from 8-week-old wild-type C57BL6/J mice. The isolated femur and tibia were centrifuged (10 000 × g, 40 sec, RT), cells were resuspended in DMEM supplemented with 10% FBS and 1%

penicillin/streptomycin and passed through a 40 μm nylon cell strainer. The single cell suspension was centrifuged (350 × g, 5 min, RT), lysed in 0.155 M NH4Cl to remove erythrocytes, and

centrifuged (350 × g, 5 min, RT) after addition of ice-cold PBS. Collected cells were resuspended in DMEM and passed through a 70 μm nylon cell strainer and centrifuged (350 × g, 5 min, RT).

Cells were resuspended in ice-cold PBS 1x supplemented with 2% FBS and 1 mM EDTA. Bone marrow cells were separated using the double gradient with Histopaque 1119 and 1077 (Sigma-Aldrich).

After centrifugation (700 × g, 30 min, RT) bottom layer enriched in neutrophils was collected and centrifuged again (350 × g, 10 min, RT). The purity of the bone marrow-derived neutrophils

was assessed by flow cytometry analysis (Fig. S5C). Cells were incubated in 0,5% BSA in PBS 1x with addition of FITC-conjugated rat anti-mouse Ly-G6 antibody (ThermoFisher Scientific,

Invitrogen, cat. no. 1-9668) and anti-mouse CD11b-PE (ThermoFisher Scientific, Invitrogen, catalog numer 12-0112-82) in 1:200 dilution for 30 minutes in room temperature. Data was acquired

by the BD LSR Fortessa system (Becton Dickinson) and analyzed by BDFACS Diva software. QUANTITATIVE REVERSE-TRANSCRIPTION POLYMERASE CHAIN REACTION (QRT-PCR) Total cellular RNA was extracted

from human peripheral neutrophils using TRIzol reagent (Invitrogen/Thermo Fisher Scientific). Briefly, 800 ng of RNA was reverse transcribed with MultiScribe Reverse Transcriptase (Applied

Biosystems) in a total volume of 20 μl according to the manufacturer’s instructions. We then amplified 20 ng of the resulting complementary DNA (cDNA) in a 15-μl reaction containing 10 mM of

each primer (Table S2) and 1× GoTaq PCR master mix (Promega). The templates were denatured at 95 °C for 5 min followed by 40 amplification cycles (Table S2) and a final elongation step at

72 °C for 10 min. Differences in gene expression were determined using the ΔΔCT method [73] and normalized against the housekeeping gene _EF-2_. PAR2 PROTEIN EXPRESSION ON THE SURFACE OF

HUMAN AND MOUSE NEUTROPHILS The cells (0,2 mln) were left untreated (human and mouse bone marrow-derived PMN), fixed with 4% formaldehyde (5 min) and washed with PBS 1x. Cells were then

permeabilized with 0,1% PBS-TritonX-100 for 3 minutes. After washing with PBS 1x, neutrophils were incubated in 0,5% BSA containing 5 μg/ml anti-PAR2 antibody (ab180953, abcam) for 30 

minutes at 20 °C. Goat anti rabbit IgG conjugated with APC (cat. no. 111-136-144, 1:1000 dilution, 30 minutes, 20 °C) was used as secondary antibody. Additionally, surface of cells were

quenched with trypan blue (0.83 mg/ml) immediatelly before measurment. Unstained cells were used as autofluorescence control (AF). Cells without incubaction with primary antibody were used

as control of secondary antibody (2nd Ab). Data were acquired by the BD LSR Fortessa system (Becton Dickinson) and analyzed by BDFACS Diva software. INDUCTION OF NETS Neutrophils (1 × 105

per well) were seeded in 96-well plates coated with 0.01 mg/ml poly-l-lysine (Sigma-Aldrich) and incubated for 30 min at 37 °C in a humidified 5% CO2 atmosphere to promote cell adhesion. NET

formation was stimulated by exposure to proteases, including bovine pancreatic trypsin (Sigma-Aldrich), kallikrein 14 (KLK14; kindly provided by Dr. T. Kantyka), FX, FXa, FXa EGR (Thermo

Fisher Scientific), cathepsin G (BioCentrum), neutrophil elastase (Athens Research & Technology) and/or RgpA. We also included the protease inhibitors aprotinin (Sigma-Aldrich) and/or

Kyt-1 (Peptide Institute). The production of NETs was also triggered using the PAR2 agonists AC 264613 and/or SLIGRL-NH2 (Tocris Bioscience). As a negative control, we used the reversed

amino acid sequence peptide LRGILS-NH2 (Tocris Bioscience). To block PAR2-dependent signaling, neutrophils were pre-treated with the peptide PAR2 antagonist FSLLRY-NH2 (Tocris Bioscience)

before NETs were induced with proteases. All treatments lasted 3 h. To investigate ERK signaling during PAR2-activated NETs, neutrophils were pre-treated for 30 min with the selective ERK

inhibitor UO126 (Cell Signaling Technology). DNA QUANTIFICATION The total DNA content of NETs was determined by incubating neutrophils with 1 U/ml micrococcal nuclease (Thermo Fisher

Scientific) for 15 min to release NETs from the cells. The free NETs were then separated from cells and debris by centrifugation (1800 × g, 10 min, room temperature) and the DNA was

quantified using 10 μM SytoxGreen nucleic acid stain (Invitrogen) at a 1:10 (v/v) ratio. The fluorescence signal was obtained by excitation at 485 nm (emission wavelength 535 nM) and is

presented as relative fluorescence units (RFU). IMMUNOFLUORESCENCE STAINING Slides coated with poly-l-lysine were seeded with 5 × 105 human or mouse neutrophils and stimulated with PAR2

agonists and proteases for 3 h, as described above. The cells were then fixed for 10 min with 3.7% formaldehyde, blocked with PBS containing 5% fetal bovine serum (FBS), 1% bovine serum

albumin (BSA), 0.05% Tween-20 and 2 mM EDTA for 1 h, and incubated with 0.1% saponin (Sigma-Aldrich) in PBS for 30 min. The cells were stained with rabbit anti-human neutrophil elastase

antibodies (Athens Research and Technology, cat. no. 16-14-051200) for 1 h and goat anti-rabbit IgG F(ab′)2 antibodies conjugated to APC (Jackson ImmunoResearch Laboratories, cat. no.

111-136-144) for 45 min. The antibodies were suspended in PBS containing 3% BSA and 0.1% saponin. Nuclei were counterstained with Hoechst 33342 (1 μg/ml) for 10 min. All steps were carried

out at room temperature with intervening washes (0.1% saponin in PBS). Confocal images were captured using a Zeiss LSM 880 microscope and ZEN software. PROTEOLYTIC ENZYMES Arginine-specific

gingipain (RgpA) was purified, followed by active-site titration as previously described [74]. TREATMENT OF MICE PRIOR TO INTRAVITAL MICROSCOPY Experimental groups, with 3 mice in each group

(6–8 weeks old), consisted of C57BL6/J wild-type and _PAR2_–/– mice that received native mouse FXa (0.06 mg/kg; abcam) or PBS (FXa control group) via cannulated jugular vein 4 h before NET

imaging. Wild-type mice were injected (i) _i.v_. with RgpA (0.8 mg/kg) 4 h before NET imaging, (ii) _i.v_. with RgpA and Kyt-1 (2.75 μg/kg) 4 h before NET imaging, (iii) _i.v_. with RgpA and

also _i.p_. with apixaban (Medchem Express; 25 mg/kg) or DMSO (apixaban control group) 30 min prior to RgpA injection, or (iv) _i.p_. with 1 mg/kg LPS (_Escherichia coli_ serotype 0111:B4;

Sigma-Aldrich) in saline to induce endotoxemia [75] followed by intravital imaging 4 h after LPS injection. PREPARATION OF MOUSE LIVER FOR INTRAVITAL MICROSCOPY Mice were anesthetized by

_i.p_. injection with a mixture of ketamine hydrochloride (200 mg/kg; Biowet Pulawy) and xylazine hydrochloride (10 mg/kg; aniMedica) and were cannulated in the right jugular vein to

facilitate the supply of anesthetics, antibodies and fluorescent dyes. Livers were prepared as previously described [76]. Briefly, the liver was exposed by a midline incision of the abdomen,

followed by a lateral incision along the costal margin to the midaxillary line. The mouse was then placed in the right lateral position, and the ligaments attaching the liver to the stomach

and diaphragm were cut, allowing the liver to be moved onto an imaging board covered with saline-soaked Kimwipes tissue. A cover glass was placed on the left liver lobe, and the space

underneath the cover glass was filled with saline to keep the tissue moist. The mouse was then placed under the upright microscope for intravital imaging. SPINNING-DISK INTRAVITAL MICROSCOPY

(SD-IVM) Livers were imaged using a Zeiss Axio Examiner. Z1 upright microscope equipped with an AMH-200-F6S metal halide light source (Andor, Oxford Instruments) with a motorized

six-position excitation filter wheel and a DSD2 laser-free confocal spinning disk (Andor, Oxford Instruments) with Zeiss EC Plan-NEOFLUAR 10×/0.3 and Zeiss EC Plan-NEOFLUAR 20×/0.5 air

objectives. The following excitation filters were used: DAPI, 390/40 nm; GFP, 482/18 nm; RFP, 561/14 nm; Cy5, 640/14 nm. These were paired with the corresponding emission filters: DAPI,

452/45 nm; GFP, 525/45 nm; RFP, 609/54 nm; Cy5, 676/29 nm. Images were captured using a Zyla 5.5 sCMOS camera (5.5 megapixels; Andor, Oxford Instruments) and iQ v3.6.1 acquisition software

(Andor, Oxford Instruments). VISUALIZATION OF NEUTROPHILS AND NETS NETs were visualized by the co-localization of neutrophil elastase, histone H2A.X and extracellular DNA. These were

detected using an AlexaFluor 647-conjugated anti-neutrophil elastase monoclonal antibody (clone G-2; Santa Cruz Biotechnology; cat. no. sc-55549 AF647; 1.2 µg per mouse), an AlexaFluor

568-conjugated anti-H2A.X monoclonal antibody (clone 938CT5.1.1; Santa Cruz Biotechnology; cat. no. sc-517336; 0.8 µg per mouse) and 0.1 mM SytoxGreen in saline, respectively. All antibodies

were injected _i.v_. via the cannulated jugular vein ~20 min before IVM. SytoxGreen stains DNA instantly, and was administrated during imaging. NETs were quantified as previously described

[77] using ImageJ v1.53e (National Institutes of Health) and expressed as the percentage of liver area covered in each field of view, with at least five fields analyzed per mouse.

BACTERICIDAL ACTIVITY OF NETS Neutrophils seeded in 24-well plates (2 × 106/well) coated with 0.01 mg/ml poly-l-lysine were centrifuged (200 × g, 5 min, room temperature) before we added 100

 µM AC 264613 and 1 µM FXa. We used 0.025 µM phorbol 12-myristate 13-acetate (PMA) to induce classical NETs. After 3 h, NETs were collected and incubated with 4 × 106 _E. coli_ ATCC 29522 or

_E. coli_ DH5α cells. As a control, the same bacterial strains were incubated in supernatant from untreated neutrophils. After 2 h, bacterial survival was estimated by plating serial

dilutions on solid agar plates and counting the colony forming units (CFUs). ENZYMATIC ASSAYS NETs generated in response to PMA (0.025 µM), AC 264613 (100 µM), trypsin (0.05 µM) and/or FXa

(1 µM) were collected, and the activities of neutrophil serine proteases were measured using specific substrates. Neutrophil elastase and cathepsin G activities were determined using

_N_-methoxysuccinyl-Ala-Ala-Pro-Val-_p_-nitroanilide (Sigma-Aldrich) and _N_-succinyl-Ala-Ala-Pro-Phe-_p_-nitroanilide (Sigma-Aldrich), respectively. Each substrate (1 mM in 100 μl 50 mM

Tris-HCl, pH 7.5) was mixed with 100 μl of supernatant from the netting and control neutrophils, and the rate of substrate hydrolysis was measured as the increase in the optical density at

450 nm (OD450) after incubation for 30 min at 37 °C. SDS-PAGE AND IMMUNOBLOTTING NETs and whole protein lysates were obtained from neutrophils by using Pierce RIPA Buffer (Thermo Scientific)

with proteinase and/or phosphatase inhibitors and equal amounts of protein were separated by SDS-PAGE. The proteins from NETs and lysates for AKT were transferred to PVDF membranes (Merck

Millipore) in 25 mM Tris-HCl, 0.2 M glycine (pH 8.3) supplemented with 20% methanol (60 V, 3 h, 4 °C) or to nitrocellulose membrane for ERK. Non-specific binding sites were blocked with 5%

skimmed milk for PVDF or 5% BSA for nitrocellulose membrane in Tris-buffered saline (pH 7.5) containing Tween-20 (TTBS) for 1 h at room temperature, followed by overnight incubation at 4 °C

with antibodies. Mouse LL-37/CAP-18 (1:500, Hycult Biotech, cat. no. mAb 3D11), PathScan® Multiplex Western Cocktail I (rabbit P-ERK1/2, Thr202/Tyr204 (1:1000, Cell Signaling, cat. no.

5301), rabbit ERK1/2 (1:500, Cell Signaling, cat. no. 9102), rabbit Rab 11 (1:1000, Cell Signaling, cat. no. 3539)), rabbit P-AKT (T308) (1:1000, Cell Signalling, cat. no. 13038), rabbit AKT

(1:1000, Cell Signalling, cat. no. 9272), rabbit GAPDH (1:5000, Cell Signaling, cat. no. 2118) were used in 5% skimmed milk or 3% BSA in TTBS, respectively for PVDF and nitrocellulose

membrane. Membranes were washed extensively in TTBS and incubated with a 1:20000 dilution of a horseradish peroxidase (HRP)-conjugated sheep anti-mouse IgG secondary antibody (Sigma-Aldrich,

cat. no. AC111P) and with goat anti-rabbit IgG (1:5000; Cell Signaling, cat. no. 7074) for 1 h in TTBS containing 5% skimmed milk or 3% BSA in TTBS. Membranes were washed (5 × 5 min) in

TTBS, and blots were developed using enhanced chemiluminescence (ECL) substrate (Thermo Fisher Scientific). INTRACELLULAR CALCIUM MEASUREMENT Neutrophils were seeded at a density of 0.5 × 

106 cells/well in black 96-well plates with clear bottoms (coated with 0.01 mg/ml poly-l-lysine) and were incubated with 80 μl 1× calcium dye 5 in 1× Hanks’ balanced salt solution (HBSS)

supplemented with 20 mM _N_-2-hydroxyethylpiperazine-_N_′-2-ethanesulfonic acid (HEPES) in the presence of 2 mM probenecid (Molecular Devices). PAR2 activators (40 μl/well) were added using

the Flex Station 3 multimode microplate reader (Molecular Devices), and fluorescence readings were acquired for 120 s at an excitation wavelength of 485 nm (emission wavelength 525 nm). The

final concentration of the compounds was 100 μM AC 264613, 0.25 μM trypsin, and 1 μM FXa. Background fluorescence was recorded in untreated cells in the same assay buffer. MASS SPECTROMETRY

(LC-MS/MS) ANALYSIS OF NETS PROTEOME Neutrophils were subjected to AC 264613 or untreated (control) for 3 hours, then treated with MNase (1 U/ml) for 15 minutes and centrifuged at 1800 × g

for 10 minutes. Ice-cold acetone was added to the supernatants containing NETs, then samples were incubated at −80 °C for 60 minutes and centrifuged at 15,000 × g for 10 minutes. The

supernatant obtained by centrifugation was withdrawn, the protein pellet was dried and dissolved in PBS. Then, samples were lyophilized in a speed vacuum concentrator before resuspension in

8 M urea, 100 mM ammonium bicarbonate. Proteins were reduced with 10 mM dithiothretiol for 30 min, followed by alkylation with 30 mM iodoacetamide for 30 min, in the dark. The urea

concentration was lowered by the addition of 100 mM ammonium bicarbonate and then digested with trypsin at a 1:50 (w/w) ratio overnight at 37 °C. Digested peptides were acidified with formic

acid and desalted on homemade reverse-phase C18 columns packed with Octadecyl C18 Solid Phase Extraction disks (Empore, 3 M). The samples were eluted from the columns with 70% acetonitrile

in 0.1% formic acid, lyophilized to near-dryness and redissolved in 0.1% formic acid. The samples were analyzed on an Orbitrap Eclipse Tribrid mass spectrometer (Thermo Fisher Scientific)

coupled to an EASY-nLC 1200 (Thermo Fisher Scientific). 750 ng of each sample was loaded onto a trap column (2 cm × 75 µm inner diameter) and separated on an analytical column (20 cm × 75 µm

inner diameter) packed with 1.9 µm C18 beads (Dr. Maisch, GmbH). The samples were eluted at a flow rate of 250 nl/min and with a gradient from 5 to 35% acetonitrile for 80 min, followed by

a steep increase to 80% acetonitrile for 10 min. The MS raw files were processed in Proteome Discoverer 2.5 using the Sequest HT search algorithm. The UniProt human reference proteome was

used as a database with the following parameters: 10 ppm precursor mass accuracy, 0.02 Da fragment mass accuracy, trypsin as digestion enzyme, max. 2 missed cleavage sites, Oxidation (M) as

variable modification and Acetyl (N-Term), Met-loss (M), and Met-loss+Acetyl (M) as variable protein N-terminus modifications. Carbamidomethyl (C) as fixed modification. Protein abundances

were label-free quantified based on precursor area and using only unique peptides. The samples were normalized to the total peptide amount. Further data analysis was conducted in Perseus

(www.perseus-framework.org) and MATLAB (R2022b) by MathWorks. Data in Venn diagram were filtered to proteins identified with a 1% FDR and minimum two unique peptides. Data in Volcano plot

were filtered to include proteins identified with a 1% FDR with a minimum of two unique peptides and quantified in all three replicates in at least one sample group. Protein abundances were

log2 transformed before further processing. Missing values were drawn at random from a normal distribution downshifted by 1.8 and with a width of 0.3 compared to the experimental data.

_P_-values were calculated with an unpaired equal variance t-test and FDR adjusted using the Benjamini-Hochberg procedure. Proteins were considered differential abundant at a fold change ≥2

and a BH adj. _P_-value < 0.05. BIOINFORMATICS ANALYSIS Principal component analysis (PCA) was performed using Python 3.12 (Python Software Foundation - PSF). The components were selected

by parallel analysis. The graph was made using the Prism software 9 (GraphPad, La Jolla, CA, USA). Venn diagrams was created using the InteractiVenn platform [78] and modified in Inkscape

(Inkscape Team). Bubble plots and Volcano plot were made using SRplot [79]. GO Enrichment, KEGG Pathways and Reactome analysis was performed using the DAVID Gene Functional Classification

Tool database. Data were considered statistically significant if _p_ value was _p_ ≤ 0.05. TURBIDITY MESUREMENTS Turbidity measurements were performed to analyze the kinetics of clot

formation in FX-deficient plasma (HYPHEN BioMed) under the influence of FXa-derived NETs and/or purified FXa. After 3 hours, 25 µl of collected NETs and 2 µg of prothrombin were resuspended

in 25 µl of 10 mM HEPES pH 7.4, 150 mM NaCl and 25 mM CaCl2. Then, 50 µl of FX -deficient plasma initiated clotting what was monitored by measuring absorbance at a wavelength of 400 nm for 2

 h at 37 °C and presented as Vmax [RFU/min]. As a control of clot formation FXa was used. SCANNING ELECTRON MICROSCOPY (SEM) Clots formed under the influence of FXa-derived NETs and/or FXa

were fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4). After fixation, the sections were washed in sodium cacodylate buffer and post-fixed in 1% osmium tetroxide.

Next, samples were dehydrated in an alcohol series, dried, and sputtered with gold. Images were captured with a JSM5410 scanning electron microscope (JEOL) at the Institute of Zoology,

Jagiellonian University, in Krakow, Poland. STATISTICS Statistical significance was determined using GraphPad Prism v7 (GraphPad Software) by applying a two-tailed unpaired t-test or by

one-way analysis of variance (ANOVA) with Tukey’s multiple comparisons test. All values are expressed as means ± standard errors (SEM) and _P_ values < 0.05 were considered statistically

significant. DATA AVAILABILITY All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Information. The raw data supporting the findings of

this study are available from the corresponding author upon reasonable request. The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE

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spectrometry-based proteomics evidences. Nucleic Acids Res. 2022;50:D543–52. Article  CAS  PubMed  Google Scholar  Download references ACKNOWLEDGEMENTS The open-access publication has been

supported by the Faculty of Biochemistry, Biophysics and Biotechnology under the Strategic Programme Excellence Initiative at Jagiellonian University in Krakow, Poland. The graphical

abstract was created with BioRender.com https://BioRender.com/h29d439. FUNDING This work was supported by the National Science Centre of Poland with funding grants number 2016/22/E/NZ6/00336

(Sonata Bis 6 to JK) and 2023/51/D/NZ5/01112 (Sonata 19 to DB). AUTHOR INFORMATION AUTHORS AND AFFILIATIONS * Microbiology Department, Faculty of Biochemistry Biophysics and Biotechnology,

Jagiellonian University, Kraków, Poland Danuta Bryzek, Anna Gasiorek, Dominik Kowalczyk, Izabela Ciaston, Ewelina Dobosz, Jan Potempa & Joanna Koziel * Department of Experimental

Hematology, Institute of Zoology and Biomedical Research, Jagiellonian University, Krakow, Poland Michal Santocki & Elzbieta Kolaczkowska * Department of Molecular Biology and Genetics,

Aarhus University, Aarhus, Denmark Katarzyna Kjøge & Jan J. Enghild * MCB, Jagiellonian University, Krakow, Poland Tomasz Kantyka * LMU Hospital, Medizinische Klinik und Poliklinik IV,

Ludwig-Maximilians University, Munich, Germany Maciej Lech * Department of Oral Immunology and Infectious Diseases, School of Dentistry, University of Louisville, Louisville, Kentucky, USA

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You can also search for this author inPubMed Google Scholar * Joanna Koziel View author publications You can also search for this author inPubMed Google Scholar CONTRIBUTIONS

Conceptualization: JK, DB, JP, EK. Methodology: JK, DB, AG, MS, KK, DK, IC, ED. Provided experimental reagents: JK, EK, ML, TK, BP, JE. Investigation: DB, JK, AG, MS, KK DK, IC.

Visualization: DB, ED. Supervision: JK. Writing—original draft: JK, DB, JP. Writing—review & editing: JK, DB, JP. CORRESPONDING AUTHORS Correspondence to Danuta Bryzek or Joanna Koziel.

ETHICS DECLARATIONS COMPETING INTERESTS The authors declare no competing interests. ETHICS APPROVAL AND CONSENT TO PARTICIPATE Peripheral blood was obtained from the Regional Blood Center

RCKiK (Krakow, Poland), which anonymizes blood materials to ensure the confidentiality of human subjects, so donor approval was not required. All animal procedures were approved by the local

Institutional Animal Experimentation Ethics Committee (Second Local Institutional Animal Care and Use Committee, permission numbers: 294/2017 and 22/2023) according to national regulations

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and permissions ABOUT THIS ARTICLE CITE THIS ARTICLE Bryzek, D., Gasiorek, A., Kowalczyk, D. _et al._ Non-classical neutrophil extracellular traps induced by PAR2-signaling proteases. _Cell

Death Dis_ 16, 109 (2025). https://doi.org/10.1038/s41419-025-07428-z Download citation * Received: 09 September 2024 * Revised: 21 January 2025 * Accepted: 04 February 2025 * Published: 19

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