Blocking peptidyl arginine deiminase 4 confers neuroprotective effect in the post-ischemic brain through both NETosis-dependent and -independent mechanisms

Abstract

Peptidylarginine deiminase 4 (PAD4) is an enzyme that modifies proteins by converting positively charged arginine residues to neutral citrulline residues. This process, termed citrullination, has been known to trigger NETosis, a neutrophil cell death pathway involving the release of neutrophil extracellular traps (NETs). Abnormal PAD4 activity and protein citrullination have been linked to various diseases, including those affecting the central nervous system. Herein we investigated the profile of PAD4 expression in an animal model of stroke induced by middle cerebral artery occlusion (MCAO). PAD4 levels were significantly elevated in the ischemic core and penumbra of the affected hemisphere at 3–6 and 6–48 h post-MCAO, respectively. Notably, NETosis induction, indicated by the upregulation of CitH3 (citrullinated histone H3, a NETosis marker), was observed between 48 and 96 h post-MCAO, peaking at 96 h. While PAD4 was present in most brain cell types of sham controls, strong PAD4 induction was primarily observed in neurons during the peak PAD4 induction period (12–24 h post-MCAO). Importantly, intranasal administration of the PAD4 inhibitor BB-Cl-amidine (BBCA) significantly reduced infarct volume and improved neurological and functional outcomes at 24 h post-MCAO, demonstrating a strong protective effect of PAD4 inhibition in ischemic stroke. Staining with an antibody that recognizing citrullinated proteins (F95) revealed an accumulation of these proteins, especially degenerating neurons, however, BBCA treatment significantly suppressed this accumulation in dying neurons. These findings indicate that PAD4-mediated protein citrullination in neurons plays a critical role in promoting ischemic brain damage. Furthermore, delayed administration of BBCA (at 48/72 h post-MCAO) suppresses the NETosis induction observed at 96 h post-MCAO, potentially ameliorating repair processes such as blood vessel regeneration. Collectively, these findings suggest a complex role of PAD4 in cerebral ischemia, with neuroprotective effects (NETosis-independent function) during the acute to subacute period and NETosis-suppressive effects at later time points.

Introduction

Peptidyl arginine deiminase 4 (PAD4) is a post-translational modification enzyme that converts positively charged arginine residues in protein substrates to neutral citrulline, via a process termed deimination or citrullination [1]. PAD4 belongs to the PAD enzyme family, and has been reported to play crucial roles in the regulation of various cellular processes, including gene expression, cell differentiation, and inflammation, in both normal physiology and pathological conditions [2, 3]. Of note, PAD4 is the only PAD family member with a nuclear localization signal [4]. Furthermore, immune cells express high levels of PAD4, and its activity has previously been linked to inflammatory regulation [5]. Specifically, PAD4 plays a well-established role in the formation of neutrophil extracellular traps (NETs) through histones citrullination in neutrophils [6]. In a recent investigation using a rat model of stroke (permanent middle cerebral artery occlusion, pMCAO), we demonstrated that NET formation plays a critical role in inducing brain damage [7, 8].

Citrullination or deamination results in the loss of positive charges in the target molecule, which can alter the conformation of target proteins, causing unfolding or misfolding. These changes can affect the stability of the target protein and its ability to interact with other proteins [9]. Consequently, dysregulated activity of PAD4 and abnormal levels of protein citrullination has been associated with numerous human diseases, including rheumatoid arthritis (RA) [10,11,12], multiple sclerosis (MS) [13], Alzheimer’s disease (AD) [14], and malignant tumors [15]. For example, in RA, PAD4-mediated citrullination of antithrombin, a protein that regulates blood clotting, results in its inactivation, which causes disease progression [10]. Furthermore, the release of active PAD isoforms from dying neutrophil into the synovial fluid may explain the generation of extracellular autoantigens in RA [11]. The importance of PAD4 in various pathological conditions is further supported by the findings of several studies showing that PAD4 deficiency or treatment with PAD inhibitors reduces severity of numerous diseases [10, 12, 13, 16, 17].

PADs have also been reported to play critical roles in various central nervous system (CNS) pathologies. Notably, PADs have been linked to an increase in citrulline-mediated responses within the CNS under hypoxic conditions. For example, studies in neonates with hypoxic ischemic brain injury showed elevated levels of citrullinated proteins, including histone 3, in affected brain regions, and treatment with PAD inhibitors significantly reduced these levels [16]. Ischemic stroke has also been shown to trigger the upregulation of PAD4, resulting in the citrullination of many proteins, which are distributed throughout the brain, particularly in the cortex [18]. Analysis of the components of the thrombi found in patients with ischemic stroke revealed that all thrombi contained NETs, with citrullinated histone 3 forming their structural backbone [19, 20]. In our previous study investigating a permanent MCAO animal model, we observed a significant induction of PAD4 in the cortical penumbra of the ischemic hemisphere 24 h following stroke induction as well as in PMNs (polymorphonuclear neutrophils) isolated 12 h after permanent MCAO [7].

In the present study, we investigated the time course of PAD4 induction and its cellular/subcellular localization in the brain following cerebral ischemia using a transient MCAO animal model. Based on this spatiotemporal expression pattern, we explored the function of PAD4 using a PAD4 inhibitor. The overall aim of this study was to elucidate the mechanisms underlying the neuroprotective effects of PAD4 inhibition. These effects were investigated in the context of both NETosis-independent and -dependent pathways.

Materials and methods

Surgical procedure of MCAO induction

Male Sprague–Dawley rats (7–8 weeks of age) were housed under diurnal lighting conditions with ad libitum access to food and tap water. All animal studies were carried out in strict accordance with the recommendations of the Guide for the Care and Use of Laboratory Animals published by the National Institute of Health (NIH, USA, 2013) and the ARRIVE guidelines (http://www.nc3rs.org/ARRIVE (accessed on August 31, 2021). The animal protocol used was reviewed for ethicality and approved prior to conduction by the INHA University Institutional Animal Care and Use Committee (INHA-IACUC) (approval number INHA210915-782). MCAO was performed as previously described (Davanyaam et al., 2022) [21]. Briefly, 8-week-old male Sprague–Dawley rats (250–300 g) were anesthetized with 5% isoflurane in 30% oxygen/70% nitrous oxide, with maintenance anesthesia during surgery comprising 0.5% isoflurane in the same gas mixture. The right middle carotid artery was occluded for 1 h by advancing a nylon suture (4 − 0; AILEE, Busan, Korea) with a heat-induced bulb at its tip (~ 0.3 mm in diameter) through the internal carotid artery for 20–22 mm from its bifurcation with the external carotid artery. Occlusion was followed by reperfusion for up to 7 d. During the procedure, the left femoral artery was cannulated to obtain blood samples, which were analyzed to assess pH, PaO₂, PaCO₂, and blood glucose levels using the I-STAT device (Sensor Devices, Waukesha, WI, USA) (Supplementary Table 1). A laser Doppler flowmeter (Periflux System 5000; Perimed, Jarfalla, Sweden) was further used to monitor regional cerebral blood flow (CBF) and relative CBF, while a thermoregulated heating pad and a heating lamp were used to maintain a rectal temperature of 37.0 ± 0.5 °C. The animals were randomly allocated to the following groups: sham, MCAO, MCAO + BB-Cl-amidine. Animals in the sham group underwent an identical procedure without MCAO.

Drug administration

Drugs were administered intranasally, as previously described by Kim et al. (2018) [22]. Briefly, rats were anesthetized with an intramuscular injection of a mixture of ketamine (3.75 mg/100 g body weight) and xylazine hydrochloride (0.5 mg/100 g per body weight). A nose drop containing BB-Cl-amidine (BBCA, HY-111347 A; MedChemExpress, Monmouth Junction, NJ, USA) dissolved in PBS (20 µL) was then carefully placed in each nostril of anesthetized animals (maintained in the supine position at a 90^◦ angle) using a pre-autoclaved pipet tip (T-200-Y; Axygen, Union, CA, USA). The procedure was repeated until entire dose was administered, with 2-min intervals between applications.

Staining with 2, 3, 5-triphenyl tetrazolium chloride

Twenty-four hours after MCAO induction, the rats were euthanized, and the whole brains were isolated and coronally sectioned at 2 mm intervals using a metallic brain matrix (RBM-40000; ASI, Springville, UT, USA). Tissue sections were immediately treated with 2% 2,3,5-triphenyl tetrazolium chloride (TTC) and incubated at 37 °C for 15 min. Slices were transferred and stored in 4% paraformaldehyde (PFA). Areas of infarcted tissue were quantified using Scion Image (Frederick, MD, USA). To adjust for edema and shrinkage effects, the ischemic lesion area was calculated as (contralateral hemisphere volume - (ipsilateral hemisphere volume - measured injury volume)). Infarct volume (in mm [3]) was calculated by multiplying the total infarct area across sections by the section thickness.

Modified neurological severity scores

Neurological deficits were assessed using the Modified Neurological Severity Score (mNSS) at 1, 4, and 7 d post-MCAO. The mNSS system includes motor, sensory, balance and reflex tests, and is graded on a scale of 0–18, with higher scores indicating more severe injuries. Motor scores were determined using two tests. (1) Rats were hung by their tails, assigning a score of 0 or 1 for each subcategory (total score 0–3), including flexion of the forelimbs, flexion of the hindlimbs, and head movement > 10° relative to the vertical axis within 30 s; (2) rats were placed on the floor and assign a score of 0 to 3 based on walking ability, as follows: 0, normal walking; 1, unable to walk straight; 2, turning to the paralyzed side; and 3, falling to the paralyzed side. Sensory tests included a placement test (score 0 or 1) and a proprioceptive test (score 0 or 1). The beam balance test assessed balance with a score from 0 to 6, as follows: 0, stable posture; 1, grasping the side of the beam; 2, hugging the beam with one limb off the beam; 3, hugging the beam with two limbs off or spinning around for 60 s; 4, attempting to balance but fell within 20–40 s. 5, trying to balance but fell within 20 s. 6, not attempting to balance or hang. Reflex test scores include scores for pinna reflex, corneal reflex, startle reflex (0 or 1, respectively), and seizures, myoclonus, or dystonia (0 or 1).

Rotarod test

Animals were trained on a rotarod unit rotating at a constant speed of 3 rpm until they could maintain on the rotating spindle for 180 s 24 h prior MCAO induction. Then, 1 d post-MCAO, each rat was tested on the rotarod at speeds of 5, 10, and 15 rpm, and the residence times on the rotarod were measured.

Serum preparation

Animals were anesthetized with an intramuscular injection of a mixture of ketamine (100 mg/kg body weight) and xylazine hydrochloride (23.32 mg/kg body weight). Blood samples were collected through cardiac puncture using a 23 G syringe, and kept at room temperature for 30 min. The blood samples were centrifuged for 15 min at 2000× g and 4 °C. The supernatant was aliquoted and stored at − 80 °C.

Isolation of peripheral blood neutrophils

Neutrophils were isolated from rat blood using Histopaque gradients, as previously described (Kim et al., 2019) [7]. In brief, 3 ml of Histopaque 1077 (Sigma Aldrich) was layered on top of 3 ml of Histopaque 1119 (Sigma Aldrich), and 3.5 ml of rat blood was carefully placed on top to form a three-step gradient in a 15 ml tube. The tube was then centrifuged at 400× g for 30 min, and the first ring containing monocytes was carefully removed. The second ring was transferred to new 15 ml tube containing phosphate buffered solution containing 10% glucose and 0.1% bovine serum albumin (PBS-BG), and centrifuged at 1500× g for 10 min. The resulting pellet was suspended in 3 ml of PBS-BG, layered on top of 3 ml of Histopaque 1119, and centrifuged at 1500 × g for 10 min. The ring containing neutrophils was obtained carefully.

Primary cortical neuron cultures

Experiments were conducted in accordance with the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health (NIH, USA, 2011). The animal protocol used in this study was reviewed and approved by the INHA-IACUC (approval number INHA 201103-734). All efforts were made to reduce the number of animals used and minimize animal suffering. Briefly, mixed neocortical cells were prepared from mouse cerebral cortices extracted at embryonic day 15.5 (E15.5) and cultured as previously described (Seol et al., 2023) [23]. Cortical cells were dissociated using a glasspasteur pipette and plated at a density of six hemispheres per well in 24-well poly-d-lysine (100 µg/mL) and laminin (100 µg/mL) (Thermo Fisher Scientific, Waltham, MA, USA)-coated plates (4 × 10⁵ cells per well). Cultures were maintained in minimum essential medium (MEM: Sigma-Aldrich) supplemented with fetal bovine serum (FBS, 5%), horse serum (5%), glucose (21 mM), and glutamine (2 mM) without antibiotics. On day 7 in vitro (DIV7), when astrocytes had reached confluence underneath neurons, cytosine arabinofuranoside (ara-C, Sigma-Aldrich) was added to a final concentration of 10 µM in MEM containing horse serum (10%) and glucose (21 mM). Cultures were maintained for 2 d to halt microglial growth. Media were changed on alternate days after DIV7, and FBS and glutamine were not supplemented from DIV7. Cultures were used on DIV12-14.

Oxygen-glucose deprivation

Primary cortical cell cultures were prepared and experiments were performed at 12 d in vitro (DIV12). After removal of the original medium, cells were rinsed with glucose-free Earle’s Balanced Salt Solution (EBSS, pH 7.4), and transferred to fresh glucose-free EBSS. Cultures were then maintained in an incubator in an atmosphere of 5% CO₂/95% N₂ at 37 °C for 90 min. Control cultures were maintained in normal EBSS under normal incubation conditions.

Quantification of cell-free DNA

Levels of cell-free DNA in serum were measured using the Quant-iT PicoGreen double-stranded DNA (dsDNA) assay kit (Invitrogen, Carlsbad, CA, USA). Fluorescence was then measured using a spectrofluorometer (Molecular Devices, Sunnyvale, CA, USA) at an excitation/emission wavelength of 480/540 nm. DNA concentrations were calculated using a standard curve prepared using kit-supplied DNA. Data were normalized relative to the NET-DNA concentration in ng/mL.

N-methyl-D-aspartate treatment

For acute excitotoxicity, cortical cell cultures were incubated in serum-free MEM containing 40 µM N-methyl-D-aspartate (NMDA, Sigma-Aldrich) for 10 min. Following treatment, the medium was removed and replaced with fresh MEM medium, and the cells were incubated for 24 h.

LDH assays

Cells were treated with BBCA and subjected to OGD or NMDA treatment. After 24 h, 50 µl aliquots of medium and 50 µl LDH assay reagent (Roche Diagnostics, Mannheim, Germany) were mixed in 96-well plates and incubated for 1 h, and then optical densities were measured at 490 nm using a 96-well plate reader.

Immunoblot analysis

Brain homogenates and whole cell lysates were extracted using RIPA buffer (50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 0.5% NP40, 0.25% sodium-deoxycholate, 0.5% Triton X-100, 10% glycerol, and Complete Mini Protease Inhibitor Cocktail tablet (Roche Diagnostics, Basel, Switzerland). Lysates were centrifuged at 12,000× g for 10 min at 4 °C, and the supernatant was loaded onto sodium dodecyl-sulphate polyacrylamide gel electrophoresis gels (10–14%). Separated proteins were subsequently transferred to polyvinylidene fluoride membranes (PVDF; IPVH00010; Merck Millipore, Darmstadt, Germany) and blocked with 5% nonfat milk or 5% BSA for 1 h. Immunoblotting was carried out using the following primary antibodies; anti-PAD4 (GTX113945; GeneTex, Irvine, CA, USA), anti-citH3 (ab5103; Abcam, Cambridge, UK), anti-Myeloperoxidase (MPO) (PA5-16672; Invitrogen, Carlsbad, CA), and anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (2118 S; Cell Signaling Technology, Danvers, MA, USA). Blots were detected using anti-rabbit HP conjugated or anti-mouse HP conjugated secondary antibody (1:2000, Merck Millipore, Burlington, MA, USA) and a chemiluminescence kit (Merck Millipore).

Immunofluorescence staining

Brains were fixed in 4% PFA solution for 2 d at 4 ^oC, post-fixed in a 30% sucrose solution at 4 ^oC, sectioned at 30 μm using a vibratome, and immunologically stained. Sections were then blocked with 5% FBS, 5% horse serum, and 2% albumin in 0.1% Triton X-100 for 1 h at room temperature. Primary antibodies for anti-neuronal nuclei (NeuN) (MAB377; Merk Millipore), anti-ionized calcium-binding adaptor molecule-1 (Iba1) (orb18542; Biorbyt, Cambridge, UK), anti-glial fibrillary acidic protein (GFAP) (556327; BD Biosciences, Franklin Lakes, NJ, USA), anti-CitH3 (ab281584, Abcam), anti-Myeloperoxidase (MPO) (ab90810; Abcam), anti-Lymphocyte antigen 6 complex locus G6D (Ly6g) (ab25024; Abcam), PAD4 (GTX113945, GeneTex), anti-F95 (MABN328, Merk Millipore), and anti-RECA (MCA970GA; BIO-RAD, Kidlington, UK) were all diluted 1:200. After incubation with the primary antibodies, brain sections were washed with PBS and incubated with FITC-conjugated anti-mouse IgG (1:200; Merk Millipore) or Rhodamine-conjugated anti-rabbit IgG (1:200; Merk Millipore) for 1 h at room temperature. Sections were subsequently mounted on slides with VECTASHIELD Antifade Mounting Solution containing DAPI (Vector Laboratories, Peterborough, UK), and examined under a Zeiss LSM 510 META microscope (Carl Zeiss Meditec AG, Jena, Germany).

Small interfering RNA (siRNA) transfection

Mouse PAD4-specific siRNA (siPAD4; 5’-AUC AAU GAA AUU CUG UCC AAC AAG A-3’ and 5’-UCU UGU UGG ACA GAA UUU CAU UGA UUG-3’) and a nonspecific siRNA (siCon; 5’-CGU UAA UCG CGU AUA AUA CGC GUA T-3’ and 55′-AUA CGC GUA UUA UAC GCG AUU AAC GAC-3′) were purchased from Integrated DNA Technologies, Inc. (Coralville, IA, USA). siPAD4 or siCon was prepared at a final concentration of 40 nM for transfection and mixed with 1 µL/per well of Lipofectamine RNAiMAX reagent (Invitrogen, Carlsbad, CA, USA) in Opti-MEM (Invitrogen)according to the manufacturer’s instructions. The siRNA-lipid complexes were added to primary cortical cultures and incubated for 24 h.

Terminal deoxynucleotidyl transferase dUTP nick end labeling staining

Apoptotic cells were detected using the DeadEnd™ Fluoromeric terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) system (Roche) according to the manufacturer’s instructions. Briefly, obtained tissue sections were rinsed three times with 0.1% Triton X-100 in PBS. The DNA end labeling was performed by incubating tissues with rTdT reaction mixture for 60 min at 37 °C followed by termination with 2 X SSC (30 mM of sodium citrate (pH 7.4) and 300 mM of sodium chloride). Sections were then immunostained with anti-NeuN or anti-PAD4 antibody, and mounted with DAPI (Vector Laboratories). Apoptotic cells were quantified by counting TUNEL-positive cells, and the data were presented as the ratio of TUNEL-positive cells to total NeuN-positive cells.

Fluorescein isothiocyanate-dextran staining

Fluorescein isothiocyanate (FITC)-labeled dextran (70 kDa) (Sigma-Aldrich) was applied according to a previously published protocol (Kim et al., 2020) [24]. Briefly, rats were subjected to induction of MCAO and administered dextran (60 mg/kg) intravenously 6 h prior to sacrifice at 7 d post-MCAO. Brains were subsequently extracted and placed in 4% PFA solution. After incubation for 2 d at 4 °C, vibratome sections were performed on the brains. These sections were mounted on slides and examined under a confocal microscope (Carl Zeiss, Oberkochen, Germany). Images depicting the distribution of FITC-dextran were captured and examined using ImageJ software (https://imagej.nih.gov/ij/download.html) for quantitative analysis of immunofluorescence intensity.

IgG staining

Rat IgG staining was conducted in accordance with a previously described protocol (Kim et al., 2020) [24]. To eliminate endogenous peroxidase activity, brain sections were first treated with 3% H₂O₂ in PBS for 30 min, and then blocked for 1 h at room temperature. Sections were then incubated in anti-rat IgG antibody (Vector Laboratories) solution at a concentration of 1:200 overnight at 4° C, incubated with biotinylated rat IgG antibody for 1 h, and then incubated with Vectastain ABC reagent (Vector Laboratories) for 1 h. Visualization of the reaction products was achieved through 3,3’-diaminobenzidine (DAB) staining, and the quantification of stained brain sections was performed using the Scion image measurement program (https://scion-image.software.informer.com).

Statistical analysis

Two-sample comparisons were performed using the Student’s t-test, while multiple comparisons were performed using one-way or two-way analysis of variance, followed by Tukey’s post hoc test. PRISM software 5.0 (GraphPad Software Inc., San Diego, CA, USA) was used for all analyses, and results are presented as the means ± SEMs. Statistical significance was set at p < 0.05.

Results

The temporal profile of PAD4 induction in the post-ischemic brain

To understand the role of PAD4 in ischemic brain injury, we investigated its temporal expression profile in the post-ischemic brain using a transient MCAO animal model induced by occlusion lasting 60 min. In the cortical penumbra of the ischemic hemisphere (Fig. 1A), PAD4 levels significantly increased starting at 6 h post-MCAO, peaking at 24 h, and gradually declining thereafter (Fig. 1B and D). In contrast, the cortical core displayed a noticeable upregulation at 3–6 h post-MCAO, which was followed by a substantial decrease to or below baseline levels (Fig. 1C and E). Notably, a significant increase in serum PAD4 levels began 6 h post-MCAO, with a gradual and further rise continuing until 96 h (Fig. 1F and G). Results demonstrate a robust induction of PAD4 in the post-ischemic brain, which was particularly pronounced in the cortical penumbra, with evidence indicating extracellular secretion.

Fig. 1

PAD4 levels in the brain tissue and serum following transient MCAO (A) Schematic representation of the cortical core and penumbra regions of the post-ischemic brain. (B-C) PAD4 protein levels in the cortical penumbra (B) and cortical core (C) were measured by immunoblotting at 3, 6, 12, 24, 48, and 96 h post-surgery. (F) PAD4 protein levels in the serum were assessed by immunoblotting at 3, 6, 12, 24, 48, and 96 h post-surgery. (D, E, G) Quantification of PAD4 protein levels from immunoblots. Data are presented as the mean ± SEM (n = 4). * p < 0.05, ** p < 0.01, *** p < 0.001 compared to sham-operated controls

Delayed induction of citrullinated histone H3 (NETosis) in the post-ischemic brain

The substantial PAD4 induction observed following transient MCAO led us to investigate the temporal profile of NETosis induction. In both the cortical penumbra and core (Fig. 1A), a significant increase in citrullinated histone H3 (CitH3, a well-established marker of NETosis) levels was detected at 48 and 96 h (Fig. 2A-D), indicating a delayed induction of NETosis within the brain tissue following transient MCAO. Circulating neutrophils isolated from the peripheral blood of sham- and MCAO-operated rats exhibited a significant increase in CitH3 levels at 48 h post-MCAO, with a further increase observed at 96 h (Fig. 2E and F). Similar results were observed for myeloperoxidase (MPO) (Fig. 2E and G). Furthermore, the amount of cell-free double-strand DNA (dsDNA, another NETosis marker) released by PMNs in serum was significantly increased at 24 h and further increased at 48 and 96 h post-MCAO (Fig. 2H). These findings indicate that neutrophil activation and NETosis induction occur around 48–96 h in both peripheral blood and brain parenchyma following ischemic stroke. Triple fluorescent staining with antibodies against PAD4 and Ly6g (a neutrophil marker) and DAPI revealed PAD4 localization in neutrophils at 48 h post-MCAO, when neutrophils were detected in brain tissue or within blood vessels (Fig. 2J, arrows and double arrows, respectively). The characteristic NETotic morphology became evident at 96 h brain sections, which were stained with antibodies against CitH3 and MPO and DAPI (Fig. 2K). Some CitH3-positive/MPO-positive cells displayed a lysed morphology, both in the brain parenchyma and within the blood vessels (Fig. 2K, arrows and double arrows, respectively). Collectively these results indicate that delayed PAD4 induction in neutrophil, specifically in PMN isolated between 48 and 96 h post-MCAO, may be responsible for the delayed NETosis induction observed. Notably, the majority of PAD4 immunoreactivity was observed in Ly6g-negative cells with neuronal morphology at all-time points examined, including sham controls (arrowheads, Fig. 2I-K).

Fig. 2

Induction of CitH3 in the brain tissue and PMNs following ischemic insult (A-D) CitH3 protein levels in the cortical penumbra and cortical core (defined in Fig. 1A) were measured by immunoblotting at 3, 6, 12, 24, 48, and 96 h after MCAO surgery. (E-G) CitH3 and MPO levels were assessed by immunoblotting in PMNs isolated at 12, 24, 48, and 96 h after MCAO surgery. (H) The amounts of cell-free dsDNA in the serum were assessed using Quant-iT PicoGreen dsDNA reagent. (B, D, F-H) Data are presented as the mean ± SEM (n = 4). * p < 0.05, ** p < 0.01, *** p < 0.001 compared to sham-operated groups. (I-K) Coronal brain sections were prepared from sham controls (I) and the cortical penumbra or core of the ischemic hemisphere (J, K) at 48 (J) or 96 h (K) post-MCAO. Triple immunofluorescence staining was conducted with anti-Ly6G antibody, anti-PAD4 antibody, and DAPI (I, J) or with anti-CitH3 antibody, anti-MPO antibody, and DAPI (K). Arrows indicate co-localization of anti-PAD4 with anti-Ly6G or anti-CitH3 with anti-MPO in brain parenchyma, while double arrows indicate the co-localization of anti-PAD4 with anti-Ly6G or anti-CitH3 with anti-MPO within blood vessels. Arrowheads indicate the co-localization of anti-PAD4 with neuron-like cells. Scale bars represent 50 μm

Induction of PAD4 occurs primarily in neurons at all time-points in the post-ischemic brain

To investigate the cell types expressing PAD4 observed in Fig. 2I and J, triple immunofluorescence staining using antibodies against PAD4, NeuN (a neuronal marker), and DAPI was performed. In sham controls, PAD4 was detected in the cytoplasm of most neurons (NeuN-positive cells) (arrows, Fig. 3B). At 24 h post-MCAO, overall PAD4 immunoreactivity was significantly enhanced in the cortical penumbra of the ischemic hemisphere compared to sham controls, while it was markedly decreased in the cortical core (Fig. 3A), which is consistent with the findings shown in Fig. 1B-E. Within the penumbra, PAD4 staining was primarily localized in the neuronal cytoplasm, particularly in the perinuclear region, depicting punctate and irregular cytoplasmic distributions (arrows, Fig. 3C-C2). Interestingly, some neurons in both the penumbra and core displayed a loss of neuronal morphology with reduced PAD4 immunoreactivity (asterisks, Fig. 3C1, C2, D1). PAD4 was also detected in many NeuN-negative cells (arrowheads, Fig. 3B-D1). We found that most of the NeuN-negative cells are microglia but not astrocytes based on the triple immunofluorescence staining results using anti-Iba1 (a microglia marker) or anti-GFAP (an astrocyte marker) antibody (Supplementary Fig. 1). Notably, PAD4 immunoreactivity was observed as scattered dot-like structures in the extracellular space (interstitium) in ischemic hemisphere in both the cortical penumbra and core (Fig. 3C-D1). These findings indicate that PAD4 is primarily localized in the cytoplasm of neurons and, to a lesser extent, in the microglia.

Fig. 3

PAD4 expression in the different brain cell types in the post-ischemic brain (A) Schematic representation of the brain region bordering the infarct core and penumbra. (B-D1) Coronal brain sections were prepared from sham controls (B) as well as the cortical penumbra (C, C1, C2) and cortical core (D, D1) of the MCAO group at 24 h post-surgery. Triple immunofluorescence staining was performed with anti-PAD4 antibody, anti-NeuN antibody, and DAPI. Arrows indicate PAD4 immunoreactivity in neurons and arrowheads indicate PAD4 immunoreactivity in non-neuronal cells. Asterisks depict PAD4 immunoreactivity in degenerating cells or cell debris. Scale bars represent 50 μm

Early PAD4 inhibition exerts a dose-dependent neuroprotection after ischemia

The induction of PAD4 observed in the post-ischemic brain and its predominant localization in neurons led us to investigate its potential role during the acute to subacute phases of ischemic injury. To evaluate the effect of PAD4 inhibition during this timeframe, we administered BB-Cl-amidine hydrochloride (BBCA, a PAD inhibitor) intranasally immediately following suture removal in MCAO animals (Fig. 4A). Administration of 5, 10, and 50 µg/kg of BBCA reduced the infarct volumes to 58.6 ± 8.3%, 39.0 ± 7.3%, and 29.2 ± 9.5% of the treatment-naïve MCAO control group, respectively, indicating a dose-dependent neuroprotective effect (Fig. 4B and C). Administration of BBCA at all doses (5, 10, 50 µg/kg) improved the mean modified neurological severity scores (mNSSs), with the most significant improvement observed at 50 µg /kg (Fig. 4D). Motor function was further evaluated using the rotarod test at 2 d post-MCAO with speeds of 5, 10, or 15 rpm. The group administered with 10 µg/kg BBCA displayed significantly longer latencies (time spent on the rod) compared to the saline-treated MCAO group, reaching levels comparable to the sham group at all tested speeds (Fig. 4E). These findings demonstrate that the neuroprotective effect of BBCA translates to improved neurological and motor outcomes.

Fig. 4

BB-Cl-amidine treatment reduces infarct volume and improves neurological function following cerebral ischemia (A) BB-Cl-amidine (BBCA) was administered intranasally immediately following suture removal in MCAO animals. Infarct volumes were measured 24 h later using TTC staining. (B-D) MCAO animals were administered with varying doses of BBCA (5, 10, 50 µg/kg) intranasally. Mean infarct volumes were measured at 1 day post-MCAO. Representative images of infarcts in coronal brain sections are shown (B), and data are presented as the mean ± SEM (n = 4 or 5) (C). Neurological deficits were assessed using the modified neurological severity scores at 1 day post-MCAO (D). (E) MCAO animals were administered with 10 µg/kg BBCA intranasally. The rotarod test was performed at 1 day post-MCAO. Animals were allowed a 1 h rest period between trials at different speeds (5, 10, or 15 rpm). Data are presented as the mean ± SEM (n = 4 or 5). Sham, sham-operated animals (n = 4); MCAO, saline-treated MCAO control animals (n = 4); MCAO + BBCA, BBCA-administered MCAO animals (n = 5 for each dose). ***p < 0.001 compared to the sham group and ^##p < 0.01, ^###p < 0.001 compared to the MCAO group

Citrullinated proteins accumulated in degenerating neurons in the post-ischemic brain

To investigate the accumulation of citrullinated proteins in the post-ischemic brain and its relationship with the potential protective effect of BBCA, we employed immunofluorescence staining with F95, anti-pan-deimination/citrullination antibody. In sham controls, F95 immunoreactivity was minimal, with only a few scattered dots observed within the cytoplasm of neurons (NeuN-positive cells ) (arrows, Fig. 5A). At 12 h post-MCAO, immunoreactivity of citrullinated proteins was elevated in the cortical penumbra, localized primarily in neurons both in cytoplasm and nuclei (arrows in Fig. 5B and C; Supplementary Fig. 2). At 24 h post-MCAO, F95 immunoreactivity increased further in neurons (arrows, Fig. 5E). Notably, F95 staining was detected in damaged neurons exhibiting a degenerating or fragmented morphology at both 12 and 24 h post-MCAO (asterisks in Fig. 5C and E). Additionally, F95 immunoreactivity was observed in non-neuronal cells (arrowheads, Fig. 5C and E; Supplementary Fig. 3) as well as within blood vessels (double arrowheads, Fig. 5B and E) both at 12 and 24 h. Importantly, intranasal administration of BBCA (50 µg/kg) immediately following surgery significantly reduced F95 immunoreactivity in neurons at both 12 and 24 h post-MCAO (Fig. 5D, F, and G). Moreover, localization of F95 in TUNEL-positive cells was markedly increased in the cortical penumbra at 24 h post-MCAO (double asterisks in Fig. 5H), however, it was significantly reduced by BBCA administration (50 µg/kg) (Fig. 5H-J). Co-localization of citrullinated proteins in degenerating neurons and its suppression by BBCA were further confirmed using Fluoro-jade C staining, which detects all degenerating neurons (Supplementary Fig. 4). Collectively, these findings indicate that citrullinated proteins accumulate in degenerating neurons and BBCA treatment suppresses the accumulation of citrullinated proteins in neurons, which might contribute to the robust neuroprotective effect of BBCA in the post-ischemic brain.

Fig. 5

Accumulation of deiminated proteins in neurons, especially in damaged neurons in the post-ischemic brain Coronal brain sections were prepared from sham controls (A) as well as the cortical penumbra (B-F, H, I) of the MCAO group at 12 h (B-D) or 24 h (E, F, H, I) post-MCAO. Coronal brain sections were further prepared from the cortical penumbra of the MCAO + BBCA group at 24 h (F, I) post-MCAO. Triple immunofluorescence staining was conducted using the anti-F95 antibody, anti-NeuN antibody, and DAPI (A-F) or with the anti-F95 antibody, TUNEL, and DAPI (H, I). (G) The ratio of F95- to NeuN-positive cells is presented (n = 12 from 3 animals). (J) The ratio of TUNEL-positive cell to F95-positive cells is presented (n = 12 from 3 animals). Arrows indicate F95 immunoreactivity localized in the neuron, while asterisks indicate localization of F95 immunoreactivity in degenerating neurons. Arrowheads highlight F95 immunoreactivity in non-neuronal cells, and double arrowheads highlight F95 immunoreactivity in vasculature. Double asterisks in H and I indicate localization of F95 immunoreactivity in TUNEL-positive cells. Scale bars represent 50 μm. **p < 0.01, ***p < 0.001 compared to the sham group and ^##p < 0.01, ^$p < 0.05 between indicated groups

PAD4 inhibition exerts protective effects in cortical neurons against oxygen-glucose deprivation and excitotoxicity

To further investigate the direct role of PAD4 in neurons, we subjected primary cortical neuron cultures to oxygen glucose deprivation (OGD), a well-established in vitro model of ischemia. We observed elevated PAD4 levels as early as 3 h after OGD induction, followed by a gradual decline (Fig. 6B and C). Pre-treatment with BBCA (0.01, 0.05, 0.1, and 0.5 nM) for 3 h prior to OGD resulted in a dose-dependent protective effect against OGD-induced cell death (Fig. 6A and D). Similarly, NMDA (40 µM, 10 min), a chemical that mimics excitotoxicity, also triggered significant PAD4 induction in primary cortical cultures (Fig. 6B and C), and BBCA treatment significantly attenuated neuronal death in these cells (Fig. 6E). These findings suggest a potential role for PAD4 in OGD- or NMDA-induced neuronal death. Importantly, silencing PAD4 expression using siRNA significantly suppressed OGD- or NMDA-induced neuronal death, further reinforcing the importance of PAD4 in this process (Fig. 6F-I). Collectively, these results demonstrate the direct protective effect of PAD4 inhibition in primary cortical neurons against both OGD-induced and excitotoxicity-mediated neuronal death, which are two primary causes of acute neuronal death following ischemia.

Fig. 6

BBCA protects primary cortical neurons from OGD- and NMDA-induced cell death (A) Primary cortical neuron cultures were subjected to oxygen-glucose deprivation (OGD) for 90 min–10 min of NMDA treatment (40 µΜ). BBCA (0.01, 0.05, 0.1, or 0.5 nM) was treated for 3 h before OGD or NMDA, and PAD4 siRNA was transfected 24 h before these treatments. (B, C) PAD4 protein levels were assessed by immunoblotting at 1, 3, and 6 h following the initiation of OGD or NMDA treatment (as indicated in A). (D, E) Cell death was measured using the LDH assay at 24 h after OGD (D) or NMDA treatment (E). (F-I) PAD4 siRNA (40 nM) was transfected 24 h before these treatments. The efficacy of PAD4 siRNA was assessed by immunoblotting analysis at 27 h post-transfection (F, G) and cell death was measured using the LDH assay at 24 h after OGD (H) or NMDA treatment (I). (C-E, G-I) Results are presented as the mean ± SEM (n = 4). *p < 0.05, **p < 0.01, ***p < 0.001 compared to untreated group. ^##p < 0.01, ^###p < 0.001 between indicated groups

PAD4 inhibition directly suppresses delayed NETosis following cerebral ischemia

As the neuroprotective effect (infarct suppression) of PAD4 inhibition by 0 h BBCA administration was evaluated at 24 h post-MCAO, we investigated whether the same PAD4 inhibition could suppress the delayed NETosis occurring at 48 ~ 96 h post-MCAO. The intranasal administration of BBCA (50 µg/kg) immediately after surgery (0 h BBCA, Fig. 7A) significantly reduced CitH3 induction at 96 h post-MCAO (Fig. 7B and C). The induction of PAD4 was also moderately suppressed (Fig. 7B). When we administered BBCA (50 µg/kg) at two later time points: 48 and 72 h post-MCAO (48/72 h BBCA) (Fig. 7A), CitH3 induction in the cortical penumbra at 96 h was also significantly suppressed (Fig. 7B and C). Notably, in PMNs isolated 96 h post-MCAO, 48/72 h BBCA administration significantly suppressed CitH3 induction to the levels comparable to that obtained by 0 h BBCA administration (Fig. 7D and E). Furthermore, reduction of Sytox green staining and free dsDNA level in serum also confirmed the efficient suppression of delayed NETosis induction by 48/72 h BBCA administration (Fig. 7F and G). Interestingly, infarct volumes and neurological deficits measured at 7 d post-MCAO were significantly reduced only in the 0 h BBCA-administration group but not in 48/72 h treatment group (Supplementary Fig. 5). Collectively, these results demonstrate that delayed PAD4 inhibition by 48/72 h BBCA administration can directly suppress NETosis induction observed 96 h post-MCAO, independent of its earlier neuroprotective effects.

Fig. 7

Suppression of delayed NETosis in the post-ischemic brain by intranasal BBCA administration (A) BBCA (50 µg/kg) was administered intranasally either once immediately following suture removal (0 h BBCA) or twice at 48 and 72 h post-MCAO (48/72 h BBCA). (B-E) Levels of CitH3 and PAD4 were examined in the brain tissue (B and C) and PMNs (D and E) at 4 d post-MCAO. (F, G) Sytox green staining was conducted with PMNs isolated at 4 d post-MCAO (F) and the amounts of free dsDNA in serum were measured using the Quant-iT PicoGreen assay kit (G). Sham, sham-operated animals (n = 4); MCAO, saline-treated MCAO control animals (n = 4); MCAO + 0 h, BBCA-administered at 0 h MCAO animals (n = 4); MCAO + 48/72 h, BBCA-administered at 48 and 72 h post-MCAO animals (n = 4). *p < 0.05, ***p < 0.001 compared to sham group, ^##p < 0.01, ^###p < 0.001 compared to the MCAO group, and ^$p < 0.05, ^$$p < 0.01 between indicated groups

PAD4 inhibition mitigates vessel damage and promotes vessel repair following cerebral ischemia

Previous studies have shown that suppression of NETosis protects blood vessels and promotes their repair in both permanent and distal MCAO models (Kim et al., 2019; Kang et al., 2020) [7, 25]. Blood vessel integrity was assessed by measuring the total length and density of blood vessels (Fig. 8A, asterisk) using RECA-1 (endothelial cell marker) immunostaining. Both the vessel density and total length were recovered by BBCA administration at 48/72 h post-MCAO, which were comparable to the value observed in 0 h BBCA administration group (Fig. 8B-D). Next, we evaluated the functionality of these newly formed vessels. In the IgG extravasation experiment, the area of the leakage region in the ischemic hemisphere was significantly reduced in the MCAO + BBCA group (both 0 h and 48/72 h) compared to treatment-naïve MCAO controls (Fig. 8E and F). For isothiocyanate (FITC)-dextran experiments, animals were administered FITC-dextran 6 h before sacrifice at 7 d post-MCAO (Fig. 8A). A significantly smaller leakage area was detected in the MCAO + BBCA group (both 0 h and 48/72 h) compared to treatment-naïve MCAO controls (Fig. 8G and H), indicating reduced blood vessel permeability following BBCA administration. When we examined effects BBCA on vessel damage at 96 h post-MCAO, similar suppressive effects were obtained (Supplementary Fig. 6), further confirming the mitigation of vessel damage by PAD4 inhibition. In conclusion, our data collectively demonstrated that delayed BBCA administration at 48/72 h post-MCAO mitigates blood vessel damage and promotes functional blood vessel repair, likely by suppressing delayed NETosis in the post-ischemic brain.

Fig. 8

BBCA suppresses blood vessel damage in the post-ischemic brain (A) BBCA (50 µg/kg) was administered intranasally either once immediately after suture removal (0 h BBCA) or twice at 48 and 72 h (48/72 h BBCA) post-MCAO. (B-D) Coronal brain sections were prepared from the cortical penumbra at 7 d post-MCAO, and double-stained with anti-rat endothelial cell antigen-1 (RECA-1) antibody and DAPI. Representative images are presented (B), while RECA-1-positive vessel densities are presented as the mean ± SEM (n = 12 from 4 animals) (C) and total vessel lengths measured using AngioTool software 0.6a (https://angiotool.sofware.informer.com/0.6) are presented as the mean ± SEM (n = 12 from 4 animals) (D). (E, F) Brain sections stained with biotinylated rat anti-IgG antibody and the IgG-positive area was measured using Scion Image 4.0 (https://scion-image.sofware.informer.com/). (G, H) FITC-dextran (60 mg/kg) was injected intravenously 6 h prior to sacrifice at 7 d post-MCAO and coronal brain sections with FITC-dextran images were acquired with confocal microscopy (G) and FITC intensity was measured using ImageJ (https://imagej.nih.gov/ij/download.html) (H). (F, H) The data are presented as the mean ± SEM (n = 4 for IgG staining and n = 12 (4 sections from 3 animals) for FITC-dextran images). Scale bars in B represent 200 μm, while those in G represent 100 μm. Sham, sham-operated animals; MCAO, saline-treated MCAO control animals; MCAO + 0 h, BBCA-administered at 0 h MCAO animals; MCAO + 48/72 h, BBCA-administered at 48 and 72 h post-MCAO animals. **p < 0.01 compared to sham controls and ^#p < 0.05, ^##p < 0.01, ^###p < 0.001 compared to MCAO + saline controls

Discussion

Immunohistochemical analysis of PAD4 expression in the post-ischemic brain revealed several intriguing findings. First, PAD4 is present in most brain cell types; however, with expression being the most prominent in neurons. Notably, PAD4 levels were significantly increased in the cortical penumbra, but markedly and rapidly decreased in the ischemic cores, as shown through immunoblotting and immunofluorescence staining. This decrease in the ischemic core may have been caused by a rapid extracellular secretion from damaged neurons, or from stimulated microglia, resulting in a rapid increase in serum PAD4 levels as early as 6 h post-MCAO. Second, the high basal PAD4 expression observed in sham controls is another noteworthy finding. Given the typically low intracellular calcium levels in healthy neurons, as well as the dependency of PAD4 activity on high calcium levels, minimal enzyme activity would be expected in the normal brain. However, our findings of high basal expression of PAD4 along with the presence of citrullinated proteins (albeit at low levels) in sham controls are consistent with previous reports [26, 27]. Further investigation is therefore required to elucidate the potential physiological roles of PAD4 under these different conditions. Third, the lack or minimal expression of PAD4 in astrocytes was interesting. However, prior studies have reported the selective expression of PAD2 and PAD4 in astrocytes and neurons, respectively [26, 28]. In line with this, Kang et al. (2020) [25] observed only minimal PAD4 overexpression in astrocytes in an experiment using a PAD4-flag overexpression system. Therefore, PAD2 may be responsible for citrullinated proteins detected in astrocyte of patients with MS [29] and AD [14, 30]. We also detected citrullinated proteins in the astrocytes of the ischemic brain (Supplementary Fig. 3). These observations collectively indicate that PAD4 and PAD2 play distinct roles in neurons and astrocytes, respectively, highlighting the need for further investigation.

The predominant cytoplasmic localization of PAD4 in neurons, observed in both sham controls and in the post-ischemic brain, is intriguing as PAD4 is known to have a canonical nuclear localization signal [4]. However, the cytoplasmic localization of PAD4 has been reported in neurons as well as in other cell types, including kidney cells [26, 31]. Moreover, various cytoplasmic targets of PAD4, for example, intermediated filaments and related proteins, have been implicated in the pathogenesis of MS [32] and RA [33]. In an acute ischemic kidney injury, PAD4 citrullinates NEMO (NF-kappa-B essential modulator, IKKγ) in the cytoplasm of proximal tubular cells, which exacerbates ischemic injury and inflammation by promoting renal tubular NF-κB activity [34]. In addition to the cytoplasm, we observed a significant increase in serum PAD4 levels following cerebral ischemia (Fig. 1F-G). Notably, PAD4 retained its activity in the extracellular environment owing to high Ca²⁺ levels, being capable of citrullinating plasma proteins, as shown in the plasma of patients with various diseases, including antithrombin in RA [10] and malignant tumors [15], fibrinogen in RA [35], and a disintegrin and metalloproteinase with thrombospondin type-1 motif-13 (ADAMTS13) in venous thrombosis [36, 37]. Consistent with these findings, our study showed a significant increase in PAD4 levels in the extracellular space of the brain tissue (as evidenced by strong immunoreactivity, Fig. 3) in addition to the serum (Fig. 1F-G), supporting a potential role for extracellular PAD4 in ischemic stroke. Therefore, PAD4 may play critical roles in both intracellular and extracellular environments, however, further investigation is required.

Our study demonstrated a robust reduction of infarct volume at 24 h post-MCAO in animals treated with BBCA immediately following suture removal. These results indicate a critical role for PAD4, particularly during the acute to subacute phase following ischemic stroke. The accumulation of citrullinated proteins and their co-localization with TUNEL and FJC staining in degenerating neurons indicate a potentially detrimental role of these modified proteins. In addition to protein unfolding, misfolding, disrupted interactions with other molecules [9], and solubility changes [38], citrullination can increase the susceptibility of a protein to degradation by proteases [9]. Supporting this, Gößwein et al. (2019) [39] demonstrated that calpain-mediated proteolysis of the Lamin B1 and HMGB1 proteins occurs only after their citrullination by PAD4. Protein citrullination can also lead to the generation of autoantibodies against these modified proteins and their breakdown products, potentially triggering autoantibody production, as it was observed in patients with RA [40,41,42]. Similar autoimmune responses against citrullinated proteins have been implicated in variety of neurodegenerative diseases [43], For example, PAD-mediated citrullination of endogenous myelin has been shown to drive a potent secondary autoimmune response in MS, inducing the progression of inflammatory demyelination and axon damage [44]. While stroke is primarily an acute event causing neuronal damage, it can also lead to massive and progressive neuronal loss and the extracellular release of cellular contents. These secreted substances include potentially immunogenic citrullinated proteins and their fragments, which can enter the bloodstream and lymphatic system, thereby triggering an immune response and autoantibody production. Therefore, PAD4 functions beyond NETosis induction and the effects of BBCA may also be interpreted in light of these potential PAD4 functions. However, present study doesn’t establish a direct causal link between specific protein citrullination and neuronal death, future research needs to investigate specific citrullination protein targets and their association with distinct neuronal death pathways and underlying molecular mechanisms. It is important to note that some studies have indicated the beneficial roles of PAD4. Nakamura et al. (2023) [45] observed PAD4 expression in surviving neurons after stroke, suggesting a potential role in the recovery. Similarly, Tanikawa et al. (2018) [46] reported PAD4-mediated inhibition of protein aggregation in ALS models. These findings highlight the complex and context-dependent functions of PAD4, thus warranting further investigation into its potentially opposing roles in various diseases.

Our study also demonstrated that delayed NETosis, occurring around 4 d post-MCAO, was significantly suppressed by BBCA administration either 0–48/72 h, and that this suppression resulted in a marked improvement in vessel regeneration at 7 d post-MCAO. Importantly, the benefits obtained by delayed BBCA administration (48/72 h) were not owing to indirect neuroprotective effects as evidenced by the lack of infarct suppression, but rather a direct consequence of the NETosis-suppressing effect. Our findings align with those of Kang et al. (2020) [25], who demonstrated that in distal MCAO animal model, NETosis peaks at 3 d post-MCAO, and PAD4 overexpression exacerbated vascular damage and reduced neovascularization by promoting NET release. Extruded NETs contain cytotoxic substances including histone, HMGB1, elastase, and MPO, which directly damage endothelial cells and increase vascular permeability [7, 47, 48], leading to exacerbated inflammation and hindering the repair process. Additionally, vessel leakage has been shown to promote the extravasation of immune cells and blood-derived toxic proteins [49, 50]. Consequently, the stability of new blood vessels and restoration of the blood-brain barrier are crucial for maintaining a stable brain microenvironment. Thus, PAD4, a key enzyme inducing NETosis, plays a crucial role in impeding delayed vascular remodeling after stroke. Future research is needed to elucidate the molecular mechanisms by which NETosis influences vessel damage and repair, as well as the regulation of blood-brain barrier markers.

Overall our study provides strong evidence to indicate a critical role of PAD4 in the post-ischemic brain. The marked increase of PAD4 specifically in cortical neurons of the ischemic hemisphere, along with its co-localization with accumulated citrullinated proteins in degenerating neurons, indicate a crucial function for PAD4-mediated protein citrullination in these cells following ischemia. This finding highlights the multifaceted nature of the function of PAD4 in the ischemic brain, operating through both NETosis-dependent and -independent mechanisms. Given the robust neuroprotection observed with PAD4 inhibition, future investigations should explore target proteins and potential therapeutic applications.

Conclusion

Overall, current study demonstrates that inhibiting PAD4 offers robust protective effects in the ischemic brain, which are achieved by suppressing both acute and subacute neuronal death, as well as delayed NETosis. This study is the first, to the best of our knowledge, to characterize the role of PAD4 in ischemia, proposing it as a potential therapeutic target. Furthermore, we showed the potential importance of protein modification (citrullination) in the ischemic brain. We believe that this phenomenon warrants exploration of the function of PAD4 in other CNS diseases including degenerative conditions.