A Novel Selective Inhibitor JBI-589 Targets PAD4-Mediated Neutrophil Migration to Suppress Tumor Progression

Polymorphonuclear neutrophils (PMN) are one of the major components of the tumor microenvironment (TME) contributing to primary tumor growth and formation of the premetastatic niche (1–5) and limiting success of cancer therapies (6). Neutrophils are derived in the bone marrow (BM) from common myeloid progenitor cells (7) and mobilized out of the BM to the blood in response to stimuli including infection, inflammation, or cancer. The mobilization of neutrophils from the BM to the circulation and their migration to the tissues is a complex process that is highly regulated by multiple factors and requires the upregulation of C-X-C chemokine receptor 2 (CXCR2) together with the downregulation of CXCR4 signaling (8, 9). A number of molecules that stimulate neutrophil release from the BM and their migration, including the CXCR2 ligands CXCL1 and CXCL2, are produced by tumor cells and are significantly upregulated in tumor-bearing hosts (10, 11).

Cancer progression is accompanied by accumulation of pathologically activated PMN-polymorphonuclear myeloid derived suppressor cells (PMN-MDSC). Despite their common origin and phenotypical similarity, these cells are distinct in their biochemical properties, transcriptome, and functional activity. In contrast to PMN, PMN-MDSC are able to suppress responses of T and B lymphocytes and NK cells and promote tumor progression and metastasis via non-immune mechanisms (12, 13). Tumor-associated neutrophils are a functionally diverse population, consisting of PMN with antitumor and PMN-MDSC with protumorigenic properties (14–16). The balance between PMN and PMN-MDSC depends on the stage of the cancer. A recent study found that the immune suppressive PMN-MDSC accumulate in tumor tissues at early stages of tumor development (16). As a result, a highly suppressive environment is created in tumors and prevents rejection of tumors via immune-mediated mechanisms. The presence of PMN-MDSC in patients with cancer is associated with poor prognosis and therapeutic outcomes (17). Therefore, targeting PMN-MDSC in cancer is an attractive therapeutic goal. The main currently unresolved challenge is how to selectively target PMN-MDSC but not PMN to avoid serious toxicity. The fact that PMN-MDSC and PMN share many functional features and receptors make this task especially difficult.

Citrullination is a posttranslational modification of proteins catalyzed by calcium-dependent peptidylarginine deiminase (PAD) enzymes, which hydrolyze a guanidino group of the arginine into a urea group, resulting in a 1 Da change in molecular mass and converting a positively charged arginine into a neutral citrulline. This reaction affects hydrophobicity, protein structure, protein–protein interactions, and substrate receptivity to other posttranslational modifications (18). The PAD family comprises five members (PAD1–4 and PAD6) encoded by distinct genes (19) that have different tissue expression patterns and substrate specificity. PADs are involved in various physiologic processes, including skin cornification, myelin sheath maintenance, and gene expression regulation (18). Excessive citrullination has been implicated in the pathogenesis of several autoimmune diseases mainly rheumatoid arthritis and lupus as well as in other pathologies including multiple sclerosis, psoriasis, Alzheimer's disease, and cancer (20). PADs may be expressed by some cancer cells and usually function in the cytoplasm or nucleus. PAD4 is the only member of the PAD enzyme family with a nuclear localization sequence, and its enzymatic activity impacts gene expression by directly citrullinating transcription factors (21, 22) or by regulating histones (23, 24). PAD4 is highly expressed in neutrophils, where it regulates citrullination of histone H3 and H4. Citrullinated histones H3 has been implicated in the formation of neutrophil extracellular traps (NET), extracellular structures formed by chromatin complexed with intracellular neutrophil proteins expelled from the neutrophils in a process called NETosis (25). An emerging role of NETs in cancer progression has been well documented (26). However, besides NETosis, the role of neutrophil PAD4 in cancer remains largely unknown.

Here, we uncovered a novel mechanism by which PAD4 in neutrophils promotes cancer progression. We found that neutrophil PAD4 regulates neutrophil trafficking, an effect mediated by transcriptional regulation of CXCR2 and specific for the PAD4 member of the PAD family. We also showed that targeted deletion of PAD4 in neutrophils, or pharmacologic inhibition of PAD4 activity with a novel selective PAD4 inhibitor JBI-589 reduced both primary tumor and metastases and significantly enhanced the effect of checkpoint inhibitors. Thus, our data indicate that PAD4 may be a promising therapeutic target for treatment of patients with cancer.

Collection of peripheral blood (PB) samples from patients with lung and colon cancer and healthy donors was approved by the Institutional Review Boards (IRB) of the Christiana Care Health System at the Helen F. Graham Cancer Center and the Wistar Institute. Collection of PB samples from patients with multiple myeloma was approved by the IRB of the University of Pennsylvania and the Wistar Institute. Written informed consent was obtained from all patients. The studies were conducted in accordance with the Declaration of Helsinki guidelines. Twenty-one patients with previously untreated stage II to IV non–small cell lung cancer (10 males, 10 females, and 1 unreported gender, aged 50 to 94, average age 70.6 ± 10.7), 12 patients with colon cancer (4 males and 8 females, aged 38–79, average age 60.7 ± 13.3), and 3 patients with multiple myeloma (1 male and 2 females, aged 51–78, average age 63.3 ± 11.1) were enrolled in this study.

LLC Lewis lung carcinoma (LL/2, RRID:CVCL_4358), LL/2-Luc2 luciferase expressing Lewis lung carcinoma (referred to as LL2, RRID:CVCL_A4CM), B16F10 melanoma (RRID:CVCL_0159), and EL4 lymphoma (RRID:CVCL_0255) cell lines were purchased from ATCC. Cells were expanded, frozen, and vials were kept in liquid nitrogen. Upon thawing, cells were kept in culture for 4 to 6 weeks. Cells were maintained in DMEM supplemented with 10% FBS and 1% penicillin–streptomycin at 37°C with 5% CO₂ and were Mycoplasma negative as detected using Universal Mycoplasma Detection Kit (ATCC).

All animal experiments were carried out in accordance with institutional guidelines and were approved by the Institutional Animal Care and Use Committee of The Wistar Institute. All mice were housed in pathogen-free conditions. Male and female mice aged 6 to 8 weeks were used in experiments.

C57BL/6 mice were purchased from Charles River Laboratories. Pmel-1 TCR-transgenic mice [B6.Cg-Thy1a/Cy Tg(TcraTcrb)8Rest, strain no. 005023, RRID:MGI:3789507], OT-1 transgenic mice [C57BL/6-Tg(TcraTcrb)1100Mjb/J. strain no. 003831, RRID:IMSR_JAX:003831], PAD4 knockout (KO) mice (B6.Cg-Padi4tm1.1Kmow/J, strain no. 030315, RRID:IMSR_JAX:030315), PAD4^fl/fl mice [B6(Cg)-Padi4tm1.2Kmow/J, strain no. 026708, RRID:IMSR_JAX:026708], and Mrp8creTg mice [B6.Cg-Tg(S100A8-cre,-EGFP)1Ilw/J, strain no. 021614, RRID:IMSR_JAX:021614] were purchased from The Jackson Laboratory. PAD4^fl/fl and Mrp8creTg mice were crossed to generate PAD4^fl/flMrp8Cre⁺ mice and their PAD4^fl/flMrp8Cre^– littermates control. PAD2 KO mice (27) were backcrossed 10 generations to the C57Bl/6 background prior to use. RET melanoma were obtained from Dr. Victor Umansky (German Cancer Center, Heidelberg, Germany) and KPC pancreatic cancer model from Dr. Robert H. Vonderheide (University of Pennsylvania, Philadelphia, PA).

Tumors were established by subcutaneous injection of 0.3 × 10⁶ LL2 cells, 0.3 × 10⁶ B16F10, or 0.5× 10⁶ EL4 cells into the right flank of the mouse. Disseminated B16F10 model was established by intravenous injection of 0.5 × 10⁶ B16F10 cells into tail vein. LL2 lung metastasis were evaluated using Perkin Elmer (Caliper) IVIS Spectrum In Vivo System.

For CD8 depletion, mice were treated with anti-CD8 antibody (clone 2.43, BioXCell, RRID: AB_1125541, 200 μg/mouse, i.p.) administered every 3 days beginning 24 hours prior to tumor cell injection. JBI-589 was administered at a dose of 50 mg/kg twice every day, daily, via oral gavage. PD-1 antibody (clone RMP1-14, BioXCell, RRID: AB_10949053, 200 μg/mouse, i.p.) and CTLA4 antibody (clone 9D9, BioXCell, RID: AB_10949609, 200 μg/mouse, i.p.) were administered twice a week beginning on day 7 to 9 after tumor cell injection for a total of four injections.

For evaluation of cell proliferation, mice were injected with BrdU (100 mg/kg, i.p.). Mice were euthanized in 12 hours, spleens were harvested, and single cell suspension prepared. Proliferation of selected cell populations was evaluated after labeling with surface markers and staining with FITC-BrdU Flow Kit (BD Biosciences).

JBI-589, (R)-(3-aminopiperidin-1-yl)(2-(1-(4-fluorobenzyl)-1H-indol-2-yl)-3-methylimidazo[1,2-a]pyridin-7-yl)methanone, was provided by Jubilant Therapeutics. JBI-589 has a molecular weight of 481.57. The purity of the compound was 97.8% as evaluated by HPLC. The detailed JBI-589 synthesis protocol is described in the patent WO 2019/077631. All other antibodies and reagents used are described in Supplementary Tables S1 and S2.

Pharmacokinetic (PK) studies in mice were approved by the Institutional Animal Ethical Committee of Jubilant Biosys (IAEC/JDC/2019/188R). Balb/c male mice were obtained from Vivo Biotech. Mice used in experiments were 6 to 8 weeks old and weighted 22 to 25 g. Mice received JBI-589 at a dose of 10 mg/kg via oral gavage. JBI-589 was given in suspension formulation prepared using Tween-80 and 0.5% methyl cellulose. Sparse sampling was done at 0.25, 0.5, 1, 2, 4, 8, 10, and 24 hours; at each time point three mice were used for blood sampling (∼100 μL). Blood samples were collected in tubes containing K2. EDTA as anticoagulant and centrifuged for 5 minutes at 10,000 rpm at +4°C to obtain plasma.

Mouse neutrophils were isolated from the BM and spleen using MACS technique. Briefly, single cell suspensions were prepared and incubated with anti-Ly6G MicroBeads (Miltenyi Biotec) followed by selection of Ly6G⁺ cells using LS columns. The purity of isolated Ly6G⁺ cells was more than 95% as detected by flow cytometry. All Ly6G⁺ cells expressed CD11b marker. Mouse CD11b⁺Ly6G^lo/–Ly6C⁺ M-MDSC were isolated from spleens by flow sort. For isolation of tumor-associated myeloid cells, single cell suspension was prepared using Tumor Dissociation Kit (Miltenyi Biotec). CD11b⁺Ly6G^–F4/80⁺ tumor-associated macrophages (TAM) and CD11b⁺Ly6G⁺Ly6C^– tumor-associated neutrophils were isolated by flow sort. Mouse hematopoietic progenitor cells (HPC) were isolated from the BM using a Lineage Depletion Kit (Miltenyi Biotec) according to the manufacturer's instructions.

Neutrophils were isolated from PB by centrifugation over a double density gradient Histopaque (Sigma) and collected from the interphase between Histopaque −1.077 and Histopaque −1.119. Cells were washed with DPBS and red blood cells were lysed using RBC lysis buffer (Thermo Fisher Scientific). Neutrophil purity was confirmed by flow cytometry staining with anti-CD15 antibody.

Some experiments required comparison between PB neutrophils (PMN) and PMN-MDSC from the same patient with cancer. For that, PMN-MDSC were isolated from the mononuclear fraction and neutrophils were isolated from granulocyte fraction obtained after Histopaque density gradient centrifugation using positive selection with CD15 MicroBeads (Miltenyi Biotech).

Tumor explant supernatants (TES) were prepared from excised LLC tumors of ∼1.5 cm in diameter. Tumor tissue was minced into small pieces less than 3 mm in diameter and resuspended in RPMI1640 medium supplemented with 10% FBS. Supernatants were collected after overnight incubation at 37°C in 5% CO₂, passed through 0.22 μm filters (EMD Millipore) and kept at −80°C.

Mouse HPC were isolated from the BM and cultured in complete RPMI1640 medium in the presence of GM-CSF (20 ng/mL) and 25% TES prepared as described above. Cells were collected on day 5 and Ly6G⁺ neutrophils were isolated using MACS technique.

Neutrophil chemotaxis was measured using 3 μm pore transwell system (Neuro Probe Inc. or Sigma-Aldrich). Advanced RPMI1640 medium with CXCL1 (BioLegend) or fMLP (Sigma-Aldrich) was placed in the bottom of the well whereas neutrophils (0.1 × 10⁶ cells) resuspended in Advanced RPMI1640 medium were placed on the top of Transwell insert. In some experiments, neutrophils were preincubated with 10 μmol/L GSK484 (Cayman) for 30 minutes prior to being placed on the top of the Transwell insert. Plates were kept for at 37°C in 5% CO₂ for 1 hour followed by counting migrated cells in the bottom well.

Full-length recombinant human PAD4 (125 nmol/L) was incubated with different concentrations of JBI-589 for 1 hour at room temperature followed by 1 hour incubation with 1.5 mmol/L α-N-benzoyl-l-arginine ethyl ester (BAEE) substrate. Ammonia produced as a result of deimination of BAEE was detected after addition of ortho-phthaladehyde, resulting in a fluorescent product. Fluorescence was measured at an excitation 410 nm and an emission 470 nm. IC₅₀ values were determined by sigmoidal dose–response curve (variable slope). Activity of JBI-589 against PAD1 and PAD2 enzymes was measured using a similar assay. Recombinant PAD1 and PAD2 were used at the concentration of 100 and 200 nmol/L, respectively.

Single cell suspension was prepared for flow cytometry analysis. For tumor tissues, Mouse Tumor Dissociation Kit (Miltenyi Biotech) was used to prepare single cell suspension. For surface staining, cells were incubated with Fc-block for 10 minutes followed by labeling with indicated specific antibodies or isotype controls on ice for 20 minutes at 4°C in dark and stained with Live/Dead Fixable Aqua Dead Cell Stain Kit (Invitrogen). Fixation/Permeabilization Solution Kit (BD Biosciences) was used to process cells for intracellular staining. For detection of apoptosis, Annexin V/DAPI staining was performed according to the manufacturer's protocol (BioLegend). At least 10,000 events were acquired using LSRII flow cytometer (BD Biosciences). Analyses was performed using FlowJo software (BD Biosciences).

The level of ROS production in neutrophils was assessed using CM-H₂DCFDA (Invitrogen). Briefly, splenocytes were labeled with surface markers, washed with PBS, resuspended in advanced RPMI1640 medium, and stimulated with 200 nmol/L phorbol 12-myristate 13-acetate (PMA) or 100 nmol/L ionomycin for 30 minutes at 37°C. Cells were then washed and incubated with 5 μmol/L CM-H₂DCFDA for 30 minutes followed by staining with Live/Dead Aqua and analysis using flow cytometry.

RNA was extracted using E.Z.N.A. Total RNA Purification Kit (Omega Bio-Tek) and cDNA was synthesized using High Capacity cDNA Reverse Transcription Kit (Applied Biosystems). qPCR was performed using SYBR Green PCR Master Mix and primers listed in the Supplementary Table S3 using ABI 7500 FAST instrument (Applied Biosystems). Expression of specific genes was normalized to the expression of a housekeeping gene β-actin.

Cells were lysed using RIPA buffer (Sigma-Aldrich) supplemented with Halt Protease and Phosphatase Inhibitor cocktail (Thermo Fisher Scientific). Lysates were subjected to 10% SDS–PAGE, followed by transfer to nitrocellulose membrane (Bio-Rad). Membranes were blocked with 5% nonfat dry milk and incubated with primary antibodies overnight at +4°C followed by incubation with secondary HRP-conjugated antibodies for 1 hour at room temperature. The following primary antibodies were used: anti-PAD4 (Antibodies Online), anti-Histone H3 (Cell Signaling Technology), anti-Histone H3 (citrulline R2+R8+R17, Abcam), and anti-β-actin (Santa Cruz Biotechnology). Densitometry was performed using ImageJ software (NIH, RRID:SCR_003070).

Neutrophils were resuspended in cell lysis buffer (Sigma-Aldrich) at 1 × 10⁸ cells/mL. Lysates were incubated with 10 mmol/L BAEE substrate for 30 minutes at 37°C followed by addition of color developing reagent [13.0% (v/v) phosphoric acid, 25.27% (v/v) sulfuric acid, 0.573 mg/mL ammonium iron sulfate, 60 mmol/L DAMO, 1.5 mmol/L thiosemicarbazide] and heating at 95°C to 100°C for 15 minutes. After cooling, 200 μL of the mixture was loaded per well of 96-well plate and absorbance was measured at 540 nm.

Single cell suspension was prepared from spleens or tumor tissues. TAMs, M-MDSC, or PMN-MDSC were isolated, plated in U-bottom 96-well plates in triplicate in complete RPMI1640 medium, and were co-cultured at different ratios with Cell Trace Violet-labeled splenocytes from either Pmel-1 transgenic mice in the presence of 1 ng/mL cognate peptide EGSRNQDWL or OT-1 transgenic mice in the presence of 0.025 ng/mL cognate peptide SIINFEKL for 48 hours. Proliferation of CD8⁺ T cells was measured using flow cytometry.

Splenocytes from C57Bl/6 wild-type (WT) and PAD4 KO mice were labeled with CellTrace Far Red dye followed by plating and stimulation with CD3/CD28 Dynabeads (Thermo Fisher Scientific) for 48 hours. Mononuclear cells (MNC) were isolated from PB of healthy donors or patients with multiple myeloma using Ficoll-Paque gradient centrifugation. PB MNC were labeled with CellTrace Far Red dye, pretreated for 2 hours with 10 μmol/L GSK-484 or vehicle control (DMSO), and stimulated with CD3/CD28 Dynabeads for 72 hours. Cell proliferation was evaluated by flow cytometry in gated population of CD4⁺ and CD8⁺ T cells.

In parallel, human PB MNC pretreated with 10 μmol/L GSK484 or vehicle control were stimulated for 5 hours with the Cell Stimulation Cocktail with or without protein transport inhibitors (Thermo Fisher Scientific) according to the manufacturer's protocol. Expression of IL2 and IFNγ was evaluated by flow cytometry in gated population of CD4⁺ and CD8⁺ T cells.

Mouse neutrophils were kept untreated or cultured in the presence of LL2 supernatant or calcium ionophore overnight, then were fixed with 4% paraformaldehyde followed by staining with Sytox Green Nucleic Acid Stain at a final concentration of 250 nmol/L. A Nikon Te300 inverted microscope equipped with a motorized XY stage was used to image NETs. Nikon Nis-Elements Ar software was used for image acquisition and was set-up to acquire 25 random locations per well. The z-stacks per location were then combined into an extended depth focused image. The total NET area was calculated by segmenting each image using a defined threshold pixel intensity setting. The spot detection tool in NIS-Elements Ar was used to count the number of cells per field. The sum of the total NET area in the 25 random fields of view was divided by the total number of cells in the 25 fields of view to obtain NET area (μM²/cell).

Statistical analysis was performed using GraphPad Prism (RRID:SCR_002798) software version 6.0. Differences between two experimental groups were determined using two-tailed Student t test. Differences in tumor growth experiments were analyzed using two-way ANOVA. P value of less than 0.05 was considered statistically significant.

The data generated in this study are available within the article and its supplementary data files.

Mouse neutrophils were defined as CD11b⁺Ly6C^loLy6G⁺ cells. In tumor-free mice, genes encoding PAD proteins, Padi2 and Padi4, were highly expressed in BM neutrophils, whereas levels of Padi1, Padi3, and Padi6 were undetectable (Supplementary Fig. S1A). In different mouse tumor models, the expression of Padi2 and Padi4 remained unchanged in BM neutrophils. These models included transplantable subcutaneous LLC lung carcinoma, B16F10 melanoma, EL4 lymphoma (Supplementary Fig. S1B), and genetically engineered KPC pancreatic cancer (28) and RET melanoma (Supplementary Fig. S1C; ref. 29). Similar levels of PAD4 protein were detected in PMN from BM and spleen of tumor-free and tumor-bearing (TB) mice (Supplementary Fig. S1D). Growth of LLC tumor was not associated with upregulation of the expression of Padi1, Padi3, and Padi6 in neutrophils. No difference was found in the expression of Padi2 and Padi4 between tumor-associated and BM PMN (Supplementary Fig. S1E). HPCs were differentiated with GM-CSF in the presence or absence of LLC TES. The presence of TES did not affect Padi2 and Padi4 expression in PMN (Supplementary Fig. S1F). Thus, no changes in the gene expression or the protein amount of the members of PAD family were detected in neutrophils in cancer compared with neutrophils in normal mice.

Next, we asked whether tumor growth affected the enzymatic activity of PADs in PMN. Markedly increased PAD activity was detected in BM neutrophils from LL2-, EL4-, B16F10-bearing mice compared with tumor-free mice (Fig. 1A). Pre-incubation of PMN with a selective PAD4 inhibitor GSK484 (30) completely abrogated the tumor-induced upregulation of PAD activity (Fig. 1B). PMN from PAD4KO tumor-free mice had markedly decreased PAD activation compared to WT mice. No difference in PAD activity was observed between PMN from TB and tumor-free PAD4KO mice (Fig. 1C). In contrast, significant PAD activation was detected in PMN from TB PAD2KO mice compared with tumor-free mice. Western blot analysis of PMN cultured overnight in the presence or absence of LLC TES demonstrated that tumor-derived factors induced citrullination of histone H3 at residues R2, 8, and 17 established to be specific substrates for PAD4 (Fig. 1D and E). Taken together, these data demonstrate that tumor growth is associated with activation of PAD4 in PMN.

To investigate the role of neutrophil PAD4 in regulation of tumor progression we generated PAD4^fl/flMRP8Cre⁺ mice that lack expression of PAD4 in PMN (Supplementary Fig. S2A) (31). LL2 and B16F10 models were chosen for our studies as these tumor cells do not express PAD4 (Supplementary Fig. S2B). In PAD4^fl/flMRP8Cre⁺ mice, growth of primary LL2 tumors was modestly but significantly delayed compared to their PAD4^fl/flMRP8Cre^– littermates (Fig. 2A). At the same time, the presence of lung metastases in PAD4^fl/flMRP8Cre⁺ mice was dramatically reduced compared to control (Fig. 2B and C). The proportion and absolute number of neutrophils also were substantially decreased in both primary tumors (Fig. 2D) and lungs (Fig. 2E) of mice with PAD4-deficient neutrophils. Similar results were observed in B16F10 mice: growth of primary B16F10 tumors was significantly delayed (Fig. 2F), the number of metastases, examined visually (Fig. 2G) or histologically (Fig. 2H and I) was dramatically reduced, and the proportion and absolute number of neutrophils in primary tumor (Fig. 2J) and lungs (Fig. 2K) were substantially reduced in PAD4^fl/flMRP8Cre⁺ mice compared with their PAD4^fl/flMRP8Cre^– littermates. In contrast, no changes in the presence of CD11b⁺Ly6C⁺ monocytic cells were observed in the tumor or lung tissues in either model (Supplementary Figs. S3A–S3D).

To alleviate a possible effect of differences in primary tumor size on the presence of lung metastases we used a disseminated tumor model where intravenous injection of B16F10 cells resulted in formation of tumor lesions in the lungs. A dramatic reduction of tumor growth was observed in PAD4^fl/flMRP8Cre⁺ mice compared with control (Supplementary Fig. S4A); this effect was associated with a significant decrease in the presence of neutrophils at the tumor site (Supplementary Fig. S4B).

A similar antitumor effect against both primary tumors and lung metastases was observed in the whole body PAD4KO mice compared to WT controls (Supplementary Figs. S4C and S4D). This effect was not associated with the lack of PAD4 in T cells as PAD4-deficient T cells had a reduced proliferation capacity (Supplementary Fig. S4E). However, observed antitumor effect was abrogated by the treatment with anti-CD8 antibody, indicating an important role played by the immune system in regulating the antitumor response in PAD4KO mice (Supplementary Figs. S4C and S4D).

In contrast to PAD4 KO, a lack of PAD2 did not affect tumor progression. No differences in primary tumor growth or lung metastases were observed between PAD2KO and WT B16F10 (Supplementary Figs. S5A and S5B) or LL2 (Supplementary Figs. S5C and S5D) bearing mice.

Thus, deletion of PAD4 in neutrophils had a strong suppressive effect on the formation of metastatic lesions, which was associated with a decreased presence of neutrophils in both the primary tumor and the lung metastases.

To understand the mechanism by which neutrophil PAD4 regulates tumor progression, we first tested for the presence of myeloid cells in PAD4^fl/flMRP8Cre⁺ mice. No differences in the proportion or absolute number of neutrophils in the BM were found between tumor-free PAD4^fl/flMRP8Cre⁺ mice and their control PAD4^fl/flMRP8Cre^– littermates (Fig. 3A). Similar results were obtained in LLC-bearing mice (Fig. 3B). In spleens, a small increase in neutrophils was found in PAD4^fl/flMRP8Cre⁺ mice, an effect observed in both tumor-free (Fig. 3C) and LLC-bearing (Fig. 3D) mice. No difference was found in the presence of Ly6C⁺Ly6G^– monocytic cells in the BM or spleens of tumor-free or LLC-bearing PAD4^fl/flMRP8Cre⁺ mice and their littermate controls (Supplementary Figs. S6A–S6D). These data suggested that the reduced presence of neutrophils at the tumor and metastatic sites observed in PAD4^fl/flMRP8Cre⁺ TB mice was not due to diminished myelopoiesis in the BM or spleen.

Lack of PAD4 did not affect in vitro differentiation of HPC toward myeloid lineage. As anticipated, differentiation of HPC in the presence of TES led to the accumulation of CD11b⁺Ly6G⁺ cells after 5 days of culture. PAD4 deficiency did not interfere with this process. Pharmacologic blocking of PAD4 activity with GSK484 had no effect on HPC differentiation in the presence or absence of TES (Fig. 3E and F). Furthermore, neither genetic deficiency or pharmacologic blocking of PAD4 interfered with the presence of monocytic cells (Supplementary Figs. S6E and S6F).

As expected, spleen and tumor-associated neutrophils from TB mice had potent immune suppressive activity defining these cells as PMN-MDSC (Supplementary Fig. S7A). Deletion of PAD4 did not change the suppressive activity of PMN-MDSC. No effect of PAD4 deletion was observed on various factors implicated in PMN-MDSC activity: expression of Mpo, Arg1, iNos, Ptgs2, S100a8, S100a9, Mmp9, and Il6 genes (Supplementary Fig. S7B) or ROS production (Supplementary Fig. S7C). Pharmacologic blocking of PAD4 activity with GSK484 during HPC differentiation did not abrogate the immune suppressive activity of Ly6G⁺ cells generated in the presence of TES (Supplementary Fig. S7D). However, PAD4 deficiency reduced the formation of NETs by mouse neutrophils (Supplementary Fig. S7E; refs. 31, 32).

Given that numbers of neutrophils in primary tumors and lungs were substantially reduced in PAD4 KO mice, we hypothesized that the lack of PAD4 in neutrophils may affect their motility and, therefore, impair their recruitment to the tumor site and premetastatic niche. To test this hypothesis, neutrophils were isolated from the BM of WT or PAD4KO tumor-free and LLC- or B16F10-bearing mice and their migration and chemotaxis were assessed using a standard transwell membrane assay. Lack of PAD4 did not affect the ability of neutrophils from tumor-free or TB mice to spontaneously migrate (Fig. 4A) and did not have any effect on migration induced by the chemoattractant fMLP (Fig. 4B). However, substantially reduced migration of PAD4-deficient neutrophils was observed in response to the chemokine CXCL1. This difference was more profound in neutrophils from TB mice (Fig. 4C). Pharmacologic blocking of PAD4 activity with GSK484 did not affect the viability of neutrophils (Supplementary Fig. S8A) but it reduced their CXCL1-induced chemotaxis (Fig. 4D).

We next tested for a mechanism underlying the impaired chemotaxis of PAD4-deficient neutrophils. CXCR2 and CXCR4 are two major receptors responsible for retention of neutrophils in the BM and their release into the circulation and tissue migration. In PAD4-deficient neutrophils, expression of CXCR2, a major receptor for CXCL1, was markedly downregulated at the gene and protein levels (Fig. 4E and F). This downregulation was more profound in neutrophils from TB mice. CXCR4 gene expression was also lower in PAD4-deficient neutrophils (Fig. 4G), but the change in surface expression of this receptor was minor (Fig. 4H).

In vitro, the presence of TES during HPC differentiation stimulated the expression of CXCR2 receptor on neutrophils. Inhibition of PAD4 with GSK484 reduced baseline CXCR2 expression and abrogated upregulation of CXCR2 in the presence TES (Fig. 4I). TES did not affect the expression of CXCR4; GSK484 slightly decreased CXCR4 expression in control and reduced its expression in the presence of TES (Fig. 4J). Thus, it appears that PAD4 regulation of neutrophil migration is mediated by CXCR2.

To determine the clinical relevance of the mouse data, we evaluated PAD4 expression and activity in neutrophils from patients with cancer. A significant increase in PAD4 gene and protein expression (Fig. 5A and B) and marked upregulation of PAD activity (Fig. 5C), were detected in neutrophils from PB of patients with lung and colon cancer compared with neutrophils from healthy donors. These changes were accompanied by markedly increased CXCR2 expression (Fig. 5D). At the same time, expression of CXCR4 was significantly upregulated only in patients with colon cancer but not with lung cancer (Fig. 5D). A direct correlation was observed between PADI4 and CXCR2 expression in neutrophils from patients with cancer (R² = 0.7698, P < 0.0001; Fig. 5E). PMN-MDSC isolated from mononuclear fraction of PB obtained from patients with cancer demonstrated significantly increased PADI4 expression compared with PB PMN from the same patients (Fig. 5F), an effect associated with significant upregulation of CXCR2. However, no changes in CXCR4 expression were observed in PMN-MDSC.

Next, we evaluated the effect of pharmacologic inhibition of PAD4 on CXCR2 and CXCR4 expression and neutrophil chemotaxis. No difference in neutrophil CXCR4 receptor expression was found between cancer patients and healthy donors and treatment with GSK484 did not affect expression (Fig. 5G). Quite differently, the surface level of CXCR2 receptor was highly upregulated on neutrophils from patients with cancer compared with healthy donors. A four-hour treatment with PAD4 inhibitor GSK484 did not affect viability of neutrophils (Supplementary Fig. S8B) but dramatically reduced CXCR2 receptor expression in neutrophils from both healthy donors and patients with cancer (Fig. 5G), resulting in decreased chemotaxis of neutrophils toward IL8 (Fig. 5H). Pharmacologic inhibition of PAD4 in T cells from healthy donors or patients with cancer slightly decreased their proliferation (Supplementary Fig. S9A) and reduced IFNγ production by CD4⁺ T cells (Supplementary Figs. S9B and S9C).

We performed a time-course analysis of CXCR2 and CXCR4 gene and protein expression in neutrophils treated with PAD4 inhibitor. One-hour treatment with GSK484 significantly downregulated CXCR2 gene expression while not affecting the surface level of CXCR2 protein (Fig. 5I). A significant downregulation of CXCR2 receptor was observed only 4 hours after treatment with PAD4 inhibitor. GSK484 did not affect gene or protein expression of CXCR4 (Fig. 5J). Taken together, these data indicate that PAD4 transcriptionally regulates CXCR2 expression in neutrophils.

We asked whether deletion of PAD4 in PMN and PMN-MDSC would improve the effect of immune checkpoint blockade (ICB). LL2-bearing PAD4^fl/flMRP8Cre⁺ mice and control PAD4^fl/flMRP8Cre^– littermates were treated with anti-CTLA4 and anti-PD-1 antibodies. As anticipated, primary tumor growth was moderately reduced in mice with PAD4-deficient neutrophils. Similar effect was observed in control mice receiving checkpoint inhibitors. However, in PAD4^fl/flMRP8Cre⁺ mice receiving the ICB, primary tumor growth was dramatically reduced (Fig. 6A; Supplementary Fig. S10A). Lung metastases in PAD4^fl/flMRP8Cre⁺ mice were substantially reduced in comparison to control mice (Fig. 6B). In control mice, treatment with checkpoint inhibitors significantly decreased presence of lung metastases, an effect that was markedly improved in PAD4^fl/flMRP8Cre⁺ mice. Similar effects of ICB on primary tumor growth (Fig. 6C; Supplementary Fig. S10B) and lung metastases (Fig. 6D) were observed in B16F10-bearing PAD4^fl/flMRP8Cre⁺ mice and control PAD4^fl/flMRP8Cre^– littermates. PAD4 deficiency in PMN and PMN-MDSC, or treatment of control mice with ICB, both led to the increased presence of activated CD8⁺ T cells in tumor tissues; this effect was significantly increased when PMN and PMN-MDSC PAD4 deficiency and ICB were combined (Fig. 6E).

We used JBI-589, a novel selective PAD4 inhibitor (Supplementary Figs. S11A; Supplementary Table S4) with an excellent oral bioavailability and potential to be translated into the clinic to investigate the effect of pharmacologic blockade of PAD4 on tumor progression. In vitro, JBI-589 inhibited PMA-induced citrullination of histone H3 in mouse neutrophils in a dose-dependent manner (Supplementary Fig. S11B) without noticeable toxicity (Supplementary Fig. S11C). JBI-589 demonstrated excellent oral bioavailability with maximum concentration in plasma achieved 0.5 hours after oral administration and half-life of 6.3 hours (Supplementary Table S5; Supplementary Fig. S11D). No effect of JBI-589 on viability of LL2 and B16F10 tumor cell lines was observed in vitro (Supplementary Fig. S11E). As anticipated, JBI-589 significantly downregulated CXCR2 expression on neutrophil (Supplementary Fig. S12A) and reduced their CXCL1-induced migration (Supplementary Fig. S12B) in vitro while it has no effect on suppressive activity of Ly6G⁺ cells generated in vitro from HPC in the presence of TES (Supplementary Fig. S12C). Treatment of LL2-bearing and B16F10-bearing mice with JBI-589 markedly reduced primary tumor growth (Fig. 7A; Supplementary Fig. S13). Treatment with ICB led to the inhibition of primary tumor growth, an effect that was significantly enhanced by the addition of JBI-589 (Fig. 7A). Analysis of lungs demonstrated the presence of metastasis in control mice while metastatic lesions were almost undetectable in JBI-589-treated mice (Fig. 7B and C). This effect was accompanied by a significant decrease in the presence of tumor-associated neutrophils (Fig. 7D) and increased infiltration of tumor tissues with activated CD8⁺ and CD4⁺ T cells (Fig. 7E and F). A significant downregulation of CXCR2, but not CXCR4, receptor expression was found in Ly6G⁺ cells from JBI-589-treated mice (Fig. 7G). Treatment with JBI-589 did not affect cellularity of the BM in TB mice but reduced splenocyte number (Supplementary Figs. S14A) due to the decreased presence of CD11b⁺Ly6G⁺ cells, CD11b⁺Ly6C⁺Ly6G^– cells, and macrophages (Supplementary Figs. S14B–S14D). No effect of JBI-589 on the presence of CD8⁺ T cells or NK cells (Supplementary Figs. S14E and S14F) and a significant decrease in Tregs in spleens (Supplementary Fig. S14G) were observed. Presence of different progenitors was assessed in spleens of LL2-bearing mice receiving JBI-589 to exclude a potential inhibitory effect of this drug on spleen myelopoiesis. No differences in frequency of LSK, GMP, CMP, and MEP (Supplementary Fig. S15A) and their proliferation (Supplementary Fig. S15B) were found in spleens of JBI-589 treated and control mice. Because PAD4 has been previously implicated in NET formation, an ability of JBI-589 to block this process in vivo was evaluated. For that, BM and spleen neutrophils isolated from control and JBI-589 treated LL2-bearing mice were restimulated in vitro with LL2 cell supernatant. Although neutrophils from control mice were undergoing extensive NETosis, BM (Supplementary Fig. S15C) and spleen (Supplementary Figs. S15D and S15E) neutrophils from JBI-589-treated mice were unable to produce NETs. As presence of M-MDSC and macrophages were reduced in spleens of JBI-589-treated tumor-bearing mice (Supplementary Figs. S14C and S14D), an effect of PAD4 inhibition of their functional activity was investigated. In vitro treatment of spleen M-MDSC (Supplementary Fig. S16A) or TAMs (Supplementary Fig. S16B) isolated from LL2-bearing mice with JBI-589 did not affect the expression of the genes encoding different soluble factors involved in their protumorigenic effect. Myeloid cells were isolated from spleens and tumors of JBI-589-treated and control mice and their functional activity was evaluated. Pharmacologic inhibition of PAD4 with JBI-589 did not affect expression of the genes encoding soluble factors by spleen and tumor PMN-MDSC (Supplementary Figs. S16C and S16D), spleen M-MDSC (Supplementary Fig. S16E), and TAM (Supplementary Fig. S16F) and did not affect suppressive activity of M-MDSC (Supplementary Fig. S16G), TAM (Supplementary Fig. S16H), and PMN-MDSC (Supplementary Fig. S16I). Taken together, these data demonstrated that targeting neutrophil PAD4 could be a new approach for reducing metastatic disease and improving the efficacy of immunotherapy in patients with cancer.

In this study we identified a novel role of PAD4 in cancer: the regulation of neutrophil migration. Previously, the role of neutrophil PAD4 in cancer has been attributed primarily to its ability to citrullinate histone H3; this led to NETosis, a process of NET formation representing one of the mechanisms of neutrophil death (33). NETs have been detected in tumor-bearing hosts and their presence was associated with cancer progression (26). PAD4 KO mice demonstrated a delay in subcutaneous tumor growth, and this has been linked to an impaired ability of PMN to form NETs (34). However, only a fraction of PMN were shown to undergo NETosis (32), prompting the question of whether PAD4 had other functions in neutrophils.

Using mice with targeted deletion of PAD4 in neutrophils, we confirmed a protumorigenic effect played by PAD4 in these cells. Although PAD4-deficient neutrophils were unable to die via NETosis, we did not find an increased presence of these cells in tumor tissues. In contrast, the number of neutrophils detected in tumor tissues and metastatic site (lungs) was significantly decreased in PAD4 KO mice. This decrease was not due to a decrease in myelopoiesis in mice with PAD4-deficient neutrophils or increased apoptosis. Instead, it was due to dramatically reduced BM PMN migration. Consistently, blocking PAD4 activity with the selective inhibitor GSK484 significantly reduced PMN migration.

CXCR4 and CXCR2 are two major receptors that regulate PMN retention in the BM and release into circulation, respectively. In TB mice, CXCR2 signaling is important for attracting MDSC to the TME; inhibition of CXCR2 suppresses the recruitment of neutrophils into the tumor and premetastatic niche (35, 36). In this study, we found that genetic deletion of PAD4 or pharmacologic inhibition of PAD4 activity dramatically downregulated CXCR2 expression at both RNA and protein levels. In contrast, the effect on CXCR4 expression was minimal. As PAD4 inhibition led first to a decrease in CXCR2 gene expression, followed by downregulation of CXCR2 protein, this suggested that PAD4 regulates CXCR2 on a transcriptional level. The cell surface levels of CXCR2 on neutrophils can be downregulated rapidly by internalization upon ligand binding (37) and by metalloprotease activity following neutrophil stimulation with various nonligand stimuli (38). However, data on transcriptional regulation of CXCR2 are scarce. A recent computational analysis of the CXCR2 promoter predicted potential binding sites for HIF-1 and NF-κB (39). However, this prediction has yet to be confirmed in PMN experimentally. Besides histone H3, a number of PAD4 substrates have been identified including fibronectin, MMP9 (40), and others (41–43). Of these, NF-κB p65 has been reported in neutrophils (44). Citrullination of NF-κB p65 regulated its nuclear translocation and production of cytokines (44). Thus, it is a possible that PAD4 may regulate CXCR2 expression via citrullination of NF-κB. Further studies are needed to clarify the specific factor(s) responsible for transcriptional regulation of CXCR2 by PAD4.

Protein citrullination is mediated by a family of five PAD enzymes. Of them, PAD2 and PAD4 are highly expressed in neutrophils (33). Although a role of PAD2 in cancer cells has been reported (45), and recent data demonstrated that PAD2 is linked to the ability of macrophages to form extracellular traps (46), the contribution of PAD2 expressed by neutrophils in cancer progression remains largely unknown. In our study, the activity of PAD4 was markedly increased in neutrophils from TB mice, but no changes in PAD2 activity were found. In agreement, lack of PAD2 in mice did not affect tumor growth, whereas PAD4 deficiency significantly delayed tumor progression.

In contrast to neutrophils from mice, PMN from patients with cancer showed significantly upregulated expression of PAD4 at the gene and protein levels, compared with healthy donors. Presently, the only transcriptional factor implicated in regulating PAD4 levels is p53. Tanikawa and colleagues demonstrated transactivation of PADI4 through an intronic p53-binding site in the U373MG cell line overexpressing p53 (47). However, it is unclear whether this mechanism occurs in human PMN and whether p53 activation status is altered in PMN from patients with cancer compared with healthy donors. Data supporting this hypothesis demonstrated activation of p53 through phosphorylation at serine 15 in myeloid cells under cellular stress including chronic inflammation (48); additional work is needed to clarify the mechanisms regulating PAD4 expression.

Overcoming PMN-MDSC-mediated immune suppression in the TME is one of the main challenges to generating potent antitumor immune responses. A number of strategies have been proposed for targeting MDSC, however, their therapeutic application remains limited (49). One of the biggest challenges is to selectively target PMN-MDSC but not PMN. CXCR2 is critically important for neutrophil migration to various tissues. However, it is not specific for PMN-MDSC and inhibition of CXCR2 blocks migration of all neutrophils. Because PAD4 is upregulated primarily in PMN-MDSC, its targeting may provide an opportunity for more selective inhibition of PMN-MDSC migration.

In this study, we showed that pharmacologic inhibitors of PAD4, including the newly developed compound JBI-589, activated antitumor immune response and significantly delayed tumor progression. As this antitumor effect was accompanied by a dramatic decrease in lung metastasis, upregulation of CXCR2, reduced PMN-MDSCs in tumors and metastatic sites, activation of T cells, and synergy with immune checkpoint blockade, our data suggest that tumor progression is regulated by PAD4-mediated neutrophil trafficking. Thus, our data describe a novel mechanism of regulation of neutrophil trafficking in cancer and demonstrate that targeting this mechanism has a potent antitumor effect. This opens an opportunity for a novel therapeutic approach targeting PAD4 in patients with solid cancers.

S. Rajagopal reports a patent for WO2019077631 pending. D.T. Vogl reports grants and personal fees from Takeda and Active Biotech and personal fees from Karyopharm, Sanofi/Genzyme, Oncopeptides, GSK, and CSL Behring outside the submitted work. D.I. Gabrilovich reports other support from AstraZeneca outside the submitted work. Y. Nefedova reports grants from NIH/NCI, DoD, and grants and nonfinancial support from Jubilant Therapeutics during the conduct of the study; grants and nonfinancial support from Active Biotech, Buzzard Pharmaceuticals, and Merck; and nonfinancial support from BMS outside the submitted work. No disclosures were reported by the other authors.

H. Deng: Formal analysis, investigation, methodology, writing–original draft. C. Lin: Investigation. L. Garcia-Gerique: Investigation. S. Fu: Investigation. Z. Cruz: Investigation. E.E. Bonner: Investigation. M. Rosenwasser: Investigation. S. Rajagopal: Investigation. M. Sadhu: Investigation. C. Gajendran: Investigation. M. Zainuddin: Investigation. R. Gosu: Investigation. D. Sivanandhan: Resources, formal analysis, writing–original draft. M.A. Shelef: Resources, writing–review and editing. B. Nam: Resources. D.T. Vogl: Resources. D.I. Gabrilovich: Writing–review and editing. Y. Nefedova: Conceptualization, supervision, funding acquisition, writing–original draft, writing–review and editing.

This work was supported by NIH/NCI grant R01CA196788 (to Y. Nefedova, D.T. Vogl), DoD award CA180496 (to Y. Nefedova, D.T. Vogl), and research funding from Jubilant Therapeutics (to Y. Nefedova). Support for the Shared Resources utilized in this study was provided by Cancer Center Support Grant P30CA010815 to The Wistar Institute. The authors thank Dr. Vonderheide for providing cells from KPC mice, Dr. Umansky for providing RET melanoma model, and Dr. Rachel Locke for critical reading and editing the manuscript.

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