apoptotic neutrophil clearance. Nevertheless, our results show that neutrophils accumulate mostly in the centrilobular areas of necrosis (**Figures 1A,B**), while KC mainly localize to liver sinusoids. Finally, the expression of G2A receptor (*Gpr132*), which drives the clearance of dying neutrophils through its binding to lyso-phosphatidylserine (lyso-PS) (61), was significantly higher in Ly6Chi monocytes than in MoMF (*p* = 5.7E−05) and recovering KC (*p* = 1.4E−06). Ly6Chi monocytes were also significantly higher for the bridge molecule thrombospondin-1 (*Thbs1*) compared to MoMF (*p* = 0.009) and KC (*p* = 6.1E−06) (**Figure 7**). While the ablation of Ly6Chi monocytes had no effect on the accumulation of dying or necrotic neutrophils during the necroinflammatory phase of AILI (**Figure 6**), they still may be involved in neutrophil clearance at 24–48 h. Noteworthy, G2A and THBS1 gene expression levels remained significantly higher on MoMF versus recovering KC (*p* = 0.0003 and *p* = 6E−05, respectively), implying on their possible involvement in MoMFgoverned neutrophil clearance during early resolution phase.

#### DISCUSSION

The concerted action of professional phagocytes, including tissue-resident macrophages, recruited monocytes, and neutrophils, is fundamental for the effective elimination of noxious agents and the restoration of tissue homeostasis after injury or infection (28). Our data uncover a new immunoregulatory role for Ly6Chi monocytes and their MoMF progenies by their regulation of neutrophil activity and clearance during AILI. Specifically, we demonstrate that liver-infiltrating Ly6Chi monocytes activate ROS production in neutrophils in a direct manner. Further transcriptomic profiling implies that Ly6Chi monocytes positively regulate neutrophil-mediated phagocytosis and inflammation. It also suggests a role for monocytes in the induction of apoptotic pathways in colocalized neutrophils. At the resolution phase, MoMF play a major role in the clearance of apoptotic neutrophils through their expression of a unique set of apoptotic cell recognition bridge molecules and receptors.

The division of labor between tissue-resident and monocytederived macrophage subsets in the resolution from injuries is under intense investigation (1). While it is well established that tissue-resident macrophages are critically involved in the initial recognition of tissue damage and the subsequent recruitment of inflammatory neutrophils and monocytes (28), emerging evidences in gut (49, 62) and liver (22) inflammation suggest that resident macrophages are also robustly imprinted to resist stimuli associated with acute inflammation. In contrast, monocytes display extreme functional plasticity and their immediate availability in the circulation makes them well-suited for a rapid recruitment and performance of acute effector functions required for promoting the initiation, propagation, and resolution of tissue inflammation (2). Indeed, monocytes were shown to play a critical role in the inflammatory and recovery phases of different tissue-specific injuries (1, 17–20). In liver fibrosis, Ly6Chi monocytes produce proinflammatory mediators that promote hepatic stellate cell activation and fibrosis (7), but subsequently give rise to prorestorative Ly6Clo macrophages (5, 8). Similarly, we have demonstrated in an acute model of AILI that recruited Ly6Chi monocytes differentiate into distinct short-lived prorestorative MoMF that outnumber the resident KC population at the early recovery phase. Transcriptomic profiling revealed that Ly6Chi monocytes activate upon their differentiation into MoMF molecular pathways that are associated with regenerative functions, including among others tissue scavenging, angiogenesis, and ECM remodeling (22).

Here, we provide a more detailed comprehension of the interplay between liver infiltrating monocytes and neutrophils, which spatially and temporally overlap in the centrilobular necrotic areas following AILI. During the initial inflammatory phase, monocytes and neutrophils become the dominant phagocyte subsets in the injured liver. Selective ablation of Ly6Chi monocytes has no effect on neutrophil generation or recruitment, as evident by their similar representation in the circulation and in the liver tissue during the first 24 h. However, in the absence of Ly6Chi monocytes, we observe a significant reduction in neutrophil activation during the inflammatory phase, as manifested by reduced ROS production, a hallmark of neutrophil activation (26–29). We also show reduced expression of NADPH oxidase 2 in the "MC-21 neutrophils," a key mediator of neutrophil-driven oxidative burst (50).  Monocyte stimulation with LPS or with apoptotic bodies of human hepatocytes, the latter as an example of environmental cues that monocytes encounter at the necrotic areas of the injured liver, can both induce significant activation of cocultured neutrophils as manifested by increased ROS production and augmented expression of the neutrophil activation marker CD66b (63). Notably, cell-free supernatants could also activate ROS production in the naïve neutrophils suggesting that monocyte-secreted mediators are likely to regulate such interaction. Of note, our results are in dispute with a previous study performed in a murine model of intestinal parasite infection (64). In that study, the authors elegantly demonstrated that Ly6Chi monocytes shut down ROS production in neutrophils that are recruited to the infected tissue (64). Given the plasticity of monocytes, there might be distinct environmental cues that affect monocyte ability to control neutrophil activation. The nature of the potentially involved monocyte-derived factors remains to be defined, but may include cytokines, and also lipid mediators (65). With respect to the latter, we have previously shown that Ly6Chi

monocytes uniquely express the Ptgs2 gene, which encodes for cyclooxygenase-2 (COX2), and the microsomal PGE synthase-1 gene (*Ptges*), both of which constitute key enzymes in PGE2 synthesis (22).

RNA-seq profiling comparing between neutrophils from normal versus monocyte-deficient livers at the necroinflammatory phase further reinforces our claim that Ly6Chi monocyte-derived signals activate innate immune functions in neutrophils. During tissue injury, recruited neutrophils play key role in the removal of damaged cells and cellular debris and prepare the tissue for regeneration (26–29). We show the inability of neutrophils to upregulate key genes involved with phagocytosis in the absence of Ly6Chi monocytes. Induction of genes related to antigen presentation has also been noted in neutrophils under different forms of activation (51–53). Our results imply that monocyte-derived substances induce the expression of MHCII genes in neutrophils. While these results do not necessarily attribute antigen presentation capabilities to neutrophils, they do provide another marker of neutrophil activation that is reduced in the absence of monocytes. Moreover, neutrophils acquire an anti-inflammatory phenotype in the absence of monocytes. This is manifested by the upregulation of various anti-inflammatory transcription factors and myeloid inhibitory receptors in response to Ly6Chi monocyte ablation. Intriguingly, we show altered expression of genes involved in the uptake of modified lipoproteins, breakdown and storage of triglycerides as well as transport and metabolism of cholesterol. While these changes are well-characterized in macrophages, especially in atherosclerosis, there are only sporadic evidences for similar gene expression alterations in activated neutrophils (51), and their mechanistic involvement in tissue injury and resolution is unclear.

Neutrophil activation has to be tightly controlled to avoid excessive tissue damage. In terms of resolution, apoptosis of neutrophils prevents further neutrophil recruitment and terminates their production of deleterious substances. We demonstrate a profound accumulation of AnnexinV+PI+ late apoptotic neutrophils specifically during the early resolution phase. Previous studies have indicated that ROS and oxidative stress can lead to neutrophil apoptosis through disruption of the mitochondria transmembrane potential [reviewed in Ref. (66)], suggesting that Ly6Chi monocyte-induced ROS production by neutrophils may facilitate their apoptosis. Indeed, we show that in the absence of monocytes, neutrophils exhibit reduction in proapoptotic genes and elevation in cell survival molecules such as BCL21A. Moreover, supernatants from LPS-activated monocytes directly increase the fraction of AnnexinV+ apoptotic neutrophils. The phagocytic removal of apoptotic neutrophils is an additional mechanism to clear effete neutrophils and ultimately facilitate the resolution of inflammation. We show here that concomitantly with the need for clearance of apoptotic neutrophils at the early resolution phase, Ly6Chi monocytes differentiate into MoMF, which become the dominant macrophage subset at this stage. Our gene-expression results further uncover that Ly6Chi monocytes upregulate the expression of various apoptotic cell recognition bridge molecules and receptors upon their differentiation into MoMF, which may qualify the latter for the clearance of apoptotic neutrophils. Notably, even in the absence of MoMF, there is still clearance of late apoptotic neutrophil levels between 48 and 72 h following AILI, suggesting that other cells are also likely to take part in the neutrophil removal process, such as KC, which start repopulating at that time. We show that the recovering KC population also expresses a wide variety of apoptotic cell bridge molecules and receptors, some shared with MoMF and others are unique. Indeed, the combined absence of KC and MoMF results in a marked delay in liver repair, greater than each one alone (24).

We have previously reported that inducible and selective monocyte ablation results in impaired liver regeneration, highlighting MoMF as pivotal players in the resolution from liver injury (22). We showed that already at the necroinflammatory phase, Ly6Chi monocyte ablation aggravates hepatic damage. In contrast, here we demonstrate that Ly6Chi monocyte ablation at both 12 and 24 h following acute AILI has no significant impact on hepatic damage. This is further supported by a similar elevation in the serum levels of ALT and AST liver enzymes during the first 24 h of AILI between PBS and MC-21-treated mice. The discrepancy in these results may be related to the different routes of APAP administration implemented in these studies. As Ly6Chi monocytes express both proinflammatory and restorative genes (22), their ablation may concomitantly interfere with inflammationinduced damage and resolution. Nevertheless, these results suggest that Ly6Chi monocyte-mediated regulation of neutrophil activity, and specifically promotion of ROS production, does not significantly contribute to hepatocyte damage. This is in alignment with a previous study reporting that neutrophils do not contribute to the initiation or progression of hepatic damage during acute AILI (35). Importantly, we corroborate our previous findings (22) demonstrating that ablation of Ly6Chi monocytes and their MoMF descendants impair liver resolution at 48 h following AILI. We also show that their ablation leads to increased accumulation of late apoptotic neutrophils. Therefore, it may be that this delay in neutrophil clearance perpetuates the inflammatory response and interferes with hepatic resolution. Of note, MoMF express large repertoire of prorestorative factors promoting hepatocyte growth, ECM remodeling, and angiogenesis (22). Thus, it is has difficult to determine to what extent the altered neutrophil activity and clearance contribute to the increased liver damage following monocyte and MoMF ablation.

Collectively, our results suggest a sequence of interactions following AILI between Ly6Chi monocytes, MoMF, and neutrophils. We show that Ly6Chi monocytes promote neutrophil activation at the injury site during the initial necroinflammatory phase, an important step for the removal of damaged tissue. Subsequently, ROS production in neutrophils may facilitate their apoptosis and their subsequent clearance by MoMF.

#### AUTHOR CONTRIBUTIONS

The corresponding author (CV) confirms that all authors agree to be accountable for the content of the work. NG and CV designed, performed, and analyzed all experiments and wrote the manuscript. MV, OM, GC, and DR substantially assisted NG with some of the major *in vivo* experiments. LCM performed the RNA library generation for the RNA-seq analysis and MPC and ED analyzed the gene expression RNA-Seq data. EB performed the histopathological assessments. EZ contributed to the design of the original idea in this manuscript. SJ mentored NG together with CV, contributed to the experimental design and analysis and critically reviewed the manuscript.

### ACKNOWLEDGMENTS

We thank Prof. Matthias Mack from the Department of Internal Medicine at Regensburg University (Regensburg, Germany) for the generous gift of the anti-CCR2 MC-21 Ab. We acknowledge Dr. Nathan Gluck for his assistance with the recruitment of healthy human subjects under informed consent. We also thank Michail Kaplan (Sourasky Medical Center) for animal care.

#### FUNDING

The postdoctoral fellowship of NG was supported by the Foundation of the Swiss Friends of the Weizmann Institute of Science. Work in the Varol laboratory has been supported by The Azrieli Foundation Canada-Israel as well as by internal funds in Tel-Aviv Sourasky Medical Center, and the "Aldema" fund in Tel-Aviv University. Work in the Jung laboratory was supported by the U.S.-Israel Binational Science Foundation (BSF).

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at http://journal.frontiersin.org/article/10.3389/fimmu.2017.00626/ full#supplementary-material.

### REFERENCES

Figure S1 | Ablation of Ly6Chi monocytes and their monocyte-derived macrophage descendants does not affect liver damage during the necro-inflammatory phase of acetaminophen-induced liver injury (AILI), but attenuates liver resolution. (A) Hematoxylin and eosin (H&E) of liver sections at 12, 24, and 48 h following AILI. Original magnification ×10. Bars, 200 µm. Note the extended necrotic area at 48 h. (B) Histopathological score at 12, 24, and 48 h following AILI of mice injected with PBS (Red circle) or MC-21 (Blue open square). (C) ALT and (D) AST levels in the blood of MC-21-injected mice compared to control mice at steady state (SS) and 24, 48, 72, and 120 h after AILI. Data were analyzed by unpaired, two-tailed *t*-test and presented as mean ± SEM with significance: \**p* < 0.05. Experiments were repeated at least three times, *n* ≥ 3 mice per group.

#### Figure S2 | Ablation of Ly6Chi monocytes reduces MHCII protein

expression in neutrophils. Flow cytometry analysis showing MHCII expression by Ly6G+ neutrophils extracted from normal (PBS) or Ly6Chi monocyte-ablated livers (MC-21), at 24 h following acetaminophen-induced liver injury. Below, graphical summary of the fraction of MHCII+ neutrophils out of total neutrophils.

#### Movie S1 | Colocalization of liver infiltrating Ly6Chi monocyte-derived

cells and neutrophils. A representing movie of 3D reconstructed confocal images extracted from 20 µm *Z*-stacks at acetaminophen-induced liver injury 48 h livers of Cx3cr1gfp/+ mice. Magnification ×63. The movie demonstrates the close proximity of Ly6Chi monocyte-derived cells (GFP) and Ly6G+ neutrophils (red).

#### Movie S2 | Liver infiltrating Ly6Chi monocyte-derived cells

uptake neutrophils at the resolution phase. A representing movie of 3D reconstructed confocal images extracted from 20 µm *Z*-stacks at acetaminophen-induced liver injury-48 h livers of Cx3cr1gfp/+ mice. Note that some of the Ly6Chi monocyte-derived cells (GFP) have engulfed neutrophils stained for Ly6G (in red). Magnification ×63.

and functional characteristics. *Hepatology* (2013) 57:385–98. doi:10.1002/ hep.26016

monocyte subsets with divergent and complementary functions. *J Exp Med* (2007) 204:3037–47. doi:10.1084/jem.20070885

**Conflict of Interest Statement:** The research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2017 Graubardt, Vugman, Mouhadeb, Caliari, Pasmanik-Chor, Reuveni, Zigmond, Brazowski, David, Chappell-Maor, Jung and Varol. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# The Procoagulant activity of apoptotic cells is Mediated by interaction with Factor Xii

*Aizhen Yang1†, Fengwu Chen1†, Chao He1 , Junsong Zhou1,2, Yi Lu3 , Jihong Dai 2,4, Raymond B. Birge5 and Yi Wu1,2\**

*1Cyrus Tang Hematology Center, Collaborative Innovation Center of Hematology, Soochow University, Suzhou, China, 2 The Sol Sherry Thrombosis Research Center, Temple University School of Medicine, Philadelphia, PA, United States, 3Wuhan Thalys Medical Technology Inc., Wuhan, China, 4Department of Pathology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, United States, 5Department of Microbiology, Biochemistry and Molecular Genetics, Rutgers University-New Jersey Medical School, Newark, NJ, United States*

#### *Edited by:*

*Amiram Ariel, University of Haifa, Israel*

#### *Reviewed by:*

*Michal Amit Rahat, Technion – Israel Institute of Technology, Israel Tal Burstyn-Cohen, Hebrew University of Jerusalem, Israel*

#### *\*Correspondence:*

*Yi Wu<EMAIL_ADDRESS>These authors have contributed equally to this work.*

*†*

#### *Specialty section:*

*This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology*

*Received: 27 July 2017 Accepted: 07 September 2017 Published: 25 September 2017*

#### *Citation:*

*Yang A, Chen F, He C, Zhou J, Lu Y, Dai J, Birge RB and Wu Y (2017) The Procoagulant Activity of Apoptotic Cells Is Mediated by Interaction with Factor XII. Front. Immunol. 8:1188. doi: 10.3389/fimmu.2017.01188*

Apoptotic cells, by externalizing phosphatidylserine (PS) as a hallmark feature, are procoagulant. However, the mechanism by which apoptotic cells activate coagulation system remains unknown. Intrinsic coagulation pathway is initiated by coagulation factor XII (FXII) of contact activation system. The purpose of this study was to determine whether FXII is involved in procoagulant activity of apoptotic cells. Using western blotting and chromogenic substrate assay, we found that incubation with apoptotic cells, but not with viable cells, resulted in rapid cleavage and activation of FXII in the presence of prekallikrein and high molecular weight kininogen (HK), other two components of contact activation system. As detected by flow cytometry, FXII bound to apoptotic cells in a concentration-dependent manner, which was inhibited by annexin V and PS liposome. Direct association of FXII with PS was confirmed in a surface plasmon resonance assay. Clotting time of FXII-deficient plasma induced by apoptotic cells was significantly prolonged, which was fully reversed by replenishment with FXII. Corn trypsin inhibitor, a FXII inhibitor, completely prevented apoptotic cells-induced intrinsic tenase complex formation. Consistently, apoptotic cells significantly increased thrombin production in normal plasma, which was not affected by an inhibitory anti-tissue factor antibody. However, blocking of PS by annexin V, inhibition of FXII, or the deficiency of FXII suppressed apoptotic cells-induced thrombin generation. Addition of purified FXII to FXII-deficient plasma recovered thrombin generation to the normal plasma level. In conclusion, FXII binds to apoptotic cells *via* PS and becomes activated, thereby constituting a novel mechanism mediating the procoagulant activity of apoptotic cells.

#### Keywords: apoptotic cells, factor XII, phosphatidylserine, coagulation, contact activation system

#### INTRODUCTION

Apoptosis, or programmed cell death, under physiologic conditions is an active process that is morphologically and biochemically different from necrosis. Apoptosis can be induced in a variety of pathological disorders, including inflammation, autoimmune diseases, atherosclerosis, tissue injury, and degeneration, as well as during radiation treatment and chemotherapy (1). When a cell undergoes apoptosis, phosphatidylserine (PS) typically becomes exposed on the cell membrane (2). If apoptotic cells are not rapidly cleared, they become procoagulant and are often

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Yang et al. FXII in Apoptotic Cell Procoagulation

associated with thrombotic disorders, such as atherothrombosis and Trousseau syndrome (3–5). Up to date, the mechanisms by which apoptotic cells activate the coagulation system and enhance blood clotting are largely unknown. Tissue factor (TF) is complexed with circulating coagulation factor VII and triggers the cascade that generates thrombin. Although TF is involved in the procoagulant activity of apoptotic cells induced by inflammatory mediators, such as LPS, only a portion of the thrombin generated during apoptosis is attributable to TF (3, 6, 7), suggesting the existence of additional mechanisms.

The plasma contact activation system, also called the intrinsic coagulation pathway, consists of factor XII (FXII), prekallikrein (PK), and high-molecular-weight kininogen (HK) (8, 9). FXII, or Hageman factor, is a zymogen of the serine protease factor XIIa (FXIIa). FXII zymogen is activated by limited proteolysis, involving cleavage of the Arg353-Val354 peptide bond, generating the two-chain molecule FXIIa (10). Two principal modes of FXII activation exist. In the first, FXII is activated by binding to negatively charged surfaces, which induces a conformational change (auto-activation). In the second, other proteases, such as kallikrein, cleave and activate FXII (hetero-activation) (10). FXIIa consists of a heavy chain and a light chain connected by a single disulfide bond between two cysteine residues (Cys340 and Cys367) (10, 11). For the last four decades, it has been known that FXII is activated by a variety of artificial and biological anionic surfaces (11, 12). These observations suggest that the contact activation system plays an important role in the enhancement of coagulation. However, whether FXII is involved in apoptotic cell-mediated procoagulant activity has never been studied.

In this study, we investigated whether FXII participates in the procoagulant activity of apoptotic cells. Our results indicate that FXII binds to apoptotic cells and rapidly becomes cleaved and activated. The binding of FXII to apoptotic cells is mediated by PS, and activation of FXII is required for apoptotic cell-mediated blood clotting and thrombin formation. Our study reveals a novel function for FXII and a new mechanism underlying apoptotic cell-mediated procoagulant activity.

#### MATERIALS AND METHODS

#### Materials

Human FXII, PK, HK, FIX, FX, and FXI were purchased from Enzyme Research Laboratories (South Bend, IN, USA). Recombinant human FVIII was obtained from American Diagnostica, Inc. The EZ-Link® Sulfo-NHS-LC-Biotinylation kit was purchased from Thermo Scientific, Inc. Chromogenic substrate for FXIIa and tissue factor blocking antibody (4501) were purchased from American Diagnostica. The chromogenic substrate for FXa was obtained from Chromogenix. Monoclonal antibody against FXII heavy chain (B7C9) was purchased from Santa Cruz Biotechnology. Monoclonal antibody against FXII light chain (C6B7) and PE-labeled anti-TF antibody were obtained from eBioscience. Polyclonal anti-HK heavy chain antibody was purchased from Abgent. Polyclonal anti-pKal antibody was obtained from Abcam (catalog number 43084). Corn trypsin inhibitor (CTI) was purchased from Merck Chemicals, Ltd. FXII-deficient plasma and pooled human normal plasma were obtained from George King Bio-Medical, Inc. Phosphatidylcholine (PC), PS, 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), and 1-oleoyl-2-{6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl}-snglycero-3-phosphocholine (NBD-PC) were obtained from Avanti-Polar Lipids. The FITC-labeled Annexin V Apoptosis Detection kit was purchased from BD Biosciences. Magnetic annexin V microbeads were obtained from Miltenyi Biotech, Inc. Annexin V was obtained from BD PharMingen. Calf intestinal alkaline phosphatase (CIAP) and DNase were purchased from New England Biolabs. RPMI 1640 medium and fetal bovine serum (FBS) were purchased from Invitrogen. All other reagents were obtained from Sigma-Aldrich unless otherwise specified.

#### Cell Culture and Induction of Apoptosis

The human T lymphoblastoid cell line CCRF-CEM was purchased from American Type Culture Collection (ATCC® Number: CCL-119™). CEM cells were propagated in complete RPMI-1640 medium supplemented with 10% FBS, 1 mM l-glutamine, and 1% penicillin-streptomycin at 37°C in a humidified atmosphere with 5% CO2 (13). To induce apoptosis, cells (2 × 105 /mL) were cultured with 10 µM dexamethasone at 37°C in a humidified atmosphere containing 5% CO2 for 24 h. Apoptosis was verified by the FITC Annexin V Apoptosis Detection kit according to the manufacturer's protocol (14). Apoptotic cells were purified with annexin V magnetic microbeads.

#### Preparation of Liposomes and Phospholipid-Coated Beads

To prepare liposomes containing 100% PC (PC liposomes) or PC and PS at 80:20 mol% (PS liposomes), the appropriate amounts of each phospholipid were dissolved with chloroform in a glass tube. Before use, the phospholipids were dried under nitrogen, suspended in PBS, and sonicated for 3 min (15, 16). The phospholipid concentration was determined by a phosphorus assay. To prepare phospholipid-coated beads, 6.5 mg/mL Nucleosil 120-3 C18 beads (Macherey-Nagel) in 1 mL of chloroform were incubated with 2.5 mg/mL DOPC (PC beads) or mixed DOPC/ PS (80:20 mol%; PS beads). The phospholipid-coated beads were dried under nitrogen, resuspended in PBS, sonicated, and then labeled with 2.5 µM NBD-PC for 20 min at 37°C. After centrifugation at 13,000 × *g* for 10 min, the beads were washed with ice-cold PBS twice to remove remaining NBD-PC, and then resuspended in PBS.

### Flow Cytometric Analysis of FXII Binding to Apoptotic Cells

Human FXII was biotinylated using the EZ-Link® Sulfo-NHS-LC-Biotinylation Kit according to the manufacturer's protocol. Biotin-labeled FXII (B-FXII) at various concentrations was incubated with apoptotic cells at 4°C for 15 min. After washing, the cells were labeled with PE-avidin and analyzed by flow cytometry.

#### Measurement of FXII Activity and Cleavage

Cells were incubated with FXII in the presence or absence of PK and HK in HEPES buffer (137 mM NaCl, 5 mM HEPES, 2.7 mM KCl, 2 mM MgCl2, 0.42 mM NaH2PO4, and 1% BSA, pH 7.5) supplemented with 50 µM ZnCl2 at 37°C for 30 min. After centrifugation at 2,500 rpm for 5 min, the supernatant was collected and a chromogenic substrate, Pefachrome FXIIa (0.5 mM), was added. The optical density of substrate hydrolysis was measured at 405 nm using a spectrophotometer (SpectraMax M5) (17). Cleavage of FXII was also detected by western blotting with a monoclonal antibody against FXII heavy chain. The density of the bands was measured by NIH Image J, and cleavage was defined as ratio of the percentage of cleaved FXIIa (48 kDa)/[uncleaved FXII (80 kDa) + cleaved FXIIa (48 kDa)].

### Surface Plasmon Resonance Assay

The experiments were performed at 25°C using HBS-N buffer (20 nM HEPES and 0.15 M NaCl, pH 7.4) containing 50 µM Zn2+ as a running buffer. PS liposomes (PS:PC = 1:9) were diluted in running buffer and immobilized onto flow cell 2 (FL2) of the L1-sensor chip, and PC liposomes were immobilized onto flow cell 1 (FL1) of the L1-sensor chip. Subsequent measurements were obtained at a flow rate of 30 µL/min. A two-fold dilution series of FXII diluted in running buffer was generated (0, 2.5, 5, 10, 20, 40, 80, 100, 200, 400 nM) and was injected over the flow cells at a flow rate of 10 µL/min for 120 s and dissociated for 300 s in order of increasing concentration. Response to PS liposome binding curves was obtained by subtracting the FL2 curve from the FL1 curve and analysis using BIAevaluation software.

#### Intrinsic Tenase Complex Activity

Apoptotic cells were incubated with 200 µL of HEPES buffer containing 2.5 mM CaCl2, 95 nM purified human FXII, 30 nM PK, 30 nM HK, 5.8 nM purified human FXI and FIX, 0.25 nM purified human FVIII, and (where indicated) 2 µM CTI at 37°C. Then, the reaction was started by the addition of purified human FX (170 nM). At various time points, a 25-µL aliquot of the mixture was removed, and 5 µL of 60 mM EDTA in PBS was added to stop FXa formation. FXa formation was monitored as the hydrolysis of the chromogenic substrate S-2222 (0.2 mM) over 30 min. Optical density at 405 nM was converted to FXa nM using a dilution curve of human FXa.

### Clotting Time Assay

Blood drawn from drug-free healthy volunteers was anticoagulated by adding 1 part sodium citrate (110 mM) to 9 parts whole blood. Our study using blood from healthy volunteers was performed after approval by the IRBs of Temple University (IRB no. 20857) and Soochow University (IRB no. 2012037), obtaining informed consent, and in accordance with the Declaration of Helsinki. Platelet-poor plasma (PPP) was prepared by centrifugation at 2,000 × *g* for 30 min. In some experiments, commercial FXII-depleted plasma and normal plasma were used to examine the role of FXII in apoptotic cell-mediated clotting and thrombin generation. After viable or apoptotic cells were mixed with plasma and incubated at 37°C for 180 s, 50 µL of warmed 20 mM CaCl2 was added to start the reaction, and clotting time was immediately recorded with an Amelung KC4A coagulometer (18).

# Thrombin Generation Assay

Thrombin generation in plasma was measured over time with a coagulation analyzer Ceveron® alpha (Technoclone, Vienna, Austria), which employs a fluorogenic thrombin substrate (Z-Gly-Gly-Arg-AMC) to continuously monitor thrombin activity in plasma. Measurements were conducted in a total volume of 150 µL, including 40 µL of normal plasma or FXII-deficient plasma. Apoptotic cells were pretreated with 5 µg/mL BSA, annexin V, mouse IgG, and anti-TF antibody (4501) as indicated. The plasma was treated with mouse IgG and anti-FXII antibody (C6B7) when there was a need to block FXII. Then, an 80-µL aliquot of cells (1 × 105 ) was added to a 40-µL plasma sample. After incubation at 37°C for 15 min, 30 µL of fluorogenic buffer (2.5 mM fluorogenic substrate and 87 mM CaCl2) was added to start the thrombin generation assay.

### SDS-PAGE and Western Blotting

Cleavage of FXII into FXIIa was detected by SDS-PAGE (12%) under reducing conditions and immunoblotting. After incubation with apoptotic or viable cells as described above, the cells were pelleted by centrifugation, and samples containing FXII were collected (14). These samples and samples from a cell-free system were mixed with SDS-PAGE loading buffer and heated at 95°C. After the samples were separated by electrophoresis and transferred to a polyvinylidene difluoride membrane (Millipore), the membrane was blocked with 5% non-fat dry milk in blocking buffer. After extensive washing, the immunoblots were incubated for 2 h with the primary antibodies, including monoclonal anti-FXII (B7C9), polyclonal anti-HK heavy chain, and polyclonal anti-pKal antibodies. Antibody binding was detected by IRDye 800-conjugated goat anti-mouse IgG (LI-COR Bioscience) or IRDye 680-conjugated goat anti-rabbit IgG (LI-COR Bioscience) and visualized with the ODYSSEY infrared imaging system (LI-COR).

### Identification of TF Antigen by Flow Cytometry

Cells (1 × 106 /mL) were incubated with PE-conjugated monoclonal TF (CD142) antibody or isotype control IgG1 for 30 min at 4°C in the dark. Cells were resuspended in 400 µL of PBS before analysis. The mean fluorescence intensity of 10,000 events was determined for each sample.

### Statistical Analysis

The results are expressed as the mean ± SEM of at least three experiments. One-way analysis of variance followed by Tukey's test (for multiple groups) or Student's *t*-test (for comparisons between two groups) was used, and a *p* value less than 0.05 was considered statistically significant. Unless stated otherwise, the data shown are from a single experiment that is representative of at least three separate experiments.

# RESULTS

### FXII Binds to Apoptotic Cells *via* Phosphatidylserine (PS)

To determine whether FXII binds to apoptotic cells, apoptotic cells were incubated with B-FXII. As detected by flow cytometry, the binding of B-FXII to apoptotic cells increased as the concentration increased (from 50 to 800 nM), and became saturated at 400 nM (**Figures 1A,B**). However, B-FXII did not bind to viable cells when added at the same concentrations (**Figure 1B**), suggesting that FXII specifically binds to apoptotic cells. PS is a phospholipid component that is usually maintained on the inner-leaflet (the cytosolic side) of the cell membrane by flippase (2). However, when a cell undergoes apoptosis, PS becomes exposed on the surface of the cell (2). Annexin V (or Annexin A5) is a member of the annexin family of intracellular proteins that binds to PS in a calcium-dependent manner. To examine whether FXII binds to apoptotic cells through PS, apoptotic cells were preincubated with annexin V or BSA in the presence or absence of 2.5 mM CaCl2. As shown in **Figure 2A**, preincubation with annexin V markedly inhibited the binding of FXII to apoptotic cells in the presence of 2.5 mM CaCl2, and its inhibitory effect was concentration dependent. In contrast, no inhibition by annexin V was observed in the

Figure 1 | FXII preferentially binds to apoptotic cells. (A) Biotin-labeled FXII (B-FXII) at the indicated concentrations was incubated with apoptotic CEM cells (2 × 105 ) at 4°C for 15 min. After washing, the cells were labeled with PE-avidin and analyzed by flow cytometry. The fluorescence tracings shown are representative data from three independent experiments. (B) Viable CEM cells (VC, closed circles) or apoptotic CEM cells (AC, squares) were incubated with B-FXII at the indicated concentrations at 4°C for 15 min. After washing, the cells were labeled with PE-avidin and analyzed by flow cytometry (*n* = 5). The data were analyzed by Student's *t-*test, \**p* < 0.05; \*\**p* < 0.01; \*\*\**p* < 0.001.

Figure 2 | Binding of FXII to apoptotic cells is dependent on phosphatidylserine (PS). (A) After preincubation with or without annexin V at the indicated concentrations in the presence of 2.5 mM CaCl2 for 20 min, apoptotic CEM cells were labeled with 100 nM B-FXII and PE-avidin as described above. The binding of FXII was analyzed by flow cytometry and indicated as a percentage compared to the binding in the absence of annexin V and CaCl2, which was set to 100% (*n* = 4). \*\*\**p* < 0.001. (B) B-FXII (100 nM) was preincubated with or without 1 mM phosphatidylcholine (PC) or PS liposomes at 4°C for 15 min, and then apoptotic cells were added. After incubation for 60 min, the binding of B-FXII to apoptotic cells was analyzed by flow cytometry as described above, and shown as the mean fluorescence intensity (MFI). \*\**p* < 0.01. (C) FXII binds to phosphatidylserine in a surface plasmon resonance assay. Serial concentrations of FXII were flowed over PS or PC liposomes immobilized on the Biacore sensor chip. The response curve of PS/FXII binding was obtained by subtracting the curve of PC from that of PS. Curves were analyzed with BIAevaluation software. The *K*D was 3.857E-9 M, and the *R*max was 787.0 Ru.

absence of CaCl2 (**Figure 2A**), suggesting that the inhibitory effect of annexin V is specific. In addition, when FXII was preincubated with PS liposomes, its binding to apoptotic cells was significantly reduced by more than 55% (**Figure 2B**). However, preincubation of FXII with PC liposomes did not affect binding to apoptotic cells (**Figure 2B**). To evaluate the binding capacity of FXII to PS liposomes, a Biacore assay was used. The sensorgrams showed an increase in response units that was reflective of PS binding, and the binding response was concentration dependent, with a *K*D of 3.857E-9 M (**Figure 2C**). These results suggest that FXII binds to apoptotic cells through a high-affinity association with PS.

### The Binding of FXII to Apoptotic Cells Leads to Its Cleavage and Activation

We next examined whether FXII binding to apoptotic cells induces its cleavage and activation. FXII is converted to its active form, FXIIa, through auto-activation induced by contact with charged surfaces. Thus, the activation of FXII is dependent on its cleavage (8). As shown by western blotting using an antibody recognizing the heavy chain of FXII (**Figure 3A**, i and ii), incubation with apoptotic cells markedly induced the cleavage of full-length FXII zymogen (80 kDa) to produce a heavy chain fragment (48 kDa). However, FXII was not cleaved when incubated with viable cells (**Figure 3A**). More strikingly, the FXII cleavage induced by contact with apoptotic cells was significantly enhanced in the presence of HK and PK (**Figure 3A**). In the absence of cells, incubation of FXII with HK and PK induced cleavage of a small percentage of FXII (**Figure 3A**), suggesting that FXII may form a complex with HK and PK on the surface of test tubes, thus leading to its cleavage. As shown by densitometric measurement of band intensity, about 85% of the FXII was cleaved in the presence of HK, PK, and apoptotic cells, which was significantly higher than that induced in the presence of viable cells (**Figure 3A**, iii). Consistent with the observations in the presence of apoptotic cells, incubation with PS liposomes induced cleavage of >80% of the added FXII (**Figure 3B**, i and ii). In contrast, <15% of the FXII was cleaved when incubated with PC liposomes (PS liposome vs. PC liposomes, *p* < 0.001; **Figure 3B**). Taken together, PS-mediated FXII binding to apoptotic cells results in its cleavage. To further examine whether the cleaved FXII induced by apoptotic cells is proteolytically active, we performed an assay with a chromogenic substrate. As shown in **Figure 3C**, in the presence of HK and PK, incubation of FXII with apoptotic cells significantly increased FXIIa activity (*p* < 0.001). This increase in activity was completely prevented by the addition of a FXII inhibitor, corn trypsin inhibitor (CTI). This result suggests that binding of FXII to apoptotic cells promotes its activation.

### FXII Is Required for Apoptotic Cell-Mediated Intrinsic Tenase Formation and Procoagulant Activity

The observation that apoptotic cells can activate FXII led us to hypothesize that this FXII activation is involved in apoptotic cell-mediated procoagulant activity. First, we determined whether the extrinsic coagulation pathway is involved in apoptotic cell-mediated procoagulant activity in our assay system. As evaluated by FACS analysis (**Figure 4A**), TF antigen remained undetectable on both viable and apoptotic CEMs. THP-1 served as a TF-positive control. Apoptotic CEM-mediated pro-coagulation is not dependent on the extrinsic coagulation pathway, which allowed us to evaluate the role of FXII in this process. In the clotting assay, the addition of apoptotic cells significantly shortened the clotting time of PPP triggered by the addition of CaCl2, compared with the clotting time for viable cells (151.8 ± 2.1 vs. 281.9 ± 2.5 s, *p* < 0.001; **Figure 4B**). To evaluate the contribution of FXII, we measured the clotting time of PPP with apoptotic cells in the presence and absence of FXII. As shown in **Figure 4C**, compared to the clotting time of normal plasma with apoptotic cells, the clotting time of FXII-deficient plasma was significantly longer (150.8 ± 3.7 vs. 281.4 ± 7.9 s; *p* < 0.001), which was reversed by replenishment with a physiological concentration of purified FXII protein (165.7 ± 6.6 s), suggesting the requirement of FXII for apoptotic cell-mediated pro-coagulation. We next tested the capacity of FXII to initiate tenase formation, which contributes to the procoagulant activity of apoptotic cells. As shown in **Figure 4D**, in the presence of FXII, PK/HK, FIX, and FVIII, apoptotic cells increased the activity of FXa as a function of time, which was 7.3-fold higher than that with viable cells at 30 s. To examine whether apoptotic cell-mediated FX activation was FXII-dependent, we tested the inhibitory effect of CTI and found that 2 µM CTI reduced apoptotic cell-mediated FXa activation to basal levels (**Figure 4D**). The above results demonstrate that FXII is critical for intrinsic tenase complex formation, contributing to the procoagulant activity of apoptotic cells.

## FXII Is Important for Apoptotic Cell-Induced Thrombin Generation

To further determine the contribution of PS to the FXII-dependent procoagulant activity of apoptotic cells, we employed a thrombin generation assay. As shown in **Figure 5A**, apoptotic cells dynamically induced thrombin generation, which was almost entirely inhibited by annexin V (AxV). Importantly, an inhibitory anti-FXII antibody (C6B7) greatly decreased apoptotic cell-mediated thrombin generation, whereas an anti-TF blocking antibody (TF Ab) did not have such an effect (**Figure 5B**). Consistently, apoptotic cells failed to induce thrombin generation in FXIIdeficient plasma. However, the addition of 375 nM purified FXII to FXII-deficient plasma increased thrombin generation (**Figure 5C**). To determine the effect of PS on thrombin generation, PS or PC liposome-coated beads were used in a test of thrombin generation activity. As shown in **Figure 5D**, PS liposomes strongly induced thrombin generation, whereas PC liposomes had only a minor effect. Interestingly, the inhibitory anti-FXII antibody C6B7 greatly diminished PS-induced thrombin generation (**Figure 5D**), suggesting that FXII contributes to apoptotic cell-mediated thrombin generation by interacting with PS. It is important to note that there is no clear distinction between apoptosis and other cell types of cell death, such as necrosis, and all dying cells exhibit increased PS exposure and enhanced coagulation. Nucleic acids and polyphosphate that are released from necrotic cells can be also associated with FXII activation

Figure 3 | Binding of FXII to apoptotic cells mediates its cleavage and activation. (A) As indicated, 95 nM FXII was incubated with viable cells (*via*) or apoptotic cells (ap) at density of 2 × 105 in the presence or absence of 30 nM prekallikrein (PK) and 30 nM high molecular weight kininogen (HK) at 37°C for 30 min. After centrifugation at 2,500 rpm for 5 min, the supernatant was collected and analyzed by western blot with an anti-FXII Ab (i). The levels of FXII, PK, and HK before incubation with cells are shown by western blotting (ii). In three independent experiments, the density of bands was measured by NIH Image J software, and the cleavage of FXII was defined as the ratio of [cleaved FXII chain (48 kDa)]/[uncleaved FXII (80 kDa) plus cleaved FXII chain (48 kDa)] shown as relative band density (iii). \*\**p* < 0.01; \*\*\**p* < 0.001. (B) In a cell-free system, 95 nM FXII was mixed with 30 nM PK and 30 nM HK followed by addition of 50 nM phosphatidylserine (PS) or phosphatidylcholine (PC) liposomes, one half of the mixture was immediately centrifuged (0 min), and the other half was incubated at 37°C for 60 min (60 min). After centrifugation at 55,000 rpm for 5 h, the supernatant was collected for analysis by western blotting using anti-FXII Ab (i, upper panel). The levels of FXII before incubation with liposomes are shown by western blotting (i, lower panel). In three independent experiments, the density of bands was measured, and the cleavage of FXII was calculated as described above and shown as relative band density (ii). \*\*\**p* < 0.001. (C) Binding of FXII to apoptotic cells mediates its activation. As indicated, FXII (95 nM) was incubated with or without apoptotic cells (AC; 2 × 105 ), PK (30 nM) plus HK (30 nM), and corn trypsin inhibitor (CTI) (2 µM) at 37°C for 30 min. After centrifugation at 2,500 rpm for 5 min, the supernatant was collected and factor XIIa (FXIIa) activity was analyzed as the hydrolysis of a chromogenic substrate as described in the Section "Materials and Methods" (*n* = 3). \*\*\**p* < 0.001.

(19, 20). However, after pretreatment with CIAP and DNase, which degrade polyphosphate and nucleic acids, respectively, there was no change in the level of thrombin generation induced by apoptotic cells (**Figure 5E**), indicating that polyphosphates and nucleic acids are not involved in the observed apoptotic cellmediated thrombin generation.

# DISCUSSION

Changes in phospholipid asymmetry, with outer surface exposure of PS, is a fundamental feature of apoptosis. On activated platelets, exposure of PS on the outer leaflet is essential for membrane assembly of the coagulation factor complexes, including the tenase complex, which are necessary for thrombin generation (21). Similarly, upon apoptotic cell death, PS is exposed on the membrane surface; concomitantly, they become procoagulant and activate the coagulation pathway. These enhanced procoagulant activities associated with membrane PS exposure have been widely observed in apoptotic endothelial cells, vascular smooth

#### Figure 4 | Continued

Apoptotic cell-mediated procoagulant activity and intrinsic tenase formation is dependent on FXII activation. (A) Tissue factor (TF) expression as determined by flow cytometry. CEM cells treated with (apoptotic cells) or without (viable cells) 10 µM dexamethasone for 24 h were stained with CD142-PE to analyze TF expression on the cell surface. THP-1 cells were used as a positive control. The background staining with an isotype control is shown in black. (B) After viable cells and apoptotic cells (2 × 105 ) were suspended in 150 µL of platelet-poor plasma, clotting was triggered by the addition of 20 mM CaCl2 and measured as described in the Section "Materials and Methods" (*n* = 3). \*\*\**p* < 0.001. (C) After apoptotic cells (2 × 105 ) were suspended in normal plasma (normal), FXII-deficient plasma [FXII(-)] and FXII-deficient plasma supplemented with 375 nM FXII [FXII(-) + FXII], respectively, clotting was triggered by the addition of 20 mM CaCl2 and measured as described in the Section "Materials and Methods" (*n* = 3). (D) Effect of apoptotic cells on intrinsic tenase complex formation. As indicated, apoptotic or viable cells were incubated with FXII, prekallikrein, high molecular weight kininogen, FXI, FIX, and FVIII. Then FX was added, and tenase complex formation was analyzed using a chromogenic substrate (*n* = 3). Some samples were also treated with 2 µM corn trypsin inhibitor (CTI). \*\*\**p* < 0.001.

muscle cells, lymphocytes, monocytes, and cancer cells. It has been shown that tissue factor does not play a major role in this process, as there is no increase in antigen levels or the functional activity of tissue factor (3). Thus, the connection between apoptotic cells and coagulation was largely unknown. In this study, we showed that the FXII zymogen preferentially binds to apoptotic cells, leading to its rapid cleavage and activation, thereby contributing to apoptotic cell procoagulant activity.

Thrombin generation is initiated by two distinct pathways, and it can be triggered by exposure of blood to either a damaged vessel wall (extrinsic) or blood-borne factors (intrinsic). The intrinsic pathway of coagulation is initiated by FXII in a reaction involving HK and PK. These factors are collectively referred to as the contact activation system. The present findings of a correlation between thrombin generation and apoptosis showed increased intrinsic tenase activity; however, there is no available model that outlines how thrombin formation could be initiated by highly negatively charged cellular surfaces when functional TF is absent. It is known that FXII is activated by a variety of artificial or biological anionic surfaces, such as kaolin (22), ellagic acid (23), polymers (24), nucleotides (25), sulfatides (26), glycosaminoglycans (27), misfolded proteins (28), polyphosphates (29), and collagen (30). In this study, we showed, for the first time, that FXII zymogen directly binds to PS liposomes and PS on apoptotic cells (**Figures 1** and **2**). The binding of FXII to PS mediates its rapid cleavage and activation in the presence of HK and PK (**Figure 3**). These results suggest that PS on apoptotic cells serves as a docking site for FXII binding, which may induce its auto-activation. The PS on apoptotic cells may also recruit HK and PK to form a complex with FXII, thereby activating PK, and activated PK may increase FXII activation. In three assays, including APTT, intrinsic tenase formation, and thrombin generation assays, we employed FXII-deficient plasma and FXII inhibitor and provided evidence that FXII plays an important role in apoptotic cell-mediated coagulation. We further showed that TF is not involved in the FXII-mediated procoagulant activity of apoptotic cells, as a TF Ab did not affect thrombin generation by Figure 4 | Continued

Figure 5 | FXII is important in thrombin generation induced by apoptotic cells. (A–E) Thrombin generation triggered by apoptotic cells in normal plasma was analyzed as described in the Section "Materials and Methods." Representative data are shown on the left, and accumulated data with statistical analysis are shown on the right. (A) Apoptotic cells were pretreated with 5 µg/mL BSA or annexin V, and then used in a thrombin generation assay (*n* = 3). (B) Apoptotic cells were pretreated with an anti-FXII antibody C6B7 or anti-tissue factor (TF) blocking antibody (4501; 5 µg/mL each), and then used in a thrombin generation assay (*n* = 3). (C) The thrombin generation assay was performed in normal plasma, FXII-deficient plasma, and FXII-deficient plasma reconstituted with human FXII (*n* = 3). (D) Phosphatidylcholine (PC) or phosphatidylserine (PS) liposome-coated beads were used to stimulate thrombin generation in normal plasma. Some samples using PS liposome-coated beads were also incubated with 5 µg/mL C6B7 (*n* = 3). (E) Apoptotic cells pretreated with PBS, 100 U/mL Calf intestinal alkaline phosphatase (CIAP), or 0.01% DNase, before thrombin generation was measured. \*\**p* < 0.01; \*\*\**p* < 0.001.

apoptotic cells, which is consistent with the absence of TF expression on the surface of apoptotic cells (**Figure 4A**). In another study, we found that HK binds to apoptotic cells *via* PS, leading to cleavage and the production of bradykinin in the presence of FXII and PK (31). Therefore, PS on apoptotic cells may serve as a novel activator of FXII and a docking site for the assembly of the contact activation system, which may account for the role of FXII in apoptotic cell-mediated procoagulant activity.

This new function of FXII in apoptotic cell-mediated coagulation provides novel insight into the pathology of apoptotic cells. In pathological settings, such as autoimmune disease, chemotherapy, and inflammation, numerous cells frequently undergo apoptosis. Rapid clearance of these apoptotic cells is crucial for maintaining an anti-inflammatory and antithrombotic state (2). However, if these apoptotic cells are not removed efficiently, they may become procoagulant and proinflammatory. For example, in patients with acute leukemia, phagocytes are overwhelmed by the large numbers of apoptotic cells because of uncontrolled leukemic cell proliferation and cytotoxic chemotherapy. In patients with leukemia, coagulation parameters could be upregulated by chemotherapy; however, this cannot account for the hypercoagulable state, as the basal levels of contact activation system components, including FXII, are fairly high (>50 μg/mL). Patients with systemic lupus erythematosus (SLE) often develop microvessel thrombi, concomitant with the accumulation of apoptotic cells (5). The enhanced functional interaction of apoptotic cells with activation of coagulation system likely plays a major role. Kunzelmann et al. (32) demonstrated that malignant hematopoietic cells (HEL cells) trigger blood coagulation through phosphatidylserine exposure (32). When apoptosis occurs in a microenvironment in direct contact with circulating coagulation factors such as FXII, they may contribute to the initiation and enhancement of unique pro-coagulants on the apoptotic cell surface (33). In view of the increasing evidence for the occurrence of vascular cell apoptosis in the above pathological settings, it is important to characterize the mechanism underlying FXII-driven contact

#### REFERENCES

system activation when vascular cells become apoptotic. Because FXII can also activate plasminogen in the fibrinolytic pathway (34), how activated FXII integrates the intrinsic coagulation and fibrinolysis systems on the surface of apoptotic cells is an interesting topic for future investigation.

In conclusion, the current study demonstrates that FXII binds to apoptotic cells *via* PS, leading to activation of FXII. Activated FXII contributes to intrinsic tenase formation and blood clotting. This study not only revealed a novel mechanism underlying apoptotic cell-mediated procoagulant activity but also identified the apoptotic cell membrane as a new activation surface for the assembly and activation of the contact activation system. These findings suggest that the role of the contact activation system in apoptotic cell-related procoagulant events requires further investigation. Moreover, it will be worthwhile to explore whether inhibition of FXII could be a therapeutic strategy for the prevention and treatment of apoptotic cell-associated thrombotic diseases.

#### AUTHOR CONTRIBUTIONS

AY, FC, CH, JZ, and JD performed research, analyzed data. YL contributed critical reagent. RB and YW designed research and interpreted the data. YW wrote the paper.

#### ACKNOWLEDGMENTS

This work was supported by grants from the National Natural Scientific Foundation (91539122, 81770138, 30971491, 81270592, 81301534, and 81670133), the National Institute of Health (AR063290), and the Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD).

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2017 Yang, Chen, He, Zhou, Lu, Dai, Birge and Wu. This is an openaccess article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

*Shira Michaeli1 , Vivian Dakwar 1 , Keren Weidenfeld1 , Ortal Granski 1 , Odelya Gilon1 , Sagie Schif-Zuck 1 , Anatolii Mamchur 2 , Imad Shams2 and Dalit Barkan1 \**

*1Department of Human Biology, University of Haifa, Haifa, Israel, 2Department of Evolutionary and Environmental Biology, Institute of Evolution, University of Haifa, Haifa, Israel*

Different subtypes of macrophages have been shown to participate in different stages

#### *Edited by:*

*Estee Kurant, University of Haifa, Israel*

#### *Reviewed by:*

*Michal Amit Rahat, Technion – Israel Institute of Technology, Israel Hynda K. Kleinman, George Washington University, United States*

> *\*Correspondence: Dalit Barkan<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology*

*Received: 06 November 2017 Accepted: 27 March 2018 Published: 25 April 2018*

#### *Citation:*

*Michaeli S, Dakwar V, Weidenfeld K, Granski O, Gilon O, Schif-Zuck S, Mamchur A, Shams I and Barkan D (2018) Soluble Mediators Produced by Pro-Resolving Macrophages Inhibit Angiogenesis. Front. Immunol. 9:768. doi: 10.3389/fimmu.2018.00768*

of inflammation and tissue repair. In the late stage of tissue repair, the macrophages, following their engulfment of apoptotic neutrophils, acquire a new phenotype termed alternatively activated macrophages. These macrophages produce growth factors, such as vascular endothelial growth factor (VEGF), that facilitate the angiogenic response as part of tissue restoration. Then, in the later stages of tissue healing, capillary regression takes place. It is presently unknown whether macrophages play an antiangiogenic role in the final stages of tissue repair. Here, we examined whether soluble mediators secreted by pro-resolving CD11blow macrophages (Mres) inhibit angiogenesis in the context of the resolution of tissue repair. Our findings indicate that soluble mediators produced by *ex vivo* generated Mres (CM-Mres) attenuate angiogenesis *in vitro* by inhibiting human umbilical vein endothelial cell (HUVEC) proliferation by lowering their cyclin D1 expression. In addition, CM-Mres lowered HUVEC survival by inducing caspase 3/7 activation, and also inhibited VEGFR2 activation *via* VEGF. HUVEC migration and differentiation to tubular-like structure was also inhibited by CM-Mres. Similarly, CM-Mres significantly inhibited neovascularization as depicted *ex vivo* by utilizing the rat aorta ring assay and *in vivo* by utilizing the chick chorioallantoic membrane assay. Notably endostatin, which was shown previously to exert its antiangiogenic effect by inhibiting proliferation, survival, motility, and morphogenesis of endothelial cells *via* inhibition of VEGFR2 activation, is produced by Mres. Taken together, our results suggest that a specialized subset of macrophages that appear during the resolution of inflammation can produce antiangiogenic mediators, such as endostatin. These mediators can halt angiogenesis, thereby restoring tissue structure.

Keywords: pro-resolving macrophages, tissue repair, angiogenesis, antiangiogenic factors, resolution of inflammation, endostatin

#### INTRODUCTION

Inflammation and tissue repair are adaptive responses to tissue damage induced by pathogen infiltration or mechanical or chemical injury. These responses involve sequential stages which are orchestrated by recruitment and activation of various non-hematopoietic and hematopoietic cells such as neutrophils, macrophages, fibroblasts, and endothelial cells (1). The return of the tissue to its homeostatic state is dependent on the tight regulation and final resolution of the inflammatory response and the wound healing processes. However, dysregulated and exaggerated tissue repair that fails to subside and resolve will result in fibrosis and consequently will lead to organ failure (2, 3). Therefore, it is important to understand the contribution of the different cellular mediators in the resolution of inflammation and the various stages of tissue repair.

Macrophages are highly versatile immune cells that can acquire functionally distinct phenotypes (4, 5). Indeed, recent reports suggest the role of specific subpopulations of macrophages in regulating the different stages of tissue repair (6) and resolution of inflammation (7). In the initial stage of the inflammatory response to injury, leukocyte infiltration is followed by recruitment of monocytes to the site of injury. The monocytes differentiate to classically active macrophages also known as M1-like macrophages (pro-inflammatory) (4, 5). The phenotype of M1-like macrophages that engulf apoptotic polymorphonuclear leukocytes cells (PMN) shifts to that of alternatively activated macrophages. The latter are also referred to as M2-like macrophages and are involved in attenuating inflammation and promoting tissue repair (8, 9). Specifically, M2-like macrophages promote tissue repair by secreting growth factors such as transforming growth factor beta-1 which induces myofibroblast differentiation and deposition of extracellular matrix, and vascular endothelial growth factor (VEGF) which promotes angiogenesis (2, 10). Hence, tight temporal regulation of macrophage phenotype is required to promote resolution of inflammation, tissue repair, and reinstatement of homeostasis. The potential role of macrophages in resolving tissue repair and inflammation has been recently described (7, 11). Schif-Zuck et al. characterized a novel subset of pro-resolving macrophages designated CD11blow macrophages that appear during the resolution of zymosan-induced murine peritonitis. These macrophages secrete pro-resolving mediators and are generated *in vivo* and *ex vivo* from M2-like macrophages following the engulfment of apoptotic leukocytes (11). However, these macrophages display a distinct enzyme expression signature from either M1 or M2, are devoid of phagocytic potential, and are prone to migrate to lymphoid tissues. We recently demonstrated that these pro-resolving macrophages can secrete anti-fibrotic mediators, thus preventing the establishment of a fibrotic-like milieu by preventing the expression of type I collagen (Col-I) by activated myofibroblasts (Gilon et al., submitted for publication). Notably, Col-I remodeling and vasculature regression are evident in the late phase of resolution. Furthermore, intraperitoneal zymosan injection is a model of acute inflammation which self-resolves within 48–72 h (12). Hence, we hypothesized that the recently characterized, pro-resolving macrophages secrete antiangiogenic mediators in addition to anti-fibrotic mediators, thus finalizing tissue repair. Here, we demonstrate that *ex vivo* generated pro-resolving CD11blow macrophages (Mres) secrete antiangiogenic mediators such as endostatin thereby inhibiting angiogenesis by endothelial cells.

#### MATERIALS AND METHODS

#### Cell Line Cultures

Human umbilical vein endothelial cells (HUVECs) (kindly provided by Prof. Gera Neufeld, Technion, Israel) were grown on 10 cm plates, coated with 0.2% gelatin in Dulbecco's phosphate-buffered saline (PBS; Biological Industries, Israel) and overlaid with growth medium comprised of Earle's salt base (M-199) medium supplemented with 20% fetal bovine serum (FBS), 1% antibiotics, 1% vitamins, and glutamine (Biological Industries, USA) and freshly added basic fibroblast growth factor (bFGF) (PeproTech, Israel) (5 ng/ml). Jurkat T cells (kindly provided by Prof. Debbie Yablonski, Technion, Israel) were maintained in RPMI-1640 (Gibco–Life Technology, USA) with high glucose, 10% heat inactivated FBS, and 1% antibiotics. All cells were incubated at 37°C, 5% CO2 incubator.

#### Animals

7- to 8-week-old male C57BL/6 mice and 6-week-female Sprague Dawley rats were purchased from Harlan Biotech Israel. All animals were maintained under specific pathogen-free conditions. Care and handling of animals was in compliance with University of Haifa's experimental protocols. This study was carried out in accordance with the recommendations of University of Haifa Animal Ethics Committee guidelines. The protocol was approved by the University of Haifa Animal Ethics Committee.

### *Ex Vivo* Generation of Pro-Resolving CD11blow Macrophages

Male C57BL/6 mice were injected i.p. with zymosan A (1 mg) purchased from Sigma-Aldrich, Israel. After 66 h, peritoneal exudates were collected, and exudate cells were stained with PE-conjugated rat anti-F4/80 (BioLegend Inc., USA). Macrophages were isolated using EasySep PE selection magnetic beads following the manufacturer's instructions (StemCell Technologies, Israel). Isolated macrophages were co-stained with FITC-conjugated rat anti-Ly-6G and PerCP-conjugated rat anti-mouse CD11b (BioLegend Inc., USA) and analyzed by FACSCanto II (BD Biosciences, USA) and the FACSDiva software. Jurkat T cells were treated with 1 µM staurosporine (Sigma-Aldrich, Israel) to induce apoptosis and washed. Then, peritoneal macrophages were incubated in the presence or absence of apoptotic Jurkat T cells [1:5 macrophage to apoptotic cell (AC) ratio]. After 8 h of incubation, the cells were washed with PBS and overlaid with fresh media; RPMI-1640 with high glucose 10% FBS, and 1% antibiotics for additional 12 h of incubation. Next, conditioned media were collected, and the macrophages were further characterized for their conversion to the CD11blow phenotype using flow cytometry.

#### Preparation of the Different Conditioned Media for the Experimental Assays

The following conditioned media, listed below, were diluted with HUVEC Assay Medium (M-199 medium supplemented with 10% FBS, 1% antibiotics, 1% vitamins, 1% glutamine, and freshly added bFGF 5 ng/ml) at a ratio of 1:1. This step was carried out to ensure viability of HUVEC in all the experimental assays described below.

Condition medium (CM): Baseline conditioned media comprise RPMI supplemented with 10% heat inactivated FBS and 1% antibiotics.

CM-Mϕ: Conditioned media collected after 12 h from culture enriched with CD11bhigh macrophages (Mϕ).

CM-Mres: Conditioned media collected after 12 h from culture enriched with CD11blow macrophages (Mres).

CM-AC: Conditioned media of un-engulfed ACs.

#### Proliferation Assay

Human umbilical vein endothelial cell grown in 10 cm plates were treated for 2 h in M-199, 5% FBS, 1% antibiotics, 1% vitamins, and 1% glutamine medium. Next, the above treated cells (3 × 103 cells/well) were cultured in 96-wells plate coated with Cultrex® growth factor-reduced basement membrane extract (BME) (Trevigen Inc., USA) and treated with the different conditioned media. After overnight incubation at 37°C, 5% CO2 incubator Cell Titer 96 AqueousOne Solution cell proliferation assay kit (Promega, USA) was added to the wells for 2 h to measure cell proliferation according to the manufacturer's instructions. The absorbance was recorded at 490 nm.

#### Immunofluorescence Staining

Human umbilical vein endothelial cell cultured in 8-well chamber glass slides coated with BME (Trevigen Inc., USA), as described previously (13), were treated for 5 min with mixture containing 0.1% Triton X-100 and 4% PFA containing 5% sucrose, and fixed for an additional 25 min with 4% PFA containing 5% sucrose. The cells were washed for 10 min with PBS and an additional 15 min with PBS containing 0.05% Tween 20 (PBS-T). Next, fixed cells were blocked with IF buffer (130 mM NaCl, 7 mM Na2HPO4, 3.5 mM NaH2PO4, 7.7 mM NaN3, 0.1% BSA, 0.2% Triton X-100, and 0.05% Tween 20) containing 10% donkey serum for 1 h followed by overnight incubation at 4°C with rabbit anti-activecaspase-3 (1:400) (Cat # 559565; BD Biosciences). The cells were washed three times with PBS for 15 min each and incubated for 1 h with donkey anti-rabbit conjugated to Alexa Fluor® 647 (Invitrogen, USA) at room temperature. Next, the cells were washed as mentioned earlier and mounted with VECTASHIELD mounting medium with 4′,6-diamidino-2-phenylindole (DAPI). Immunofluorescent images were captured by Zeiss LSM 700 confocal laser scanning microscope (magnification 40×).

For F-actin staining, cells were incubated overnight with Alexa Fluor 488 Phalloidin (1:40) (Molecular Probes, USA), washed three times with PBS for 15 min each and mounted with VECTASHIELD mounting medium with DAPI.

#### Caspase 3/7 Activity

Human umbilical vein endothelial cells grown in 10 cm plates were treated for 2 h in M-199, 5% FBS, 1% antibiotics, 1% vitamins, and 1% glutamine medium. Next, the aforementioned treated cells were cultured in 96 wells coated with 50 µl BME (3 × 103 cells/well) and were overlaid with the different conditioned media. After overnight incubation at 37°C, 5% CO2 incubator, Caspase-Glo® reagent (Promega, USA) was added for each well according to the manufacturer's instructions, and plates were incubated at room temperature for 1 h. Luminescence of each sample was measured using a plate-reading infinite M200PRO, TECAN luminometer.

#### Wound Migration Assay

Wound migration assay was performed using a 12-well plate coated with 0.2% gelatin in PBS (10 × 104 HUVEC/well). 17 h post seeding, a wound was induced by mechanical application of a 1,000 µl sterile tip. Images of wound formation and healing were acquired at time 0 and at 2.5, 5.5, 7, and 8.5 h post-induction, using a light inverted microscope magnification 10× (Nikon Eclipse TS100). Analysis of the wound healing was carried using the Nikon A1R confocal laser scanning microscopy software (NIS Elements AR version 4.3, by Nikon). The area of the scratch was quantified and normalized to the area of the scratch at time 0.

#### Time-Lapse Microscopy

Human umbilical vein endothelial cells (6 × 104 ) were plated on top of gelatinized 15 mm glass-bottom cell culture dishes (Nest Scientific USA Inc.) and overlaid with either CM-Mϕ or CM-Mres. Cells were incubated at 37°C, 5% CO2 incubator for 30 min to allow adherence of the cells. Thereafter, cell motility was followed by time-lapse video microscopy using Nikon A1R confocal laser scanning microscope (20× magnification). Differential interference contrast (DIC) microscopy images were acquired every 1 min for a period of 2 h. Motility of the cells for a period of 2 h was measured by determining the average velocity of 10 different cells for each treatment utilizing ImageJ software (with the "win 64" plug).

#### Tube-Formation Assay

Human umbilical vein endothelial cells grown in 10 cm plates were treated for 2 h in M-199, 5% FBS, 1% antibiotics, 1% vitamins, and 1% glutamine medium. These cells were then cultured in eight-chamber glass slides (Lab-TEK® II, Naperville, IL, USA) coated with BME and overlaid with the different conditioned media for 16 h. For positive control M-199, 20% FBS, 1% antibiotics, 1% vitamins, and 1% glutamine medium was applied. Pictures were acquired by Nikon Eclipse TS100 light microscopy (10× magnification), and the number of bifurcations per field was quantified using ImageJ software.

#### Aortic Ring Assay

Aorta ring assay was carried out as previously described (14) with slight modification. Briefly, thoracic aorta rings were prepared from female Sprague Dawley rats according to the protocol by Bellacen and Lewis (14) and placed in 48-well plates coated with 150 µl Cultrex® growth factor-reduced BME. Plates were incubated for 10 min at 37°C in a 5% CO2 incubator. Following incubation, each well was overlaid with an additional 150 µl BME and incubated at 37°C, in a 5% CO2 incubator for 20–30 min. Next, the aorta rings were subjected to the different treatments for a period of 6 days. For a negative control, vascular cell basal medium (ATCC® PCS-100-030™) was applied, and for positive control vascular cell basal medium supplemented with components listed in Table S1 in Supplementary Material was utilized. Images were acquired by Stemi SV 6, ZEISS light microscope (10× magnification). The micro-vessel sprouting area was analyzed by ImageJ software. Sprouting area for each treatment at time point *T* = 0 was subtracted from the sprouting area at 2, 4, and 6 days of treatment.

## Chick Chorioallantoic Membrane (CAM) Assay

The CAM assay was carried out as previously described (15) with slight modification. Embryonated chicken eggs (~30 per treatment) were incubated at 38°C incubator. At day 3, ovalbumin was removed (3 ml per egg), a window was opened [according to the protocol of Ponce and Kleinmann (15)], and inserts treated with the different conditioned media were applied. The inserts were composed of autoclaved filtered paper (5.5 mm in diameter), which were treated either with CM-Mϕ or CM-Mres (10 µl/ insert). Eggs were incubated for additional 48 h at 38°C incubator. Images were acquired by binocular (0.8× magnification) at time 0 and after 48 h incubation with the above inserts. Quantification of blood vessel density was carried out using ImageJ (by using "win 64" plug). For each treatment, an area of 15 cm2 was analyzed with the filter paper at its center. Vascular density measured at 48 h posttreatment was normalized to the vascular density measured at time 0.

# Mouse Angiogenesis Array

Mouse angiogenesis array kit (R&D Biosystems) was used according to the manufacture's instructions using either CM-Mϕ or CM-Mres. Membranes were analyzed using ImageQuant LAS-4000 analyzer (GE Healthcare Life Sciences, Pittsburgh, PA, USA) and "ImageQuant LAS-4000" software (GE Healthcare Life Sciences). Densitometry analysis was performed using ImageQuant total lab-7 (GE Healthcare Life Sciences) image analysis software.

# VEGFR2 Phosphorylation

Confluent HUVEC cells (cultured on gelatin as described earlier) were treated in M-199, 5% FBS, 1% antibiotics, 1% vitamins, and 1% glutamine medium for 12 h. Next, the cells were overlaid with the different conditioned media for 30 min followed by VEGF supplementation (25 ng/ml) for 5 min. Cell pellets were prepared for western blot analysis.

### Western Blot Analysis

Cell pellets were lysed in WCE (whole-cell extract) buffer [25 mM HEPES, pH 7.7, 0.3 M NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 0.1% Triton X-100, 100 µg/ml PMSF, and 25 mM protease inhibitor cocktail (Roche)], and for p-VEGFR2 detection the cell pellets were lysed in WCE buffer supplemented with 20 mM NAF, 2 mM Na3VO4, and 0.5 mM DTT. The proteins from cell lysate or from the conditioned media were separated by SDS-PAGE followed by transfer on to a nitrocellulose membrane. The membranes were blocked with 5% (w/v) non-fat dried skimmed milk powder either in PBS supplemented with 0.05% Tween 20 (PBS-T) for protein detection or for phosphorylated VEGFR2 in Tris-buffered saline supplemented with 0.1% Tween 20 (TBS-T) for 1 h at room temperature. Membrane was then probed either with rabbit anti-GAPDH (1:500), rabbit anti-cyclin D1 (1:200), or rabbit anti-VEGFR2 (1:1,000) (Santa Cruz, Dallas, TX, USA). Rabbit anti-phospho-VEGFR2-Tyr951 (1:500) (Cell Signaling, Danvers, MA, USA), rabbit anti-VEGF (Abcam, Cambridge, UK), or monoclonal anti-endostatin (1:1,000) (Merck, Darmstadt, Germany) at 4°C overnight. Next, the membranes were incubated with the appropriate horseradish peroxidase-conjugated secondary antibodies (1:10,000; Jackson ImmunoResearch Laboratories, West Grove, PA, USA) for 1 h at room temperature and washed 15 min 3× with either PBS-T or TBS-T (for phosphor-protein detection). Western Bright ECL (Advansta, Menlo Park, CA, USA) was added to the membrane for 1 min and analyzed using ImageQuant LAS-4000 analyzer (GE Healthcare Life Sciences, Pittsburgh, PA, USA) and "ImageQuant LAS-4000" software (GE Healthcare Life Sciences). Densitometry analysis was performed using ImageQuant total lab-7 (GE Healthcare Life Sciences) image analysis software.

## Statistical Analysis

Student's unpaired *t-*test was used accordingly. Two-tailed *p* values of 0.05 or less were considered to be statistically significant. Repeated measures ANOVA comparison test was used accordingly. Values of 0.05 or less were considered to be statistically significant.

# RESULTS

### *Ex Vivo* Generation of Secreted Factors of Pro-Resolving CD11blow Macrophages

To this end, peritonitis was induced, and 66 h later peritoneal exudates were collected. The percentage of macrophages (Mϕ) was determined in peritoneal exudates, based on their size and granularity and positive staining for F4/80 as previously described (11). Next, CD11bhigh Mϕ were collected and were either untreated or treated with apoptotic Jurkat cells {a common apoptotic leukocyte target for macrophages in experimental procedures [(11, 16) at a ratio of 1:5 respectively]}. Incubation with ACs resulted in 79% conversion of CD11bhigh-Mϕ to CD11blow-Mϕ compared with untreated macrophages where only 13% of CD11bhigh-Mϕ were converted to CD11blow-Mϕ, as determined by surface expression of CD11b by FACS analysis (Figure S1 in Supplementary Material). Conditioned media were collected after 12 h from cultures enriched with either CD11bhigh macrophages (CM-Mϕ) or pro-resolving CD11blow macrophages (CM-Mres) and from un-engulfed AC (CM-AC).

### Proliferation of HUVECs is Modulated by Factors Secreted by Pro-Resolving CD11blow Macrophages

Angiogenesis, characterized by sprouting of preexisting vasculature to form new vessels, requires several coordinated endothelial cell activities, such as proliferation, migration, and morphogenesis (17, 18). Therefore, initially we determined whether CM-Mres was able to inhibit the proliferation of endothelial cells. To this end, conditioned media (condition medium (CM), CM-Mϕ, CM-Mres, or CM-AC) were overlaid on HUVECs cultured on top of growth factor-reduced reconstituted BME. The proliferation was measured after overnight incubation. One-way repeated measures ANOVA analysis was conducted followed with repeated contrasts to probe the differences between the four groups of treatments. Significant difference was found between the different treatments [*F*(3, 12) = 5.699, *p* = 0.012]. Furthermore, significant difference was found between the treatments CM-Mϕ vs. CM-Mres (*p* = 0.022) (**Figure 1A**). By contrast, no significant difference was found between CM-Mϕ vs. CM-AC (*p* > 0.05). Given that we observed a reduction in cell number of HUVEC treated with CM-Mres, we next tested whether this inhibition is also due to an increase in apoptosis of HUVEC. HUVECs were cultured as described earlier and scored for apoptosis by (1) percentage of TUNEL positive cells (Figure S2 in Supplementary Material), (2) activated caspase 3 detected by immunofluorescence staining (**Figure 1B**), and caspase 3/7 activity (**Figure 1C**). Increase in the percentage of HUVEC positive for TUNEL staining was evident upon treatment with CM-Mres compared with CM-Mϕ (Figure S2 in Supplementary Material). Activated caspase 3 was apparent by immunofluorescence staining (red staining) in HUVEC treated with CM-Mres (**Figure 1B**), whereas no staining was evident in CM-Mϕ. In addition, a significant increase in caspase 3/7 activity was observed in HUVEC treated either with CM-Mres or CM-AC compared with CM-Mϕ as determined by one-way repeated measures ANOVA analysis followed with repeated contrasts. Specifically, significant difference was found between the different treatments [*F*(4, 20) = 4.549, *p* = 0.009]. Furthermore, significant difference was found between CM-Mres compared with CM-Mϕ (*p* = 0.014) (**Figure 1C**) and CM-AC compared with CM-Mϕ (*p* = 0.030). Hence, our results suggest that CM-Mres inhibits proliferation. Of note, activation of caspase 3/7 by CM-Mres may be also attributed to the presence of mediators secreted by un-engulfed ACs.

#### CM-Mres Inhibits the Motility of HUVEC

Migration of vascular endothelial cells plays an important role in angiogenesis (18). Therefore, we tested whether CM-Mres may impact the motility of HUVEC. To this end, wound migration assay was utilized to study the rate of the wound closure in the plate (cell migration toward the wound/scratch) as detailed below. Wounded monolayers of HUVECs were incubated with the different conditioned media (CM, CM-Mϕ, CM-Mres, or CM-AC). The filling of the "wound" was monitored in a period of 2.5–8.5 h by measuring the % of remaining clear surface, compared with the *T* = 0 (scratch initiation). Our results demonstrate that treatment with CM-Mres delayed overtime the closure of the wound (**Figures 2A,B**) compared with treatment with CM-Mϕ. Repeated measures ANOVA and repeated contrasts were conducted with treatment (CM, CM-Mϕ, CM-Mres, and CM-AC) as repeated measures, at time 8.5 h. A significant difference between all treatments was found [*F*(3, 3) = 24.261, *p* = 0.013].

In addition, at 8.5 h posttreatment, CM-Mres significantly inhibited the closure of the wound compared with CM-Mϕ as determined by repeated contrast [*F*(1, 5) = 7.548, *p* = 0.040].

to five replicates. (B) Representative confocal images of HUVEC stained for activated caspase 3 (red) and nuclei (DAPI, blue). Magnification 40×, bar = 50 µm. (C) Caspase 3/7 activity in HUVEC either starved [CM-no-fetal bovine serum (FBS); positive control] or treated with the indicated conditioned media overnight. Columns; mean, bars; STD, *n* = 6 with three replicates for each experiment. One-way repeated measures ANOVA analysis with repeated contrasts, \**p* ≤ 0.05.

cells) over time. Points; mean, bars; STD, *n* = 4. (C) Quantification of the wound closure at 8.5 h post wounding. Columns; mean, bars; STD, *n* = 6, one-way repeated measures ANOVA analysis with repeated contrasts, \**p* ≤ 0.05. (D) Motility of HUVEC cultured on gelatin for a period of 2 h upon treatment with either CM-Mϕ or CM-Mres was monitored by time-lapse microscopy. An average velocity of 10 different cells for each treatment was determined. Columns; mean, bars; STD, *n* = 10 cells for each treatment, *t*-test, \**p* ≤ 0.05, \*\**p* ≤ 0.01.

Specifically, 51% of the original area remained (**Figure 2C**) upon treatment with CM-Mres whereas, 36% of the original area remained upon treatment with CM-Mϕ (**Figure 2C**). Whereas, there was no significant difference between CM-Mϕ compared with CM-AC (*p* > 0.05).

To further validate that the delay in wound closure was due to inhibition of motility of HUVEC, we conducted time-lapse video microcopy for a period of 2 h (DIC images were captured every minute). Indeed, CM-Mres treatment significantly attenuated the motility of HUVEC cultured on gelatin (Video S1 in Supplementary Material; **Figure 2D**) compared with treatment with CM-Mϕ (Video S2 in Supplementary Material; **Figure 2D**). Interestingly, CM-Mϕ altered HUVEC morphology and induced cell expansion and contraction often associated with membrane blebbing upon their movement, whereas treatment with CM-Mres induced a spindle shape morphology and no cell expansion or membrane blebbing was apparent in the migrating HUVEC.

In summary, our findings suggest that CM-Mres inhibits the motility of HUVEC.

### Soluble Mediators Secreted by Pro-Resolving Macrophages Prevent HUVEC Morphogenesis to Tubular Structures

Next, we determined whether soluble mediators secreted by pro-resolving macrophages were able to inhibit the differentiation of endothelial cells to capillary-like networks. To this end, HUVECs were cultured on BME and treated with CM, CM-Mϕ, CM-Mres, CM-AC, or Assay Medium that promotes HUVEC differentiation to tubular structures. After 16–18 h, the extent of HUVEC differentiation to tubular structures was determined by light microscopy (**Figure 3A**). Quantification of the number of bifurcations of vessel-like tubular structures was carried out in three independent fields per each experimental condition using ImageJ software (**Figures 3A,C**). F-actin organization of the tubular-like structures was determined by phalloidin staining (**Figure 3B**).

HUVEC grown on BME with Assay Medium undergo spontaneous alignment into hollow tubes, forming capillary-like networks (19). However, we found that endothelial tubule formation and stability was impaired in the presence of CM-Mres compared with endothelial cells treated with CM, Assay Medium, or CM-Mϕ (**Figures 3A,B**). This was further supported by one-way repeated measures ANOVA analysis demonstrating significant differences between the different treatments [*F*(4, 16) = 9.997, *p* < 0.001]. Utilizing repeated contrasts analysis, we found a significant reduction in the number of bifurcations in the tubular network formed by the endothelial cell treated with CM-Mres in comparison to treatment with CM-Mϕ (*p* = 0.012) (**Figure 3C**). By contrast, there was no significant difference between CM-AC in comparison to CM-Mϕ (*p*> 0.05). The tube-formation assay is based on the ability of endothelial cells to form three-dimensional capillary-like tubular structures in the 3D BME system. In this system, endothelial cells proliferate, differentiate, directionally migrate to align, branch, and form the tubular polygonal networks of blood vessels. This is a well-established assay to study angiogenesis (19, 20). Given that a significant effect on formation of the capillary-like tubular structures was only evident upon treatment with CM-Mres, we proceeded to further validate the antiangiogenic effect of CM-Mres and compared it to CM-Mϕ, as described below.

#### Sprouting Angiogenesis *Ex Vivo* and *In Vivo* are Attenuated by CM-Mres

To further verify our *in vitro* findings, we utilized the rat aorta ring assay as an *ex vivo* model of angiogenesis. This organ culture assay scores for sprouting angiogenesis from the segmented aorta ring cultured on BME. Our results demonstrate that CM-Mϕ promoted neovascularization (see black arrow; **Figure 4A**; Figure S3 in Supplementary Material), whereas, CM-Mres restrained neovascularization (see white arrows; **Figure 4A**). We conducted repeated measures two-way ANOVA analysis, and we found significant difference between the different treatments [*F*(4, 40) = 10.828, *p* = 0.003]. Furthermore, we found significant interactions between time and treatment [*F*(8, 40) = 14.127, *p* < 0.001]. Planned comparisons were carried out on data from day 6 to probe the interaction. We found significant difference between CM-Mres compared with CM-Mϕ [*F*(1, 5) = 55.574, *p*= 0.001]. Furthermore, a significant difference was found between negative control vs. CM [*F*(1, 5) = 6.663, *p* = 0.049]. Whereas, no significant difference was found between CM-Mres vs. negative control [*F*(1, 5) = 4.839, *p* = 0.079] (**Figure 4B**). Hence, CM-Mres contain soluble mediators that can inhibit sprouting angiogenesis in the rat aorta ring assay. Next, we tested whether CM-Mres can inhibit angiogenesis *in vivo* by using the chick CAM assay. Similarly, to the *in vitro* and *ex vivo* results, exposure of the CAM to CM-Mres reduced vessel density (compare vessel density at time *T* = 0 vs. *T* = 48 h post CM-Mres treatment; **Figure 4C**, see white arrows and **Figure 4D**) whereas, treatment with CM-Mϕ for 48 h increased vessel density compared with *T* = 0 (**Figure 4C** see black arrows and **Figure 4D**). Altogether, these results suggest that CM-Mres contains antiangiogenic mediators.

### Increased Levels of the Antiangiogenic Mediator Endostatin and Decrease in Angiogenic Factor VEGF in CM-Mres Compared with CM-M**ϕ**

Angiogenesis is regulated by a balance between angiogenic and antiangiogenic factors. Given that CM-Mres inhibited angiogenesis *in vitro*, *ex vivo*, and *in vivo*, has promoted us to determine the presence and predominance of antiangiogenic mediator/s over angiogenic factors in CM-Mres. To this end, mouse angiogenesis array was utilized to detect pro- and antiangiogenic factors in CM-Mres compared with CM-Mϕ (**Figures 5A–C**). Initial dot blot analysis revealed several pro-angiogenic factors with lower levels in CM-Mres compared with CM-Mϕ such as osteopoetin (21), HGF (22), CXCL16 (23), and CCL2 (24) (**Figure 5B**). Whereas, increase in the levels of the antiangiogenic factors endostatin, PEDF and thrombospondin-2 [reviewed in Nyberg et al. (25)] was observed in CM-Mres compared with their levels in CM-Mϕ (**Figure 5C**). Notably, the levels of the central pro-angiogenic mediator VEGF (26) decreased significantly in CM-Mres compared with its levels in CM-Mϕ (determined by western blot analysis; **Figures 5D,E**), whereas the levels of endostatin; an inhibitor of VEGF mediated signaling (25), was significantly higher in CM-Mres compared with CM-Mϕ (determined by western blot analysis; **Figures 5D,F**). Next, we tested VEGFR2 phosphorylation, given that endostatin was previously shown to block VEGF-induced tyrosine phosphorylation of VEGFR2 in HUVEC (see Materials and Methods) (27). Indeed, VEGFR2 phosphorylation on Y195 was reduced upon treatment with CM-Mres compared with treatment with CM-Mϕ (**Figures 5G–H**). Furthermore, CM-Mres inhibited

significantly Cyclin D1 expression in HUVEC compared with treatment with CM-Mϕ (determined by western blot analysis; **Figures 5I,J**). These results are in accordance with previous studies demonstrating downregulation of Cyclin D1 expression upon endostatin treatment (28). Overall, our results suggest that reduction in the levels of VEGF and increase in endostatin levels in CM-Mres may mediate in part the antiangiogenic effect of CM-Mres by inhibiting VEGFR2 mediated signaling.

#### DISCUSSION

Macrophages play a fundamental role in wound healing by generating bioactive mediators that stimulate angiogenesis and fibroplasia (29, 30). However, the potential role of macrophages in resolving tissue repair by inhibiting angiogenesis is largely unknown (9).

Angiogenesis is a multifaceted process required to facilitate restoration of the damaged tissue during wound healing. This

control), basal media with supplementations (positive control, see Materials and Methods), condition medium (CM), CM-Mϕ, or CM-Mres. Vascularized area is indicated by black arrow heads, and avascular area is indicated by white arrow heads. (B) Quantification of the sprouting area of endothelial cells relative to *T* = 0. Columns; mean, bars; STD, *n* = 2 with three replicates for each treatment, two-way repeated measures ANOVA analysis with repeated contrasts \**p* ≤ 0.05, \*\*\**p* ≤ 0.001. (C) Representative images of the chick CAM treated for 48 h with either CM-Mϕ (*n* = 15) or CM-Mres (*n* = 12).  (D) Quantification of vessel density in panel (C). Columns; mean of the fold change in vessel density from *t* = 0 for each treatment, bars; STD, *t*-test, \*\*\**p* ≤ 0.001.

process is orchestrated by: (1) remodeling of the extracellular matrix, (2) proliferation and migration/chemotaxis of endothelial cells, and (3) assembly of endothelial cells to vessel tube and its stabilization by pericytes and smooth muscle cells (17). In this study, we demonstrate for the first time to our knowledge that mediators generated by pro-resolving CD11blow macrophages (Mres) that participate during resolution of acute murine peritonitis (11) inhibit angiogenesis *in vitro* and *in vivo*. The multiple parameters of angiogenesis, proliferation, viability, and motility, were modulated by CM-Mres thus culminating in overall significant and robust inhibition of the angiogenic process.

*In vitro*, we demonstrated that bioactive mediators produced by Mres inhibited HUVEC proliferation significantly and enhanced their apoptosis. This was depicted by increase in the percentage of cells positive for TUNEL and increase in caspase 3/7 activity. Furthermore, a significant reduction in the migration capacity

Figure 5 | Antiangiogenic factors levels are higher in CM-Mres compared with CM-Mϕ. (A) The Proteome Profiler Mouse Angiogenesis Array Kit (Catalog # ARY015) was used to simultaneously assess the relative levels of 53 mouse angiogenesis-related proteins in either CM-Mϕ (upper panel) or CM-Mres (lower panel). (B) Quantification of pro-angiogenic factors; HGF, osteopontin, CXCL16, and CCL2. (C) Quantification of antiangiogenic factors; endostatin, thrombospondin-2, and PEDF. The histograms (B,C) were generated by quantifying the mean spot pixel density from the arrays using image software analysis. (D) Representative western blot analysis of vascular endothelial growth factor (VEGF) and endostatin levels in CM-Mϕ compared with CM-Mres. (E,F) Quantification of VEGF (E) and endostatin levels (F). Densitometry values in panels (E,F) were normalized to CM-Mϕ. *n* = 3, *t*-test, \**p* < 0.05. (G) Representative western blot of VEGFR2 phosphorylation in human umbilical vein endothelial cell (HUVEC) induced by VEGF (25 ng/ml) in the presence of the different conditioned media and its quantification (H). (H) Densitometry values of p-VEGFR2 were normalized to treatment with condition medium (CM) + VEGF (*n* = 2). Columns; mean, bars; STD. (I) Representative western blot of cyclin D1 expression levels in HUVEC treated with either CM-Mϕ or CM-Mres and its quantification (J). (J) Densitometry values were normalized to treatment with CM-Mϕ. Columns; mean, bars; STD, *n* = 3, *t*-test, \**p* < 0.05.

of the cells was observed upon exposing HUVEC to CM-Mres compared with treatment with CM-Mϕ as determined by the wound migration assay and time-lapse live video microscopy. Furthermore, tubular formation of HUVEC on BME was also significantly reduced upon exposure to CM-Mres in comparison to treatment with CM-Mϕ. This was further supported by a significant reduction in the number of bifurcations in the tubular network formed by the endothelial cells treated with CM-Mres in comparison to treatment with CM-Mϕ. Similarly, CM-Mres restrained vascular outgrowth, whereas CM-Mϕ promoted vascular outgrowth, in the rat aorta ring and CAM model systems. Notably, rat aorta ring assay and CAM assay exhibit multiple cell processes involved in angiogenesis as depicted *in vitro* and allow analysis of angiogenesis in an environment composed of multiple cell types and in the physiological context, respectively (31–33). Hence, our results demonstrate that bioactive mediators generated by Mres can inhibit different stages in the angiogenesis process.

Inhibition of angiogenesis is dependent on tilting the delicate balance between pro- and antiangiogenic factors. Therefore, if antiangiogenic factors predominate the angiogenic factors then angiogenesis will not occur (17, 34). Initial insight into the balance between pro- and antiangiogenic factors in CM-Mres vs. CM-Mϕ was obtained by performing a mouse angiogenesis array. Preliminary results demonstrate reduction in some of the pro-angiogenic growth factors and chemokines such as HGF (22), osteopontin (21), CCL2 (35), and CXCL16 (36) in CM-Mres compared with CM-Mϕ. Notably, these mediators were shown previously to modulate motility proliferation and/or survival of endothelial cells. Furthermore, an increase in some of the antiangiogenic mediators such as PEDF, thrombospondin-2 and endostatin [reviewed in Nyberg et al. (25)] was observed in CM-Mres in comparison to CM-Mϕ. Endostatin is a potent inhibitor of angiogenesis. It is a 20-kDa proteolytic fragment of collagen XVIII, which exerts its antiangiogenic effect by inhibiting VEGF binding to VEGFR2 and thus prevents VEGFR2 phosphorylation and activation (27). Notably, along with the increase in endostatin levels in CM-Mres, VEGF levels were significantly reduced in comparison to CM-Mϕ. Hence, this tilt in the balance between VEGF and endostatin in CM-Mres may have resulted in the antiangiogenic effect of the CM-Mres on HUVEC. To further explore this, we tested whether CM-Mres can inhibit VEGFinduced tyrosine phosphorylation of VEGFR2 in HUVEC (27). Indeed, VEGFR2 phosphorylation on Y195 was reduced upon treatment with CM-Mres compared with treatment with CM-Mϕ. Furthermore, we demonstrated reduction in cyclin D1 expression in HUVEC treated with CM-Mres compared with treatment with CM-Mϕ. Notably, in these experiments HUVECs were exposed to angiogenic factors that were added, for instance, bFGF (supplemented in all prepared conditioned media) or present in the CM-Mϕ, such as VEGF. This is in line with a previous report demonstrating endostatin-induced downregulation of cyclin D1 (28) resulting in G1 arrest of endothelial cells that were either treated with bFGF or VEGF. Similarly, the reduction in motility and increase in caspase 3 activity upon treatment with CM-Mres can also be attributed to endostatin angiostatic activity, as described previously (37). Overall, our results suggest that reduction in the levels of VEGF and increase in endostatin levels in CM-Mres may mediate in part the antiangiogenic effect of CM-Mres by inhibiting VEGFR2-mediated downstream signaling. VEGFR2 activation and induction of its downstream signaling by VEGF is one of the key pathways in the angiogenesis process and wound healing repair (38, 39). Hence, the potential antiangiogenic activity of CM-Mres *via* inhibition of VEGFR2 downstream signaling warrants further future experimentation. Overall, our results suggest that pro-resolving CD11blow macrophages can resolve tissue repair by secreting angiostatic mediators, such as endostatin, thus insuring tissue restoration to its homeostatic state.

### ETHICS STATEMENT

This study was carried out in accordance with the recommendations of University of Haifa Animal Ethics Committee guidelines. The protocol was approved by the University of Haifa Animal Ethics Committee.

### AUTHOR CONTRIBUTIONS

SM, VD, KW, and OGranski performed the experiments and analyzed the data. SM, VD, and KW prepared the figures. SS and OGilon prepared the conditioned media of the macrophages and performed FACS analysis. AM and IS performed the aorta ring assay and its analysis. IS reviewed the manuscript. DB conceived the project, designed the experiments, analyzed the data, and wrote the manuscript.

#### ACKNOWLEDGMENTS

We thank Dr. Amiram Ariel for the use of FACSCanto II (BD) equipment, Dr. Ram Reshef for assisting with the CAM assay and Dr. Haim Shapiro and Shany Barkan for reviewing the manuscript.

### FUNDING

This research was supported by the British and Swiss friends of University of Haifa, University of Haifa seed money (DB), and by John Templeton Foundation (grant # 53057; AM and IS).

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at https://www.frontiersin.org/articles/10.3389/fimmu.2018.00768/ full#supplementary-material.

VIDEO S1 | Time lapse video microscopy of HUVEC treated with CM-Mres.

VIDEO S2 | Time lapse video microscopy of HUVEC treated with CM-Mϕ.

### REFERENCES

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

The handling Editor declared a shared affiliation, though no other collaboration, with the authors.

*Copyright © 2018 Michaeli, Dakwar, Weidenfeld, Granski, Gilon, Schif-Zuck, Mamchur, Shams and Barkan. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# The "Phagocytic Synapse" and Clearance of Apoptotic Cells

*Nicole D. Barth, John A. Marwick, Marc Vendrell, Adriano G. Rossi and Ian Dransfield\**

*MRC Centre for Inflammation Research, Queen's Medical Research Institute, University of Edinburgh, Edinburgh, United Kingdom*

Apoptosis and subsequent phagocytic clearance of apoptotic cells is important for embryonic development, maintenance of tissues that require regular cellular renewal and innate immunity. The timely removal of apoptotic cells prevents progression to secondary necrosis and release of cellular contents, preventing cellular stress and inflammation. In addition, altered phagocyte behavior following apoptotic cell contact and phagocytosis engages an anti-inflammatory phenotype, which impacts upon development and progression of inflammatory and immune responses. Defective apoptotic cell clearance underlies the development of various inflammatory and autoimmune diseases. There is considerable functional redundancy in the receptors that mediate apoptotic cell clearance, highlighting the importance of this process in diverse physiological processes. A single phagocyte may utilize multiple receptor pathways for the efficient capture of apoptotic cells by phagocytes (tethering) and the subsequent initiation of signaling events necessary for internalization. In this review, we will consider the surface alterations and molecular opsonization events associated with apoptosis that may represent a tunable signal that confers distinct intracellular signaling events and hence specific phagocyte responses in a context-dependent manner. Efficient molecular communication between phagocytes and apoptotic targets may require cooperative receptor utilization and the establishment of efferocytic synapse, which acts to stabilize adhesive interactions and facilitate the organization of signaling platforms that are necessary for controlling phagocyte responses.

Keywords: macrophage, phagocytosis, apoptotic cells, cell–cell interactions, phagocytic receptor, phosphatidylserine, opsonin

# INTRODUCTION

Elimination of injured or metabolically stressed cells in multicellular organisms is controlled *via* engagement of apoptotic programs together with efficient tissue clearance mechanisms (1–3). Phagocyte/apoptotic cell interactions also initiate anti-inflammatory reprogramming that regulates inflammation and immunity (4). Deficient clearance of apoptotic cells contributes to the development and/or exacerbation of many autoimmune and inflammatory diseases [reviewed in Ref. (5)].

The diversity of molecular pathways mediating recognition and phagocytosis of apoptotic cells (efferocytosis) reflects the fundamental importance of this process (4). There are several mechanisms by which viable cells avoid phagocytosis (6). However, altered plasma membrane lipid composition (7, 8) and/or oxidation status (9), together with changes in cell surface molecule repertoire and patterns of glycosylation (10) termed "apoptotic cell associated molecular patterns" (11) (**Figure 1**), allow phagocytes to distinguish viable and apoptotic cells. Here, we consider the formation of an "efferocytic synapse" and assembly of molecular platforms that facilitate phagocytosis and subsequent signaling events.

#### *Edited by:*

*Kirsten Lauber, Ludwig-Maximilians-Universität München, Germany*

#### *Reviewed by:*

*Raymond B. Birge, Rutgers University, The State University of New Jersey, United States Michael R. Elliott, University of Rochester, United States*

> *\*Correspondence: Ian Dransfield<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Molecular Innate Immunity, a section of the journal Frontiers in Immunology*

*Received: 28 August 2017 Accepted: 20 November 2017 Published: 04 December 2017*

#### *Citation:*

*Barth ND, Marwick JA, Vendrell M, Rossi AG and Dransfield I (2017) The "Phagocytic Synapse" and Clearance of Apoptotic Cells. Front. Immunol. 8:1708. doi: 10.3389/fimmu.2017.01708*

Figure 1 | Schematic representation of surface molecular changes associated with apoptosis. Reduced cell surface molecule expression may occur through metalloprotease-mediated proteolytic shedding. Reduced receptor expression may act to limit apoptotic cell function and generate a distinct cell surface profile from viable cells. For example, reduced expression of CD47 or disabled CD31 may lead to loss of signals preventing phagocytosis. In addition, loss of integrin regulation may result in functional uncoupling in apoptotic cells, leading to cell detachment. Altered association of cell surface molecules with lipid rafts may alter functional activity, including gain-of-function of some receptors, e.g., FcγRIIa on myeloid cells. Altered carbohydrate processing may result in reduced sialic acid exposure and appearance of accessible mannose residues. Downregulation of complement regulatory molecules (e.g., CD55 and CD46) may lead to opsonization with complement components including C3b. Exposure of anionic phospholipids, including phosphatidylserine (PtdSer), allows binding of a broad range of opsonins to apoptotic cells. Protein S and Gas6 bind to PtdSer in a Ca2+-dependent manner, whereas milk fat globule EGF-factor 8 (MFG-E8) binds independent of Ca2+. Other less well defined apoptotic cell surface changes may allow binding of other opsonins including thrombospondin, C-reactive protein, and surfactant protein A. Finally, proteins with intracellular localizations may appear on the surface of apoptotic cells, including heat-shock proteins and calreticulin. Apoptotic cell surface molecules are shaded blue whereas apoptotic cell opsonins are shaded green.

## PHOSPHATIDYLSERINE (PtdSer) AS A LIGAND FOR APOPTOTIC CELL RECOGNITION

A near universal membrane alteration associated with apoptosis is the caspase-dependent exposure of PtdSer on the outer leaflet of the plasma membrane (12–14) *via* the XK-related protein 8 (15). Exposure of PtdSer affects the biophysical characteristics and organization of the plasma membrane through recruitment of proteins to PtdSer-enriched regions *via* electrostatic interactions (16). Phagocytes express transmembrane receptors that bind PtdSer directly, e.g., brain-specific angiogenesis inhibitor-1 (BAI-1) (17) and stabilin-2 (18). In addition, soluble molecules such as transthyretin-like protein TTR-52 (19), milk fat globule EGF-factor 8 (MFG-E8) (20, 21), protein S (Pros1), Gas6 (22) and C1q (23) also bind to (and opsonize) PtdSer, providing a scaffold for phagocyte recognition *via* a diverse array of counter-receptors.

Phagocytes fail to engulf viable cells that expose low levels of PtdSer during activation (24–26) or when PtdSer exposure is induced by overexpression of a phospholipid scramblase, transmembrane protein 16F (TMEM16F) (27), suggesting that additional signals are necessary to initiate efferocytosis. A critical threshold of PtdSer exposure on the cell surface may be required to trigger efferocytosis (28). For example, recognition of PtdSer *via* T-cell immunoglobulin and mucin-domain-containing molecule (TIM)-4 was dependent on ligand density, allowing phagocytes to distinguish between high and low level PtdSer exposure (28). Further modifications of PtdSer during apoptosis, e.g., oxidation or formation of lyso-PtdSer (29) may also be important.

#### CELL SURFACE RECEPTOR ALTERATIONS ASSOCIATED WITH APOPTOSIS

Apoptosis-dependent loss of cell surface receptors or appearance of "new" molecules may confer recognition by phagocytes. For example, signaling *via* Signal regulatory protein-α (SIRPα) inhibits myosin-II-mediated phagocytosis (30). Downregulation of ligands for SIRPα, e.g., CD47 (31), from the surface of apoptotic cells would be predicted to promote efferocytosis (32, 33). SIRPα-mediated signaling has also been reported to be triggered by binding of surfactant protein (SP)-A and SP-D to phagocytes. However, SP-A may have a dual role in regulation of phagocytosis as binding to apoptotic cells/debris results in promotion of phagocytosis *via* a calreticulin/CD91-mediated pathway (34). Early experiments identified a unique charge-sensitive mechanism for apoptotic cell recognition (35). The cell glycocalyx provides a negative surface charge "repulsive" force that counters cell–cell interactions (36). Loss of N-terminal sialic acid and exposure of mannose and fucose moieties during apoptosis reduces electrostatic forces that counter phagocyte recognition (37–39). In addition, the surface charge of apoptotic cells is further altered by reduced expression of heavily sialylated proteins (e.g., CD43, CD45, and CD162) (37). Consistent with this suggestion, removal of sialic acid from the cell surface by neuraminidase treatment enhances phagocytosis (37, 39).

Apoptosis is associated with loss of expression of complement regulatory proteins such as CD46/CD55 (40, 41). As a result, deposition of complement may occur, providing a cue for recognition by phagocytes. Additional signals for phagocytosis may occur as a result of exposure of intracellular proteins such as calreticulin (42) and annexin I (43). Following binding to PtdSer on apoptotic cells, oxidation of Pros1 induces oligomerization, which promotes Mer-dependent phagocytosis (44). Similarly, altered glycosylation of membrane proteins or oxidization of lowdensity lipoprotein-like moieties on apoptotic cells (8) may also contribute to the specific recognition by phagocytes. In addition to increased expression of ligands for phagocytic receptors on apoptotic cells, patching and/or clustering of surface molecules may also have important consequences for triggering phagocyte responses. Clustering might occur through specific association with membrane microdomains. For example, FcγRIIa redistributes to membrane microdomains during neutrophil apoptosis (45). In addition, specific proteolysis of adhesion molecules (e.g., CD62L) (46, 47) and uncoupled β2 integrin-mediated adhesion (47) during apoptosis is likely to provide additional molecular cues for phagocytosis.

### PHAGOCYTE MOLECULES THAT MEDIATE APOPTOTIC CELL RECOGNITION

Phagocytes are capable of direct recognition of PtdSer exposed on the apoptotic cell surface. BAI-1 binds PtdSer *via* thrombospondin (TSP) type 1 repeats present in the extracellular domain (48). Binding induces the formation of a trimeric complex of BAI-1 with the Rac-GEF ELMO and DOCK180 that promotes subsequent engulfment of apoptotic cells (17, 49). This pathway is homologous to the genetically defined pathway for removal of apoptotic cells in *Caenorhabditis elegans* (Ced2-CrkII, Ced5- DOCK180, Ced10-Rac, and Ced12-ELMO) (50).

Phosphatidylserine is also recognized by the CD300 family of molecules with an extracellular IgV-like domain and intracellular adaptor molecule binding sites (51). CD300b localizes to phagocytic cups and binds DAP12, activating Syk and PI3K/ Akt to promote phagocytosis (52). Stabilin-2 binds PtdSer and also lacks direct signaling activity (18). However, the cytoplasmic domain of stabilin-2 can interact with GULP to facilitate phagocytosis (53). GULP also binds to NPxY motifs present in the cytoplasmic domains of CD91/LRP (low-density lipoprotein receptor-related protein) and the *C. elegans* scavenger receptor Ced-1 (54). In contrast, TIM-4 confers Ca2+-dependent PtdSerdependent apoptotic cell recognition, but lacks intracellular signaling potential (55). Thus, TIM-4 functions cooperatively with other receptors that trigger apoptotic cell internalization.

Indirect recognition of apoptotic cells by phagocytes is also achieved by phagocyte receptors that bind to soluble apoptotic cell opsonins. In *C. elegans*, TTR-52 bridges apoptotic cellexposed PtdSer to phagocyte Ced-1 (19), which together with Ced-6 initiates rapid and efficient engulfment of apoptotic cell corpses by neighboring cells. This module of proteins (Ced1-MEGF-10; Ced6-GULP; and Ced7-ABCA1) has been defined genetically in *C. elegans* (50). Pros1 and Gas6 contain a Gla-domain that binds PtdSer in a Ca2+-dependent manner (22), bridging to the Tyro3/Axl/Mer receptor tyrosine kinases that signal particle internalization *via* intrinsic kinase activity (56). By contrast, MFG-E8 binds PtdSer in a Ca2+-independent manner and bridges to phagocyte integrins αvβ3/5 *via* arginine– glycine–aspartic acid (RGD) peptide motifs in the C1 and C2 domains (21). TSP-1 also bridges apoptotic cells *via* the phagocyte integrin αvβ3 and CD36 (57, 58).

#### COOPERATIVE RECEPTOR UTILIZATION IN PHAGOCYTOSIS OF APOPTOTIC CELLS

Phagocytes in different tissue settings or microenvironments express distinct repertoires of efferocytic receptors. Whether a single phagocytic cell utilizes multiple receptor pathways to recognize and internalize a single apoptotic target is not clear. However, the distinct molecular requirements for the capture and subsequent internalization may require that multiple receptors are involved (42). In addition, the complex topology of apoptotic cell surface molecules and co-opsonization of PtdSer with different proteins may determine the spectrum of signal transduction pathways engaged, controlling internalization and subsequent phagocyte responses in a context-dependent manner.

Tethering of IgG-opsonized particles to FcγR occurs at 4°C (59) whereas internalization requires cytoskeletal reorganization and metabolic activity (60). Similarly, apoptotic cells can also be tethered by phagocytes *via* Mer at low temperature (61). However, the avidity of low affinity receptors is influenced by receptor density and rapid lateral movement of receptors to facilitate target capture (62, 63). Receptor mobility is controlled by cytoskeletal constraint, association with membrane lipid microdomains and/or other membrane proteins (64). For example, cytoskeletal-associated CD44 restricts membrane lipid and receptor motility *via* interactions with hyaluronan, forming a glycosaminoglycan barrier that reduces binding of phagocytic targets (36). Interestingly, CD44 cross-linking with antibody augments macrophage phagocytosis of apoptotic cells, possibly as a result of changes in cytoskeletal regulation (65, 66). Phagocytic targets are bound at dynamic extensions of phagocytic cells, including filopodia and membrane ruffles (67), and receptor aggregation is required for orchestration of cytoskeletal alterations necessary for internalization [see Ref. (68) for a comprehensive review of cytoskeletal regulation in phagocytic synapses].

In contrast with IgG or complement attached to components of the microbial cell wall, molecules on the surface of apoptotic cells may exhibit unrestricted lateral mobility as a consequence of proteolytic cleavage of actin during apoptosis (69). Thus, there may be key mechanistic differences between efferocytosis and FcγR-dependent phagocytosis. Engagement of freely mobile molecules by phagocyte receptors may lead to assembly of receptor microclusters and significantly impact upon phagocytosis (33). Alternatively, opsonization of apoptotic cells may result in formation of immobile molecular complexes [like annexin V (70, 71)] that promote redistribution of phagocytic receptors necessary for signaling of internalization.

#### MOLECULAR SEGREGATION AND THE FORMATION OF A PHAGOCYTIC "SYNAPSE"

The cellular contact between phagocyte and apoptotic cells has parallels with those of antigen-specific T and B cells with antigen presenting cells that leads to establishment of an immunological synapse (72). This specialized intercellular contact zone stabilizes adhesion and facilitates efficient molecular communication following antigen-specific interactions. A number of different biophysical factors (including receptor density, ligand-binding affinity, molecular dimensions, and interactions with cytoskeletal elements) all contribute to the dynamic redistribution of adhesion and signaling receptors into distinct regions in the plasma membrane (73, 74).

Following phagocyte contact with IgG-coated surfaces or on supported lipid bilayers, formation of FcγRII nanoclusters suggests activation-driven organization of receptor redistribution (75). FcγR clusters are localized in front of ruffles on extending pseudopods, with rapid recruitment of Syk to advancing pseudopods and subsequent retrograde movement toward the cell center (76). PI3K co-localized with actin around FcγR clusters, suggestive of signal propagation from FcγR and consistent with PI3K-dependent control of actin cytoskeletal rearrangements (76).

Although phagocyte contact with apoptotic cells has not been examined with the high-resolution imaging techniques used in T cell–APC interactions, molecular segregation may also be a key feature of the formation of efferocytic synapses (see **Table 1**). For example, exclusion of phosphatases (e.g., CD45) from the immune synapse is a critical early event in the initiation of T cell receptor-mediated phosphorylation of Zap70 and Lck (77). Changes in the distribution of CD45 may also represent a general feature of membrane alterations that control signaling events associated with phagocytosis. The C-type lectin containing receptor for β-glucan, Dectin-1, mediates the recognition Table 1 | Comparison of phagocytic synapse formation in FcγR-mediated and apoptotic cell phagocytosis.

*a See Elliott et al. (80) for a comprehensive overview of receptor signaling. ITIM, immunoreceptor tyrosine-based inhibition motif; ITAM, immunoreceptor tyrosinebased activation motif; BAI-1, brain-specific angiogenesis inhibitor-1.*

and phagocytosis of yeast particles. During Dectin-1-mediated phagocytosis, exclusion of CD45 and the receptor type protein tyrosine phosphatase CD148 from the nascent phagocytic cup (78) is important for signaling associated with particle internalization (79). Redistribution of phosphatases in the phagocyte membrane would likely be necessary for internalization of apoptotic cells.

Analysis of the molecular basis of redistribution of CD45 following ligation of FcγR suggests non-linear pattern with formation of an exclusion barrier which restricts access of CD45 to the contact site (79). CD45 is a relatively rigid molecule that extends axially from the plasma membrane approximately 20 nm (81). Similar to the molecular redistribution that occurs during immune synapse formation, exclusion from phagocytic synapses was dependent on the axial molecular dimensions of CD45. In experiments using chimeric constructs in which the extracellular portion of CD45 was replaced by either CD43 (similar length) or CD2 (shorter length), the CD43/CD45 molecule was excluded from the phagocytic synapse, whereas CD2/CD45 was not (79). A requirement for integrins and cytoskeletal regulation was shown to be necessary to establish a CD45 exclusion zone that extended beyond the IgG layer (79). These results suggest that integrin-mediated contact between phagocyte and phagocytic targets facilitates engagement of phagocytic receptors at low ligand densities or when binding to larger particles.

For apoptotic cell recognition, it is intriguing that integrins have been proposed to act cooperatively with other receptors to mediate phagocytosis. For example, αv integrins and CD36 are both required for the recognition of TSP-1 bound to apoptotic cells (58). Similarly, phagocytosis of apoptotic cells *via* TIM-4 requires β1 integrins and activation of integrin-dependent signaling involving Src family kinases and FAK (82). Furthermore, interactions between the β5 integrin and stabilin-2 were found to promote phagocytosis of apoptotic cells (83). Similarly, co-expression of αvβ5 with Mer increased activation of Rac-1, cytoskeletal regulation and the phagocytosis of apoptotic cells (84). Integrins can directly mediate the recognition of apoptotic cell opsonins, for example, RGD-dependent binding of MFG-E8 (21) or fibronectin (85). However, phagocytosis of apoptotic targets is increased following macrophage adhesion to extracellular matrix *via* β1 integrins (86) and is compromised following exposure to oxidized extracellular matrix molecules (87). We would speculate that, as for FcγR-mediated phagocytosis, integrin signaling can regulate cytoskeletal organization and facilitate tethering/phagocytosis of apoptotic cells with low-level opsonization (36).

For human macrophage-like cells, CD47 expression acts to limit phagocytosis of IgG-opsonized erythrocytes (30). CD47 binds to phagocyte SIRPα, resulting in recruitment to the phagocytic synapse, decreasing the accumulation of non-muscle myosin IIa and levels of tyrosine phosphorylation (30). Localization of SIRPα to the site of cell contact would recruit inhibitory tyrosine phosphatases such as SHP-1 *via* ITIM motifs present in the SIRPα cytoplasmic domain (88). In the absence of CD47 or when CD47 was blocked with antibody, phagocytosis was increased. Specific membrane receptors are organized into protein islands in unactivated T cells that subsequently coalesce as a consequence of T cell receptor-mediated signaling (89). The membrane distribution of SIRPα and FcγR during phagocytosis has been further analyzed using super-resolution microscopy. When macrophages were plated onto poly-l-lysine-coated slides, molecular clusters containing both FcγRI and SIRPα were observed (75). These molecules were found to segregate into discrete nanoclusters when macrophages were plated onto IgG. Interestingly, IgG promoted the formation of concentric rings of FcγRI and FcγRII, with FcγRI redistributing more rapidly (<10 min). Similar results were obtained when macrophages interacted with IgG in a supported lipid bilayer. When recombinant CD47 was included into the supported lipid bilayer, segregation of FcγR and SIRPα and the formation of concentric rings of FcγR were blocked. Thus co-localization of SIRPα and FcγR inhibits cellular activation following FcγR ligation, whereas segregation of these two molecules leads to activation (75). We would speculate that efferocytic receptors would also be present in dynamically regulated nanoclusters in the phagocyte membrane.

#### MOLECULAR DIMENSIONS AND APOPTOTIC CELL RECOGNITION

Size-dependent redistribution of molecules within the phagocyte membrane may represent an important organizing principle for the assembly of molecular platforms that are essential for signaling the cytoskeletal alterations required for the internalization of apoptotic cells. Estimation of the molecular dimensions of receptors involved in efferocytosis using published structural data (51, 90–106) reveals considerable differences in axial dimensions (**Figure 2A**).

First, it is notable that CD300 (single Ig-like domain) and TIM-4 (single Ig-like domain with a potentially rigid mucin-like stalk), which mediate direct recognition of PtdSer, are predicted to span a relatively short intermembrane distance between phagocyte and target (4 and 7 nm, respectively, **Figure 2B**). By contrast, stabilin-2, which also binds to PtdSer, is a much longer molecule extending some 23 nm. Assuming a degree of structural flexibility, BAI-1 may be capable of functioning in a broad range of intermembrane distances. The extracellular region of BAI-1 contains 5 TSP type 1 repeats (around 5 nm in size) with LPS and PtdSer binding motifs, which together with the GAIN/HBD regions could extend to ~33 nm from the plasma membrane. Following initial tethering of PtdSer by the N-terminal TSP repeat, BAI-1 could align parallel to the apoptotic cell surface as additional TSP repeats become ligated (**Figure 2B**). Nevertheless, it seems likely that a degree of molecular segregation would be required for these different receptors to be involved in apoptotic cell uptake on the same phagocyte.

Second, receptors such as SR-A or LRP are predicted to be highly extended molecules (~40–50 nm). In addition, some of the well characterized opsonins for apoptotic cells are extremely large. For example, C1q is approximately 30 nm and TSP-1 is 40–50 nm. Together with the relatively large counter receptors for these opsonins [e.g., SCARF1 for C1q (107) and αvβ3 for TSP], the predicted intermembrane contact distance would likely be incompatible with those of BAI-1, αvβ5/MFG-E8, or Mer/Protein S. One possibility is that these extended structures are efficient in the initial capture of apoptotic targets, facilitating subsequent engagement of receptor/counter-receptor pairs that span a smaller intermembrane distance and are influenced by electrostatic repulsion between cells.

Third, receptors of the Tyro3/Axl/Mer family, BAI-1, and the integrin αvβ5 are able to initiate intracellular signaling that controls particle internalization. If there are parallels with immunological synapse formation, these receptors might be expected to become localized at the center of a contact zone. However, the intermembrane distance for TAMs and integrins to engage their counter-receptors (~30 nm) is considerably larger than that of the immunoreceptors that are responsible for signaling during the establishment of the immunological synapse (~14 nm for the TcR/MHC class II and co-stimulatory receptors such as CD80/ CD28). We would speculate that the organizing principles for immune and efferocytic synapses would be different due to the distinct requirements for signal propagation following cognate interaction of receptors. In an immune synapse, delivery of a signal that controls T cell proliferation or target cell killing requires maintenance of intercellular adhesion and redistribution of antigen-specific recognition molecules to the contact zone. By contrast, an efferocytic synapse would likely be a more dynamic structure that facilitates particle tethering and allows dynamic regulation of cytoskeletal organization as particle internalization proceeds. As discussed earlier, the localization

phosphatidylserine (PtdSer) binding (either directly or indirectly). (B) Example receptor–counter-receptor pairings are shown to illustrate the likely differences in intermembrane "working" distance during phagocytosis, particularly for the molecules involved in the recognition of PtdSer (shown in yellow). Phagocyte receptors are shown in blue, opsonins are shown in green, and the apoptotic cell membrane is shown in red.

of receptors involved in phagocytosis may involve initial tethering mediated by larger molecules as a prerequisite for engagement of smaller PtdSer binding receptors. The mechanisms for exclusion of phosphatases from an efferocytic synapse might also be distinct.

Finally, although phagocytosis of intact apoptotic targets is readily observed *in vitro*, the tight apposition of the plasma membranes of phagocytes and apoptotic targets in certain tissues *in vivo* may require a mechanistically distinct clearance process. For example, in the retina clearance of the outer segments of photoreceptors by retinal pigment epithelial cells has been likened to a phagocytic "pruning" of the photoreceptor outer segments (108). The molecular basis of removal of photoreceptor outer segments by retinal pigment epithelial cells involved Mer, Pros1 and Gas6, and the integrin αvβ5 (108–110). Electron microscopy analysis of retinal cell architecture reveals the exquisitely close contact between RPE cells and photoreceptors (111). Elegant *in vivo* imaging studies have revealed the diurnal exposure of PtdSer in localized a manner, which then triggers the "pinching off " of the distal tips of the photoreceptors by the adjacent RPE cells (112). The exposure of PtdSer on viable photoreceptors may have some parallels with a process termed phagoptosis (113) in which viable cells are recognized by phagocytes. However, during phagoptosis, recognition triggers apoptosis in the target cell (114). Although similar molecular pathways are involved in the recognition of PtdSer exposure on viable cells (e.g., MFG-E8, stabilins, αv integrins, and Mer) (115), the intercellular communication events that are involved are likely to be distinct from those required for efferocytosis.

In summary, the establishment of an efferocytic synapse may be required for efficient recognition of apoptotic cells by phagocytes. Cooperativity of receptor engagement may act to facilitate and stabilize adhesive interactions and lead to the assembly of signaling platforms that ultimately determine phagocyte responses to apoptotic cell binding and internalization.

#### AUTHOR CONTRIBUTIONS