Loss of cell surface CD47 ‘clustering’ formation and binding avidity to SIRPα facilitate apoptotic cell clearance by macrophage

Abstract

CD47, a ‘self’ recognition marker expressed on tissue cells, interacts with immunoreceptor SIRPα expressed on the surface of macrophages to initiate inhibitory signaling that prevents macrophage phagocytosis of healthy host cells. Previous studies have suggested that cells may lose the surface CD47 during aging or apoptosis to enable phagocytic clearance. In the present study, we demonstrate that the level of cell surface CD47 is not decreased but the distribution pattern of CD47 is altered during apoptosis. On non-apoptotic cells, CD47 molecules are clustered in lipid rafts forming ‘punctates’ on the surface, whereas on apoptotic cells, CD47 molecules are diffused on the cell surface following the disassembly of lipid rafts. We show that clustering of CD47 in lipid rafts provides a high binding avidity for cell surface CD47 to ligate macrophage SIRPα, which also presents as clusters, and elicit SIRPα-mediated inhibitory signaling that prevents phagocytosis. In contrast, dispersed CD47 on the apoptotic cell surface is associated a significant reduction of the binding avidity to SIRPα and failure to trigger SIRPα signal transduction. Disruption of lipid rafts with methyl-β-cyclodextrin (MβCD) disrupted CD47 cluster formation on the cell surfaces, leading to decrease of the binding avidity to SIRPα and a concomitant increase of cells being engulfed by macrophages. Taken together, our study reveals that CD47 normally is clustered in lipid rafts on non-apoptotic cells but is diffused in the plasma membrane when apoptosis occurs, and this transformation of CD47 greatly reduces the strength of CD47-SIRPα engagement, resulting in the phagocytosis of apoptotic cells.

Introduction

CD47 is an immunoglobulin (Ig) superfamily transmembrane protein that is universally expressed on mammalian cells and tissues. Through its trans interactions with SIRPα on macrophages, CD47 triggers tyrosine phosphorylations in the SIRPα cytoplasmic immunoreceptor tyrosine-based inhibitory motifs (ITIMs) and recruitment of protein tyrosine phosphatases SHP-1/SHP-2, which further mediate negative signaling events that inhibit macrophage phagocytosis. For this, CD47 acts as a “self” marker and prevents macrophage engulfment of host cells (1, 2). This self-recognition system mediated by CD47-SIRPα interaction plays a critical role in restraining macrophages. Disruption of CD47-SIRPα interaction would lead to normal tissue damage (3–6) on one hand, while preservation of this self-recognition could result in failure of clearing apoptotic cells, pathogen-infected cells, or tumor cells (7) on other hand.

Recent studies of cell apoptosis and how apoptotic cells are cleared by macrophages suggest that there are three kinds of potential signals controlling macrophages to target apoptosis cells. The first signal is a ‘find me’ signal. The apoptotic cells release soluble factors such as lysophosphatidylcholine (LPC) (8) that act as chemoattractants for recruiting macrophages or other phagocytes. Following macrophages approaching, previous studies have shown molecules that are especially increased on apoptotic cells, such as phosphatidylserine (PS) (9) and calreticulin (10, 11), initiate the next ‘eat me’ signaling, the second class of signal (7,8). Meanwhile, CD47, through ligation of macrophage SIRPα, provides an additional control - the “don’t eat me” signal, which should restrain the process initiated by the first two classes of signaling. As apoptotic cells do indeed get engulfed by host macrophages, some explanations regarding the impotence of this usually effective final veto is required. Evidence suggests that apoptotic cells, as well as senescent cells, may lose their surface CD47 or change the cell surface localization pattern of CD47 (12–14), resulting in a dysfunction of “don’t eat me” signaling. However, the mechanism that governs the changes of both cell surface expression level and the pattern of CD47, and how the CD47 pattern change affects the CD47-SIRPα interaction during apoptosis is incompletely understood.

In the present study, we monitored the kinetics of the cell surface level and the pattern of CD47, and also the CD47-SIRPα interaction following UV-induced cell apoptosis or apoptosis induced by other means. Our results showed that cell apoptosis does not decrease the CD47 level on the cell surface but alters the cell surface pattern of CD47 from ‘punctate’ clusters into diffused distribution, which dramatically decreases the avidity of CD47-mediated cell binding to SIRPα and incapacitates SIRPα-mediated inhibitory signaling in macrophages. Our data further suggest that dispersion of surface CD47 is related to apoptosis-induced disruption of lipid rafts in the plasma membrane.

Material and Methods

Cells, antibodies and reagents

Human colonic epithelial cell HT-29, human mammary gland epithelial cells T47D, MCF7, MDA435 and HS578T, and primary cultured human foreskin fibroblasts (HFF-1) (all from American Type Culture Collection (ATCC)) were maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS). Human microvascular endothelial cells (HMEC-1) initially primarily cultured by Dr. E.W. Ades (Centers for Disease Control and Prevention, Atlanta) (15) were maintained in MCDB 131 medium with 10 mM/L L-glutamine, 10 ng/ml mouse epidermal growth factor (mEGF, BD Biosciences), 1 µg/ml hydrocortisone (Sigma) and 10% FBS and were used within 15 passages (16). Human acute monocytic leukemia cells THP-1 (from ATCC) were cultured in RPMI 1640 supplemented with 50µM β-mercaptoethanol and 10% FBS (16). Human peripheral polymorphonuclear leukocytes (PMN) were isolated from whole blood of healthy volunteers (16–18). Murine splenocytes were isolated from spleens of healthy C57BL6 mice. Mouse mAbs against the human CD47 extracellular domain, B6H12.2 and PF3.1, and the human SIRPα extracellular domain, SE5A5 and SE7C2, and a rabbit polyclonal antibody against the SIRPα extracellular domain (anti-SIRPα.ex) were used previously (19, 20). The IgG Fab fragment of PF3.1 was prepared by cleavage with papain protease using a Fab Preparation Kit (Thermo Scientific, Rockford, IL). Mouse mAb clone 4G10 (EMD Millipore) was used to detect protein tyrosine phosphorylation and anti-SHP-1 antibody was purchased from Santa Cruz Biotechnology. Rat mAb against mouse CD47, miap301, was purchased from BD Biosciences. The anti-actin antibody and FITC-conjugated cholera toxin B subunit (CT-B) were obtained from Sigma. A recombinant fusion protein containing the entire extracellular domain of human SIRPα and rabbit Fc, SIRPα.ex-Fc, was generated and used previously (21, 22). Another recombinant fusion protein containing the entire extracellular domain of human CD47 and alkaline phosphatase (AP), CD47-AP, which functionally ligates SIRPα, was generated in the lab (17, 22, 23). Octylglucoside (OG) was from Millipore. Other essential reagents including Methyl-β-cyclodextrin (MβCD), lipopolysaccharide (LPS), lectin from Psophocarpus tetragonolobus, PMA, protease inhibitors and cocktails were purchased from Sigma.

Cell apoptosis

HT-29, T47D, HS578T, MCF7, HFF-1 and HMEC-1 were induced apoptosis by UV irradiation in 35 mm Petri dishes in the absence of cell culture medium. Specifically, after removal of culture medium, the dishes were placed under a UV lamp/cross-linker (model XL-1000, Spectronics Corporation, New York) and irradiated for various time periods, all within 200 seconds, to achieve induction of cell apoptosis starting at 2–4 h post-UV, but not necrosis or instant death (the total energy strength for different cell types are listed in Table 1). After UV irradiation, the culture medium was added back to the dishes and the cells were cultured for various time periods until analysis. Cell apoptosis was assessed by labeling with FITC-conjugated Annexin V (Sigma) for cell surface PS and YO-PRO-1 iodide (Life Technologies) for cell nuclei. Cell necrosis or dead cells were detected by staining with propidium iodide (PI). Cell apoptosis was also assessed by Western blot to detect cleavage of Poly ADP-Ribose Polymerase (PARP) using a rabbit polyclonal anti-PARP antibody (Roche).

Table I. Cell apoptosis and necrosis after UV irradiation.

Healthy cultured cells (≥95 viability) of various types were UV irradiated to induce apoptosis. At different time point post-UV irradiation, cells were stained with YO-PRO-1 (green) and PI (red) to determine apoptosis and necrosis, respectively. The data (means ± SD) represent one of three independent experiments in which % of apoptotic cells (only green) and necrotic cells (green and red double staining (orange)) were microscopically determined in ten visual fields against the total cells.

Cell type	UV dose (mJ/cm2)	Post-UV (h)	Apoptosis (%) YO-PRO-1 (green)	Necrosis (%) PI (red)
HT29	150	0	3.2 ± 1.0	2.3 ± 0.9
		2	11.8 ± 4.1	2.0 ± 1.0
		4	60.7 ± 7.3***	5.9 ± 2.0
		6	87.6 ± 6.2***	16.2 ± 3.0**
		8	100***	28.1 ± 5.8***
		18	100***	100***
T47D	150	0	2.4 ± 0.9	1.9 ± 0.5
		6	89.6 ± 6.2***	18.2 ± 6.6**
MCF7	125	0	1.9 ± 1.1	2.7 ± 0.8
		6	91.3 ± 8.4***	20.3 ± 7.5**
MDA231	150	0	3.0 ± 0.8	2.6 ± 0.8
		6	92.8 ± 5.9***	17.9 ± 3.2**
MDA435	180	0	2.9 ± 1.2	2.5 ± 0.7
		6	93.4 ± 9.1***	21.3 ± 6.0**
HS568T	150	0	2.2 ± 0.6	3.4 ± 0.9
		6	87.3 ± 5.8***	17.1 ± 4.6**
MDA468	100	0	3.0 ± 0.9	2.8 ± 0.5
		6	88.6 ± 9.8***	15.3 ± 7.8**
HFF-1	180	0	2.1 ± 0.8	1.7 ± 0.6
		6	71.4 ± 5.8***	14.6 ± 4.8**
HMEC-1	150	0	3.8 ± 0.6	2.1 ± 0.6
		6	79.3 ± 6.7***	20.6 ± 5.3**

^**p≤0.01

^***p≤0.001

Cell adhesion assay

Cell adhesion to immobilized recombinant proteins was performed according to previously described methods (22, 24) with modifications. Briefly, the purified SIRPα.ex-Fc (10 µg/ml), lectin (50 µg/ml), control BSA (5%), or anti-CD47 mAb PF3.1 (10 µg/ml) in HBSS were immobilized in 96-well microtiter plates by incubation for 2 h at 25°C or overnight at 4°C. After blocking with HBSS containing 2% BSA and 5% normal goat serum (1 h, 25°C), the wells were incubated with suspensions of cells (1–5 × 10⁵) for 30 min (25°C) in the presence or absence of inhibitory antibodies or peptides. After incubation, the wells were gently washed (3x) using a 27-gauge needle connected to vacuum followed by analysis for cell adhesion microscopically. In some experiments, cells were labeled with a fluorescent dye 2”,7”-bis (carboxyethyl)-5,6- carboxyfluorescein (BCECF) prior to incubation with immobilized proteins. Cell adhesion was quantified using a fluorescence plate reader (Perkin Elmer) by reading the fluorescence intensity of the total load versus the after wash.

Surface plasmon resonance (SPR) measurement

The binding avidity of cells to immobilized SIRPα was analyzed using the SPR technology. Briefly, a new CM5 chip (Biacore, GE Bioscience) with four binding surfaces/channels was placed into the Biacore T100 system (Biacore, GE Healthcare) followed by binding surface activation using 400 mM 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide (EDC) and 100 mM N-hydroxysuccinimide (NHS) according to a standard protocol in Biacore Sensor Surface Handbook (GE Healthcare). After activation, one channel that was used as the reference and three were for binding experiments. To immobilize SIRPα.ex-Fc on the chip, purified SIRPα.ex-Fc (100 µg/ml) in NaAc/HAc buffer (pH 5.0) was injected into activated channels. The final response unit (RU) change with immobilized protein was about 4500. After protein coating, 1 M ethanolamine was then injected into the channels to neutralize residual activated carboxyl groups. The reference channel was directly blocked with ethanolamine. For cell adhesion experiments, cells suspended in HBSS (1 × 10⁷ cells/ml) were injected into the channels at a flow rate of 3µl/min, which was lasted at least 500 sec. Continual flow of HBSS at the same rare was given after the cell injection. To determinate cell binding, the channels were washed/regenerated by running 5 M LiCl in HEPES, pH 7.3, for 15 min at a flow rate of 3µl/min.

In vitro binding assay of purified CD47 to SIRPα

CD47 purification from various cells was described previously (17). Briefly, non-apoptotic cells and cells that were 6 h post-UV (> 5 × 10⁸ each) were lysed in a cold buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 2 mM MgCl₂, 1% NP-40, 1% TritonX-100, 0.5% sodium deoxycholate, protease inhibitor cocktail (Sigma) and 1 mM PMSF). After pre-clearing by a control IgG-conjugated Sepharose, the lysates were incubated with Sepharose conjugated with PF3.1 (2 mg of IgG/ml). After incubation, the beads were washed with the same lysis buffer but with the detergent changed to 1% octylglucoside. After washing, CD47 was eluted in a buffer containing 50 mM TEA, pH 10.5, 1% octylglucoside and 100 mM NaCl, followed by neutralization. To assay CD47 binding to SIRPα.ex-Fc, purified CD47 (~100 µg/ml in buffer containing 1% OG) was diluted 10-fold in HBSS in ELISA plate wells and allowed protein binding for 2 h (25°C). The wells were then blocked, and incubated with SIRPα.ex-Fc (5 µg/ml) in HBSS for 30 min in the presence or the absence of anti-CD47 or anti-SIRPα mAb (5µg/ml). After washing, SIRPα.ex binding was detected using a peroxidase-conjugated goat anti-rabbit Fc antibody and the substrate 2,2’-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS).

Immunofluorescence labeling, FACS and microscopy

Trypsinized cells were blocked with 5% normal goat serum followed by cell surface labeling of CD47 with antibody B6H12.2 or PF3.1 (30 min). Cells labeled with mouse normal IgG were used and controls. To label cells growing in the culture dishes, cells were briefly treated with 1 mM EDTA to dissemble the intercellular junctions (21) before blocking and CD47 labeling. After primary antibody labeling, cells were washed and labeled with AlexaFluor–conjugated anti-mouse secondary antibody (Invitrogen) followed by FACS, fluorescence microscopy or confocal microscopy in the presence of anti-fade reagent (Molecular Probes). CD47 granularities and particle fluorescence intensities were analyzed by NIH software Image J and Nikon camera software NIS-Elements B.R. 4.20.00. In some experiments, CD47-expressing cells were also incubated with SIRPα.ex-Fc (5 µg/ml) in HBSS (30 min, 4°C). After washing, SIRPα.ex-Fc binding was detected with an AlexaFluor–conjugated anti-rabbit Fc antibody. Plasma membrane lipid rafts were labeled with FITC–conjugated cholera toxin B (30 min, 4°C).

Immunoprecipitation (IP) and Western blot (WB) analysis

Cells were lysed by an ice-cold lysis buffer containing 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% NP-40, 1% TritonX-100, 0.5% sodium deoxycholate, a cocktail of protease inhibitors (Sigma) and 1 mM PMSF. Clear cell lysates were obtained by centrifugation at 14,000 rpm (10 min). For WB of CD47, cell lysates were supplemented with NaIO4 (10 mM, to enhance CD47 detection) prior to mixing with non-reducing SDS-PAGE sample buffer and heating (5 min, 60°C). After transferring proteins onto nitrocellulose, the membrane was blocked with 5% nonfat milk followed by detection using anti-CD47 mAb B6H12.2 (2 µg/ml, 1 h, 25°C). For WB of actin and PARP, SDS-PAGE was performed under reducing conditions. For WB of SIRPα, anti-SIRPα.ex was used. For IP of SIRPα, the THP-1 macrophage lysates were incubated with 1–2 µg anti-SIRPα.ex and protein A-conjugated Sepharose for 4 h (4°C). After washing, the beads were heated in 1x SDS-PAGE sample buffer to release proteins, and samples of pre-bound lysates, post-bound lysates and IP were separated by SDS-PAGE followed by WB analyses for SIRPα. To detect SIRPα phosphorylation, THP-1 macrophages were treated with freshly prepared pervanadate (2 mM, 3 min, 37°C) prior to cell lysis and IP. The tyrosine phosphorylated SIRPα was WB detected using mAb 4G10, and SIRPα associated SHP-1 was detected using an anti-SHP-1 antibody.

Phagocytosis assay and cytokine detection

Cultured THP-1 cells were induced to differentiate into phagocytic macrophages using PMA (100nM) for 3 days (37°C). After induction, THP-1 changed morphology and adhered to the culture dishes. To assay THP-1 macrophage phagocytosis, healthy cultured cells (non-phagocytosis control) and cells that were induced apoptosis by UV or other methods were loaded with carboxyfluorescein diacetate succinimidyl ester (CFSE) or labeled with YO-PRO-1 iodide (for apoptotic cells) prior to incubation with THP-1 phagocytes at 37°C. After 30 min, the dishes were gently washed to remove floating cells, and THP-1 phagocytes were stained with a PE-conjugated anti-CD11b antibody followed by fluorescence microscopy. To determine phagocytosis, macrophages (CD11b⁺, red) in 5–6 view fields with a total cell number > 200 were analyzed for ingestion of target cells that were stained with CSFE (green) or YO-PRO-1 (apoptotic marker, green). Successful phagocytosis were those macrophages in which green cells were ingested or partially ingested hence showing green staining, while macrophages without green staining within the cells were not considered phagocytic. Alternatively, a fluorescence plate reader (Perkin Elmer, Waltham, MA) was used to assess phagocytosis and in those experiments only target cells were labeled green. After co-incubation with THP-1 macrophages (without labeling), the wells were washed to remove cells that were not phagocytized or ‘stuck’ to phagocytes, followed by reading fluorescence intensities that reflected cells captured by macrophages. This method is normally used in combination with microscopic determination. To detect cytokines produced during phagocytosis, THP-1 macrophages (2 × 10⁵) were incubated with healthy cells or cells that were treated by UV or other methods (1 × 10⁵ each cell sample). At various time points, the cell-free media from the THP-1-target cell co-incubation were collected and used for ELISA detection of IL-10, IL-6 and TNFα using capture and detection antibodies (27, 28).

Lipid raft staining and disruption

HT-29 cells were stained with FITC-conjugated cholera toxin B subunit to identify lipid rafts (29). To disrupt the lipid rafts on cell surfaces, cells were incubated with 15 mM MβCD for 20 min at 37°C as described previously (30).

Statistical analysis

All images of immunofluorescence labeling, FACS and Western blots represent the results of at least three independent experiments. Data are presented as the mean ± SD or SEM for three or more independent experiments. Differences were considered statistically significant at P<0.05, as analyzed using Student’s t-tests for paired samples and one-way ANOVA for k >2 samples.

Results

Cell apoptosis does not reduce the cell surface level of CD47 expression

It is generally believed that the interaction between the phagocyte surface SIRPα and the self-recognition marker protein, CD47, controls the phagocytosis of self-cells by phagocytes such as macrophages, and that during cell aging or apoptosis, cells may lose surface CD47 and/or the interaction with SIRPα in order to be phagocytized (5, 6). To test whether cells lose their surface CD47 when apoptosis occurs, we examined the CD47 expression level on multiple types of cells prior to and after induction of apoptosis. For the experiments, human colonic epithelial cell HT-29, breast cancer cells T47D and MCF7, and other types of cells were exposed to UV (energy levels of UV used for different cells were listed in Table 1) or other ways to induce apoptosis. As shown in Figure 1A, the UV-treated HT-29, MCF7 and T47D cells started to display apoptosis at about 4 h post-UV, indicated by the cell surface labeling of PS by a FITC-conjugated Annexin V, which started to show positive staining at 4 h and grew stronger thereafter. Cell apoptosis was also confirmed by detecting the cleavage of poly ADP-ribose polymerase (PARP), a DNA repairing polymerase that is cleaved under apoptosis (31) (Figure 1B). As cell necrosis (secondary necrosis) generally occurs following apoptosis if the apoptotic cells were not immediately phagocytically removed (32–34), co-staining with YO-PRO-1 iodide (green) and PI (red) was applied to differentiate cells that were apoptotic (YO-PRO-1 positive but PI negative) or necrotic (PI/YO-PRO-1 double positive). As shown in Table 1, cells began apoptosis 2 h after UV irradiation and, at 8 h, all cells were positive of YO-PRO-1 staining. Cell necrosis started to appear 6 h post-UV, and gradually increased with the time extension. At 18 h post-UV, all cells became necrotic and positively stained by PI. Therefore, in our analyses of apoptotic cells, cells within 8 h post-UV were generally used and YO-PRO-1/PI staining distinguishing apoptosis from necrosis was included in experiments.

Figure 1.

Cell apoptosis is not associated with CD47 reduction. A) HT-29, T47D and MCF7 were induced apoptosis by UV irradiation (~150 mJ/cm²) followed by analyses at 2, 4, 6, 8 or 18 h post-UV for the cell surface CD47 and cell apoptosis by labeling with mAb B6H12.6 and a FITC-conjugated Annexin V for PS, respectively. B) WB detection of the total cellular CD47 using mAb B6H12.6 and cleavage of PARP, an indicator of apoptosis, using a rabbit anti-PARP antibody in UV-irradiated cells collected at different time points post-UV. WB of actin was used as the loading control. C) HT-29 cells collected at various time points after UV irradiation were assessed for the cell surface CD47 by labeling with mAb B6H12.6 (w/o cell permeabilization) and FACS. Cells labeled with mouse normal IgG were used as negative control. D) Detailed flow cytometry analyses of HT-29 cells. Cells collected at various time points post-UV were gated according their sizes (FSC) into two populations: FSC^high (R1) and FSC^low (R2). With apoptosis progression to necrosis, cells in R1 were reducing and, by 18 h post-UV, all cells were shifted to FSC^low R2. Cells in R2 were positive of PI staining thus were dead/ necrotic (Table 1). For cells in R1, increasing in YO-PRO-1staining displayed 4h after UV irradiation, while the cell surface CD47 levels were not significantly changed. The surface CD47 level for cells in R1 was consistently higher than those in R2. E) Plasma membrane blebs and CD47 distribution. Note that the blebs (marked with white arrowheads) emerged at the stage of apoptosis (4–8 h post-UV) were generally depleted of CD47, while the blebs formed during necrosis (18 h post-UV) were CD47-positive.

As shown in Figure 1 (A-C), despite that cells had undergone apoptosis, cell surface labeling of CD47 with mAbs against the CD47 extracellular domain (B6H12.2 and PF3.1) showed that CD47 was continuously expressed on the cell surface without significant reduction until later time points when cells were necrotic (18 h post-UV, Figure 1C). Detailed flow cytometric analyses of cells after UV irradiation (Figure 1D) showed that the cells can be separated into two populations, the FSC^high large cells (gated as R1), and the FSC^low small cells (gated as R2). The non-apoptotic cells (0–2 h) were all FSC^high large cells in R1 region, whereas the PI-positive necrotic cells (18 h post-UV) were FSC^low small cell particles. As shown, at 4–8 h post-UV, although cells were apoptotic (positive YO-PRO-1 staining), they generally maintained the cell sizes (FSC^high, R1) and the surface level CD47. In contrast, when cells became necrotic, they significantly shifted towards the low values of FSC, becoming FSC^low cells with smaller size (R2 region). At 18 h post-UV, nearly all cells were FSC^low and also had significant reduced CD47 on the surface (~40% reduction).

Furthermore, we observed that membrane blebs emerged around the cell bodies when cells started apoptosis and then necrosis. This phenomenon, which has been reported previously, likely reflected vigorous dynamics and/or destabilization of the plasma membrane during apoptosis, resulting in partial plasma membrane dissociation from the underlying cytoskeleton cortex (35–38). For cell apoptosis especially at early stages (4–6 h post-UV), the membrane blebs were small relative to the cell body, transient and deficit of CD47 (Figure 1E, marked by white arrowheads). However, when apoptosis progressed to necrosis, large membrane blebs formed (relative to the remnant cell body) and on which CD47 staining was positive (Figure 1E, marked by white arrowheads). Shedding of these large membrane blebs might serve as a reason for loss of both the CD47 molecules and the cell size for necrotic cells.

Similar results were obtained when studying apoptosis of other cell types, such as human fibroblasts HFF-1 and human microvascular endothelial cells HMEC-1 (both non-transformed cells), the breast cancer cells MDA435, MDA468, MDA231 and HS568A, murine cell lines B16 and MC38, murine freshly isolated splenocytes (> 90% being lymphocytes), and freshly isolated human neutrophils (PMN). As shown in Figure 2, A and B, and Table 1, UV irradiation induced apoptosis of HFF-1, HMEC-1 and various breast cancer cells, indicated by the positive YO-PRO-1 labeling and the cleavage of PARP, while without noticeably affecting CD47 expression on the cell surface as determined by anti-CD47 labeling, as well as immunoblot analysis. Apoptosis in murine splenocytes was induced by treatment with H₂O₂ (5 mM) and this apoptosis was not associated with cell surface CD47 reduction as assessed by a murine CD47 extracellular domain-specific antibody (mAb miap301) (Figure 2C). Similar results of no CD47 reduction were also observed in B16 and MC38 cells when these cells were induced apoptosis by UV irradiation (not shown). Different from other cells, freshly isolated human PMN without stimulation reportedly express only low level CD47 on the cell surface but store majority CD47 in intracellular specific granules (18); chemoattractant stimulation induces PMN degranulation and hence mobilizes CD47 to the cell surface (18). In this study, we treated PMN with the chemoattractant fMLF for 15 min to increase cell surface CD47 (Figure 2D), prior to inducing apoptosis by incubating the PMN on ice in the presence of 20 mM EDTA (EDTA also prevents PMN aggregation). As shown in Figure 2D, PMN that were incubated on ice overnight (> 14 h) were apoptotic and displayed positive of YO-PRO-1 staining; however, such change of PMN also did not result in reduction of CD47 on the cell surface. In conclusion, our results indicate that cell apoptosis, per se, does not lead to loss of CD47 on the cell surface. However, when apoptosis proceeds to secondary necrosis the cell sizes are reduced and as such CD47 proteins remained on the remnant cell surfaces are coordinately decreased.

Figure 2.

No reduction of CD47 on various types of cells during apoptosis. A-B) Multiple types of cells were induced apoptosis by UV irradiation (refer to Table 1) and their cell surface (A) and total (B) CD47 levels were assessed by FACS and WB and compared to cells that were non-apoptotic. Apoptosis was confirmed by positive YO-PRO-1 labeling (A) and cleavage of PARP (B). Panel A also shows the mean fluorescence intensity (MFI) values of CD47 labeling on non-apoptotic cells (red) and apoptotic cells (blue). Cells labeled with mouse normal IgG were used as negative control. C) No apparent alteration of the CD47 level on mouse splenocytes following H₂O₂-induced cell apoptosis. Freshly isolated mouse splenocytes were treated with H₂O₂ (5 mM, 37°C) to induce apoptosis (positive YO-PRO-1 staining). The cell surface CD47 was assessed by labeling with a rat anti-mouse CD47 (mAb miap301) followed by FACS. D) No reduction of the cell surface CD47 on fMLF-stimulated PMN following cell apoptosis. PMN were freshly isolated from peripheral blood of human donors and were then stimulated with fMLF (1 µM) for 15 min (37°C) to induce degranulation and increase cell surface CD47. Apoptosis, monitored by YO-PRO-1 labeling, was induced by incubating the fMLF-stimulated PMN on ice overnight (> 14 h) in the presence of 20 mM EDTA.

Macrophage phagocytosis of apoptotic cells regardless of CD47 expression

Given that CD47 serves as a ‘self’-marker and its interactions with SIRPα on phagocytes inhibit macrophage phagocytosis, we asked whether apoptotic cells with CD47 expression could be phagocytized by macrophages. For these experiments, HT-29 cells were loaded with a fluorescent tracer CFSE prior to the induction of apoptosis by UV irradiation. At various time points post-UV, HT29 cells were incubated with THP-1 differentiated macrophages to allow phagocytosis. As shown in Figure 3, A and B, THP-1 macrophages (labeled with anti-CD11b antibody, red) displayed aggressive phagocytosis towards HT-29 cells 4 h post-UV despite the fact ample CD47 remained on the target cells. Apoptotic cells (post-UV 4–8 h) were rapidly grabbed and engulfed by macrophages and thus formed a wide area of orange labeling in merged images. THP-1 macrophages also aggressively ingested necrotic cells but completely avoided non-apoptotic cells. We also assayed cytokines produced by THP-1 macrophages during phagocytosis. As shown in Figure 3C, cell-free supernatants collected from the THP1 macrophage-HT29 incubation (37°C, 3 h) confirmed that phagocytosis of the CD47-expressing apoptotic HT29 cells (4–8 h post-UV) was associated with induction of the anti-inflammatory cytokine IL-10 while suppression of IL-6 and TNFα. Conversely, phagocytosis of necrotic cells collected at 18 h post-UV was associated with significant induction of proinflammatory IL-6 and TNFα. Similar results were obtained when testing THP-1 phagocytosis of apoptotic and necrotic HFF-1, HMEC-1, and various breast cancer cells (data not shown). Together, these results suggest that, while CD47 expression remains on apoptotic cells, it does not prevent macrophage clearance, and that phagocytosis of CD47-expressing apoptotic cells is a physiologically relevant, immunosuppressive event, whereas phagocytosis of necrotic cells is associated with proinflammatory conditions.

Figure 3.

A-B) Phagocytosis of apoptotic cells and necrotic cells by THP-1 macrophages. HT-29 cells loaded with CFSE (green dye) were collected before UV irradiation (CTL) or at 4, 6, 8 and 18 h post-UV irradiation and then incubated with THP-1 macrophages. After 30 min (37°C), the dishes were gently washed with HBSS and THP-1 macrophages were stained with a PE-conjugated anti-CD11b antibody (red). Represented images (A) and quantitative analysis (B) of phagocytosis of non-apoptotic cells (CTL), apoptotic cells (4–8 h post-UV) and necrotic cells (18 h post-UV) by THP-1 macrophages are shown. C) Cytokine production by THP-1 macrophages during phagocytosis. HT-29 cells (1 × 10⁵) collected without UV irradiation (CTL) or at 0, 2, 4, 6, 8 and 18 h post-UV irradiation were incubated with THP-1 macrophages (2 × 10⁵) at 37°C. After 3 h, the cell-free supernatants were collected from the HT29- macrophage incubation and tested for cytokines by ELISA. The data represent three independent experiments with triplicates in each detecting point. *** P<0.001; ** P<0.01. D) Apoptotic cells failed to trigger SIRPα ITIM tyrosine phosphorylation and/or SIRPα association with SHP-1 in THP-1 macrophages. Healthy and apoptotic (6 h post-UV) HT29 cells (1 × 10⁵) were incubated with THP-1 macrophages (2 × 10⁵) for 5 min at 37°C. The cells were then treated with freshly prepared pervanadate (2 mM) for 3 min followed by lysis. SIRPα was immunoprecipitated (IP) by anti-SIRPα.ex and protein A-sepharose followed by WB detections of SIRPα (anti-SIRPα.ex), SIRPα phosphorylation (mAb 4G10) and SHP-1 (rabbit anti-SHP-1 Ab). The data represent five independent experiments.

Although CD47 molecules were on the surface of apoptotic cells, they were incapable of ligating SIRPα and inducing the SIRPα-mediated inhibitory signaling in macrophages. Addition of a SIRPα-binding CD47 extracellular domain (CD47-AP) and a SIRPα-ligating mAb (SE7C2), both capable of triggering SIRPα-mediated inhibitory signaling, inhibited macrophage phagocytosis of apoptotic cells (Figure 3B). As shown in Figure 3D, direct incubation of apoptotic cells with THP-1 macrophages failed to trigger SIRPα ITIM tyrosine phosphorylation (SIRPα^pY) and/or recruitment of SHP-1. The same manner of co-incubating healthy, non-apoptotic cells with macrophages resulted in SIRPα phosphorylation and SIRPα association with SHP-1, suggestive of the successful induction of SIRPα-mediated signaling in macrophages. Consistent with their inhibition in phagocytosis, addition of CD47-AP and SE7C2 into the THP-1-apoptotic cell co-incubation led to SIRPα phosphorylation and SHP-1 association.

CD47 on apoptotic cell surface has low binding avidity to SIRPα

Since CD47 molecules on the apoptotic cells do not prevent macrophage phagocytosis, we speculate that CD47 on apoptotic cells might not interact with SIRPα on macrophages. To test this possibility, we first evaluated whether CD47 molecules in apoptotic cells were intact and could bind to SIRPα. CD47 was affinity-purified from various, non-apoptotic and apoptotic cells, and then tested for binding to SIRPα using a recombinant SIRPα extracellular domain fusion protein (SIRPα.ex-Fc). As shown in Figure 4, CD47 proteins purified from normal cultured, non-apoptotic cells and UV-induced apoptotic cells displayed equal binding to SIRPα.ex-Fc. The binding interactions between the CD47 and SIRPα extracellular domains were confirmed by including the extracellular domain-specific, inhibitory mAbs for CD47 (B6H12.2) or SIRPα (SE5A5) that abrogated the binding.

Figure 4.

Purified CD47 from apoptotic cells displayed an intact binding ability to SIRPα. CD47 was affinity purified from normal (Normal) cultured cells including HT-29, T47D and MCF7 and freshly isolated human PMN (hPMN), and apoptotic cells (Apop.) including UV-irradiated HT-29, T47D and MCF7 cells (6 h post UV), and apoptotic hPMN (overnight on ice with 20 mM EDTA). The purified CD47 samples were coated on microtiter plates followed by testing for binding to SIRPα extracellular domain using SIRPα.ex-Fc. Binding to the BSA-coated wells served as control. The anti-SIRPα antibody (SE5A5) and anti-CD47 antibody (B6H12.2) were used to block the specific SIRPα-CD47 binding. Data are presented as the mean ± SD (n=3).

Although the above assays using purified CD47 suggested that the binding capacity of CD47 to SIRPα was not affected by apoptosis, the cell adhesion assays suggested otherwise. As shown in Figure 5, A-C, stationary incubation of immobilized SIRPα (SIRPα.ex-Fc-coated wells) with healthy cultured, non-apoptotic cells resulted in effective, CD47-SIRPα interaction -mediated cell adhesion. In support that the cell adhesion was specifically mediated by CD47-SIRPα interaction, the extracellular domain inhibitory mAbs against CD47 (B6H12.2) and SIRPα (SE5A5) were applied and these mAbs completely blocked healthy cells from adhesion to immobilized SIRPα.ex-Fc. However, the same manner of SIRPα.ex-Fc-coated wells completely failed to mediate adhesion of apoptotic cells (Figure 5, A-C). Interestingly, apoptosis did not diminish cell adhesion mediated by lectin or anti-CD47 antibody (Figure 5D). We further analyzed the differences of apoptotic cells and non-apoptotic cells in adhesion to immobilized SIRPα by SPR. For these experiments, healthy cultured cells and apoptotic cells of the same type were set to flow through CM5 chip surfaces coated with SIRPα.ex-Fc. As shown in Figure 5E, non-apoptotic cells exhibited rapid attachment/retention to the chip and demonstrated effective, CD47-mediated cell adhesion to SIRPα. In contrast, apoptotic cells displayed no evident retention/attachment to the SIRPα.ex-coated chip under flow. These results conclude that apoptotic cells lost the capability to mediate strong (high avidity) cell surface binding to immobilized SIRPα via CD47, resulting in being washed away by the flow shear force.

Figure 5.

CD47 on the apoptotic cell surface failed to mediate high avidity cell binding to immobilized SIRPα extracellular domain. A-C) CD47-mediated cell adhesion to SIRPα. SIRPα.ex-Fc was immobilized on 96-well plates. After blocking, cells (2.5×10⁴/per well) of different types were added in the presence or the absence of anti-CD47 mAb (B6H12.2) or anti-SIRPα mAb (SE5A5) (10 µg/ml of each) followed by incubation for 30 min. Non-adherent cells were then removed by washing. The figure shows representative images (A-B) of cell adhesion to SIRPα.ex-Fc-coated wells or BSA (control) under various conditions. For quantitative analysis (C), cells were loaded with BCECF prior to incubation in SIRPα.ex-Fc coated wells. Fluorescent intensities before and after washing were recorded by a fluorescence plate reader and were used to calculate percentiles of cell adhesion (22). Data represent cell adhesion results from five independent experiments. D) Apoptotic cells adhered to immobilized lectins and anti-CD47 antibody but not SIRPα.ex-Fc. Lectin from Psophocarpus tetragonolobus (50 µg/ml in HBSS) and anti-CD47 mAb PF3.1 (10 µg/ml in HBSS) were coated on 96-well plates followed by blocking and incubation with healthy and apoptotic (6 h post-UV) HT29 cells. **, P < 0.01. E) SPR analysis of CD47-mediated cell adhesion to immobilized SIRPα.ex-Fc under flow in the presence or absence of anti-SIRPα mAb SE5A5. The adhesion curves represent the similar results from three independent experiments. F) Adhesion of different cells to immobilized SIRPα.ex-Fc. G) Cell binding to SIRPα.ex-Fc in solution. HT29 cells with and without UV irradiation were incubated with SIRPα.ex-Fc dissolved in HBSS (final concentration of 5 µg/ml) for 30 min. After washing, cells were labeled with AlexaFluor-conjugated secondary antibody followed by FACS. The binding specificity through CD47-SIRPα interaction was confirmed by including the inhibitory anti-CD47 (B6H12.2) or anti-SIRPα (SE5A5) antibody in the assays (data not shown).

Multiple additional types of cells were tested in cell adhesion assays and all yielded the similar results indicating that apoptosis led to loss of high avidity, CD47-mediated binding to SIRPα (Figure 5F). However, despite of failure to mediate cell adhesion, incubation of SIRPα.ex-Fc in solution with apoptotic cells resulted in SIRPα-CD47 binding (Figure 5G) in a manner similar to that of anti-CD47 antibody labeling. This result again indicates that failure of apoptotic cells to mediate the firm adhesion to the immobilized SIRPα is not due to depletion of CD47 on apoptotic cells or the change of CD47 extracellular domain structure, but the protein binding avidity.

Distribution change of CD47 from ‘cluster’ to ‘diffuse’ pattern on cell surface during apoptosis

Our previous study found that SIRPα forms clusters on the macrophage surface and such formation of SIRPα plays a key role in mediating high avidity cell interactions with CD47 (16). In this study, we asked whether CD47 also clusters on the cell surface and if such distribution is essential for SIRPα ligation and triggering SIRPα-based inhibitory signaling. We compared the cell surface distribution patterns of CD47 on non-apoptotic cells and apoptotic cells. To avoid possible antibody cross-linking, we used Fab fragments of both anti-CD47 and fluorophore-conjugated secondary antibodies to label cell surface CD47. As shown in Figure 6A, immunofluorescence microscopy revealed that CD47 on non-apoptotic HT29 cells was distributed in an uneven, punctate pattern, suggestive of cluster formation (Figure 6A, left panel, arrowheads). In contrast, CD47 on apoptotic cell surfaces were diffused resulting in relatively weak visual effects when microscopically analyzing the staining images (Figure 6A, right panel), despite that the actual level of CD47 was not significantly changed. Analyses of multiple other types of cells found the same CD47 distribution change before and after apoptosis (Figure 6B). Using NIH Image J and Nikon camera software (NIS-Elements B.R. 4.20.00) to analyze cell surface CD47 distribution and CD47 clusters/granules revealed remarkable differences between normal non-apoptotic cells and apoptotic cells. As shown in Figure 6C, Image J analysis showed that both the number and the sizes of CD47 granules/clusters on the cell surfaces were significantly decreased after apoptosis. In particular, CD47 granules/clusters larger than 50 pixel² were distributed on non-apoptotic cells but deficit on apoptotic cells. In contrast, apoptotic cells were chiefly distributed with small CD47 ‘dusts’ of sizes < 20 pixel². Figure 6D shows that on non-apoptotic cells, cluster-formed CD47 emitted high intensity, scattered peaks of fluorescence, whereas on apoptotic cells diffused CD47 emitted broad but weak fluorescence. As depicted in Figure 6E, this marked change of CD47 distribution on the cell surface before and after apoptosis correlates with the alteration of CD47 binding avidity to SIRPα in adhesion assays, suggesting that while individual CD47 is capable of binding, the overall low binding avidity of CD47 on apoptotic cells is insufficient to hold the cell in place. For our previous studies demonstrated that SIRPα clusters on macrophages, thus it is essential that CD47 also forms clusters in order to mediate high avidity cell surface interactions and strong inhibitory signal transduction effectively suppressing macrophage phagocytosis. Apoptosis-associated dispersal of CD47 significantly reduces the avidity of SIRPα ligation and hence facilitates macrophage phagocytosis.

Figure 6.

Disruption of ‘clustering’ pattern of CD47 on the cell surface during apoptosis. A) Represented images of surface CD47 pattern before and after HT-29 cell apoptosis from five independent experiments. Note that CD47 was largely clustered in punctate particles with various sizes on the surface of non-apoptotic cells (left panel), whereas the punctate pattern of CD47 was disappearing and CD47 distributed diffusely on the surface of apoptotic cells (right panel). Due to the diffused distribution, the visual intensity of CD47 labeling on apoptotic cells was relatively weak. B) Represented images of the surface CD47 pattern on other cell types prior to and after induction of apoptosis. Note, cells were co-stained by anti-CD47 mAb (red) and YO-PRO-1 (green), and apoptotic cells were susceptible to YO-PRO-1 labeling in the nucleus (green). C) NIH Image J analysis of the number and the sizes of CD47 granules/clusters on the cell surfaces prior to and after cells undergone apoptosis. A hundred (100) cells of each type were analyzed simultaneously per assay, and the data represent three independent assays. Note, the large granules/clusters of CD47 on non-apoptotic cells were disappearing after cells undergone apoptosis. D) Analysis of fluorescence intensity of CD47 on the cell surface by Nikon camera software (NIS-Elements BR4.20.00). Note, healthy, no-apoptotic cells displayed scattered CD47 labeling peaks with high fluorescence intensities, whereas apoptotic cells displayed broad CD47 labeling patches with low fluorescence intensities. Fifty (50) cells of each type were analyzed simultaneously per assay, and the data represent more than five independent assays. E) A sketch demonstrating a possible link between cell surface pattern of CD47 and the CD47 binding avidity to SIRPα.

Disrupting lipid rafts results in dispersion of surface CD47 clusters and reduction of CD47 binding avidity to SIRPα

It has been widely reported that clustering of cell membrane receptors requires the location of receptors in lipid rafts in the plasma membrane and such lipid raft-supported receptor clustering plays an essential for maintaining the structure and function of receptors (39–41). To test whether the clustering of CD47 on healthy cell surfaces is associated with lipid rafts and whether the dynamic distribution of lipid rafts is the mechanism underlying CD47 aggregation and/or disassembly, we performed several experiments. First, we double stained cell surface CD47 and lipid rafts using anti-CD47 Fab and FITC-conjugated cholera toxin B subunit, respectively, and studied their distribution patterns and potential co-localization. As shown Figure 7 (A-B), confocal microscopy taking images of HT29 cells at different focus layers (panels A & B) revealed that CD47 is largely co-localized with lipid rafts marked by cholera toxin B subunit in large punctate particles on healthy, non-apoptotic cell surface. This suggests that CD47 clustering is likely due to the assembly of CD47 within lipid rafts. Next, we disrupted lipid rafts with the cholesterol depletion agent MβCD and then examined the cell surface pattern of CD47. As can be seen, treating healthy HT-29 cells with MβCD (15 mM) led to a diffused staining pattern of FITC-conjugated cholera toxin B subunit, suggestive of lipid raft disruption in the plasma membrane (Figure 7, A-B). Clustering of CD47 on the cell surface also disappeared following MβCD treatment and CD47 staining displayed a diffuse pattern. This result therefore suggests that clustering of CD47 on the cell surface may be supported by lipid rafts. Analyses of the cell surface CD47 labeling using Image J and Nikon camera image software revealed that MβCD treatment caused CD47 distribution changes in a way similar to that of apoptotic cells, mainly diminishing large CD47 clusters that emitted high intensity of fluorescence (data no shown).

Figure 7.

‘Clustering’ punctate pattern of CD47 on epithelial cell surface is associated with and supported by lipid rafts in the plasma membranes. A-B) Confocal fluorescence images showing the surface distribution of CD47 (red) and lipid rafts (green) on non-apoptotic HT-29 cells with or without lipid raft ablation. Healthy cultured HT29 cells, with and without treatment with MβCD (15mM, 20min), were labeled without membrane permeabilization by mAb B6H12.2 for CD47 (red) and by FITC-conjugated cholera toxin B subunit for lipid rafts (green). Confocal images were taken at different cell layers and two images, one taken at the top layer (A) and one being at the middle of the cells (B), were shown. Note that without MβCD treatment, CD47 was largely co-localized with lipid raft at sizable punctate structures on cell surface, whereas disruption of lipid rafts by MβCD defused the ‘punctate’ localization of CD47 on the cell surface. Multiple types of cells were analyzed and the images represent typical results of five independent experiments. C) Disruption of cell surface lipid rafts by MβCD decreases cell adhesion to immobilized SIRPα.ex-Fc. CTL, cells without MβCD treatment. D) Disruption of cell surface lipid rafts by MβCD increases THP-1 macrophage phagocytosis towards the cells.

We next performed two additional experiments to confirm that non-apoptotic cells treated with MβCD would lose the high avidity CD47 binding to SIRPα and would promote macrophage phagocytosis. Cell adhesion assays were performed and cells with and without MβCD treatment were tested for adhesion to immobilized SIRPα.ex-Fc. As shown in Figure 7C, in contrast to HT-29, MCF7, T47D without MβCD treatment readily bound to SIRPα.ex-Fc, the same cells treated with MβCD completely lost stable cell adhesion to immobilized SIRPα.ex-Fc, indicating that raft disruption abrogated the high avidity cell surface CD47 binding. Phagocytosis assays (Figure 7D) by incubating MβCD-treated cells with THP-1 macrophages also showed enhanced phagocytosis of MβCD-treated cells compared to non-treated cells, and the presence of CD47-AP and anti- SIRPα mAb SE7C2, both ligating SIRPα, significantly inhibited THP-1 macrophage phagocytosis towards MβCD-treated cells.

Discussion

As a marker protein for self-recognition, CD47 plays a critical role in protecting host cells/tissues from being attacked by host phagocytes. Accumulating evidence has shown that ligation of macrophage SIRPα by CD47 expressed on the encountered cells prohibits macrophage phagocytosis, whereas failure of SIRPα engagement by CD47, or deficiency of SIRPα ITIM-mediated signaling, promotes macrophage engulfment of the host cell (1, 3, 42–45). Through over-expressing CD47 on the surface, certain types of cancer cells can escape from macrophage-mediated immune clearance (46, 47). It has been reported that cells likely lose their surface CD47 when aging or undergoing apoptosis, and this change results in abrogation of the CD47-SIRPα-mediated inhibitory signaling and hence facilitates macrophage phagocytic clearance.

However, contrary to previous reports, we demonstrate in the present study that the level of CD47 on the cell surface is not reduced during cell apoptosis; instead, the localization pattern of CD47 on the cell surface is drastically altered. On non-apoptotic cells, CD47 is largely clustered into numerous punctates on the cell surface. After apoptosis, the punctate pattern of CD47 disappears and replaced with a diffuse CD47 distribution on the cell surface. The shift of cell surface CD47 from ‘clustering’ punctates into diffuse distribution is a critical factor for the loss of CD47 capacity to suppress macrophage engulfment through ligating SIRPα. Our cell adhesion data derived under both static and flow conditions showed that, although apoptotic cells maintain the expression of CD47 on the cell surface, these CD47 proteins have a low binding avidity and are incapable of mediating firm cell adhesion to immobilized SIRPα. On the contrary, the CD47 molecules expressed on healthy, non-apoptotic cells display a high binding avidity and strongly hold cells to immobilized SIRPα even under a flow shear force (SPR technique). Consistent with these results, apoptotic cells showed failure to elicit CD47-SIRPα interaction–mediated inhibitory signaling indicated by the SIRPα cytoplasmic ITIM phosphorylation and association with SHP-1 when co-incubated with macrophages, whereas macrophages co-incubating with non-apoptotic cells resulted in SIRPα signaling (Figure 3D). Moreover, we show that loss of CD47 binding avidity on apoptotic cells is not due to biochemical modifications or defect of CD47 molecules, for purified CD47 from either healthy or apoptotic cells displayed the same binding activities towards SIRPα. As SIRPα also present as clusters on macrophages (16), it only makes sense that both these counter-receptors cluster on the cell surfaces in order to mediate high-density ligations effectively driving a strong signal transduction leading to inhibition of macrophage phagocytosis. In addition, a previous study by Burger et al. (13) showed that CD47 on erythrocytes undergo a conformational change that alters CD47 binding avidity to TSP-1. Those interesting results together with our results in the present study reveal a more complex role for CD47 in cell apoptosis and/or senescence, as well as the determination of phagocytic clearance of apoptotic or senescent cells by phagocytes.

Interestingly, we observed that while apoptosis does not lead to CD47 loss, the subsequent cell necrosis following apoptosis (‘secondary necrosis’) is associated with reduction of CD47. However, necrosis is also associated with significant reduction of the cell sizes. Therefore, decreases in CD47 on necrotic cells could be the result of reduction of the cell sizes and the loss of plasma membrane, even though CD47 molecules / per membrane area are not reduced. Our data also suggest that the loss of cell sizes during necrosis is likely due to the progressive membrane destabilization resulting in formation of large membrane blebs that eventually detach from the main cell bodies. Indeed, membrane blebs instantly appear when cells start apoptosis, and this result is consistent with observations by others (38, 48). As shown in our study, formation of blebs around the cell body appeared at 4 h post-UV, the same time point when other apoptotic signs such as surface PS and PARP-1 cleavage became apparent. To us, the appearance of these blebs becomes another indicator, which is easily visualized, for detecting apoptosis in addition to PS and PARP. As suggested by Charras et al. (37), membrane blebs are a result of membrane dynamics during which the plasma membrane transiently dissociates/ruptures from the underlying cytoskeleton cortex due to non-uniform intracellular hydrostatic pressure. Formation of membrane blebs is not apoptosis-specific. Certain active cellular processes, e.g. cell migration and cell proliferation, or when the cytoskeleton cortex integrity is compromised (e.g. filamin depletion), are also associated with cell blebbing (49–51). We observed that from apoptosis to the subsequent secondary necrosis, plasma membrane blebs are increased in size (from small to large blebs), suggestive of increased levels of plasma membrane destabilization and membrane dissociation from the underlying cortex. We consider that the shrinkage of cell sizes during necrosis is likely due to progressive damage/diminishment of the membrane cortex and detachment of large membrane blebs from the cell body. Along these dramatic cellular changes, CD47 at the stage of apoptosis shows remaining on the cell body without distribution to blebs, in a way very similar to the pattern of actin as reported by Charras et al (37), suggesting that CD47 may through its cytoplasmic domain connect to the underlying cytoskeleton cortex. Not only are they CD47-negative, the blebs at the stage of apoptosis are small and dynamic (continuously appearing and disappearing). In contrast, at the stage of necrosis membrane blebs are large and tend to detach from the cell body, and these blebs are CD47-positive. Therefore, the decrease in CD47 on necrotic cells could be the result of shrinkage of the cell size and loss of the plasma membrane, even though CD47 molecules per membrane area may not reduce. This explains that even under necrosis the CD47 level on the remnant cell body is never diminished completely. In our study, we specially utilized YO-PRO-1 and PI to distinguish apoptosis from necrosis. To confirm our results, we also compared the cytokines released by macrophages during their phagocytosis of apoptotic or necrotic cells. As shown by our data, macrophage phagocytosis of CD47-expressing apoptotic cells is accompanied with release of IL-10, while phagocytosis of those CD47-reduced necrotic cells is associated with IL-6 and TNFα production.

Our data further revealed that the functional clustering of CD47 on the cell surface is supported by lipid rafts in the plasma membranes. Immunofluorescence staining showed that cell surface CD47 is largely co-localized with lipid rafts, and disruption of lipid rafts also leads to diffusion of CD47. Clustering of CD47 within lipid rafts is in agreement with the previous finding by McDonald et al. (52), who showed that CD47 is associated with cholesterol-rich lipid rafts on human ovarian carcinoma cells, and also recruits αvβ3 integrin and its associated signaling molecules into these microdomains in the plasma membranes. An original study by Green et al. (53) showed that cholesterol directly binds to the multiple membrane-spanning domain of CD47 and is an essential component of the αvβ3/CD47/G protein-signaling complex.

In conclusion, this study demonstrates that cell apoptosis does not reduce CD47 expression on the cell surface but disperses CD47 from clustering in lipid rafts into the plasma membrane, and that this change of CD47 distribution on the cell surface results in loss of the high binding avidity of CD47 to SIRPα and renders CD47 incapable to trigger SIRPα-mediated inhibitory signaling in macrophages and hence facilitates phagocytic clearance. For healthy cells in tissues, the clustering of CD47 on the cell surface is a functional form supported by lipid rafts in plasma membranes and this form of CD47 enables effective ligation to SIRPα and triggers SIRPα-mediated inhibition in macrophages.

The abbreviations used are

PMN

polymorphonuclear leukocyte

HBSS

Hank’s balanced salt solution

SIRP

signal regulatory protein

SHP

Src homology 2 domain-containing tyrosine phosphatase

mAb

monoclonal antibody

BSA

bovine serum albumin

fMLF

formyl-methionyl-leucyl-phenylalanine