Information

What is the end-point of material from an apoptotic cell, after phagocytosis?

What is the end-point of material from an apoptotic cell, after phagocytosis?



We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

Apoptosis occurs. The organelles and interior material form apoptotic bodies that are packed in vesicles. The cell membrane breaks apart (cell no longer exists) and apoptotic bodies enter the extracellular space. Phagocytes "eat" the apoptotic bodies.

What happens then?

Is the material processed again and effluxed into the blood, sent to other cells,… ?


Frontiers in Immunology

The editor and reviewers' affiliations are the latest provided on their Loop research profiles and may not reflect their situation at the time of review.


  • Download Article
    • Download PDF
    • ReadCube
    • EPUB
    • XML (NLM)
    • Supplementary
      Material
    • EndNote
    • Reference Manager
    • Simple TEXT file
    • BibTex


    SHARE ON

    Enhanced efferocytosis of apoptotic cardiomyocytes through myeloid-epithelial-reproductive tyrosine kinase links acute inflammation resolution to cardiac repair after infarction

    Rationale: Efficient clearance of apoptotic cells (efferocytosis) is a prerequisite for inflammation resolution and tissue repair. After myocardial infarction, phagocytes are recruited to the heart and promote clearance of dying cardiomyocytes. The molecular mechanisms of efferocytosis of cardiomyocytes and in the myocardium are unknown. The injured heart provides a unique model to examine relationships between efferocytosis and subsequent inflammation resolution, tissue remodeling, and organ function.

    Objective: We set out to identify mechanisms of dying cardiomyocyte engulfment by phagocytes and, for the first time, to assess the causal significance of disrupting efferocytosis during myocardial infarction.

    Methods and results: In contrast to other apoptotic cell receptors, macrophage myeloid-epithelial-reproductive tyrosine kinase was necessary and sufficient for efferocytosis of cardiomyocytes ex vivo. In mice, Mertk was specifically induced in Ly6c(LO) myocardial phagocytes after experimental coronary occlusion. Mertk deficiency led to an accumulation of apoptotic cardiomyocytes, independently of changes in noncardiomyocytes, and a reduced index of in vivo efferocytosis. Importantly, suppressed efferocytosis preceded increases in myocardial infarct size and led to delayed inflammation resolution and reduced systolic performance. Reduced cardiac function was reproduced in chimeric mice deficient in bone marrow Mertk reciprocal transplantation of Mertk(+/+) marrow into Mertk(-/-) mice corrected systolic dysfunction. Interestingly, an inactivated form of myeloid-epithelial-reproductive tyrosine kinase, known as solMER, was identified in infarcted myocardium, implicating a natural mechanism of myeloid-epithelial-reproductive tyrosine kinase inactivation after myocardial infarction.

    Conclusions: These data collectively and directly link efferocytosis to wound healing in the heart and identify Mertk as a significant link between acute inflammation resolution and organ function.

    Keywords: efferocytosis inflammation macrophages myocardial infarction phagocytosis.

    Conflict of interest statement

    Figures

    Figure 1. Macrophages phagocytose cardiomyocytes (CMs) and…

    Figure 1. Macrophages phagocytose cardiomyocytes (CMs) and Mertk is specifically required for CM efferocytosis

    Figure 2. Identification and kinetics of Mertk…

    Figure 2. Identification and kinetics of Mertk expression post MI in experimental mice

    Figure 3. Chemokine and cytokine mRNA and…

    Figure 3. Chemokine and cytokine mRNA and inflammatory cells in hearts from Mertk +/+ vs.…

    Figure 4. Quantitation of cardiomyocyte (CM) apoptosis…

    Figure 4. Quantitation of cardiomyocyte (CM) apoptosis and association with CD68+ phagocytes in Mertk +/+…

    Figure 5. Acute myocardial infarct size is…

    Figure 5. Acute myocardial infarct size is increased in Mertk -/- mice post MI

    Figure 6. Quantitation of scar formation in…

    Figure 6. Quantitation of scar formation in remodeled hearts post myocardial infarction (MI) in Mertk…

    Figure 7. Assessment of heart function by…

    Figure 7. Assessment of heart function by echocardiography after myocardial infarction (MI) in Mertk deficient…


    Materials And Methods

    Preparation of Recombinant MFG-E8.

    Recombinant MFG-E8-L and its mutant proteins were prepared as described previously (18). In brief, the expression plasmid of wild-type MFG-E8-L, D89E carrying RGE sequence instead of RGD in the second EGF-like domain, or E1E2PT lacking the C1C2 domains were introduced into human 293T cells by the calcium phosphate precipitation method. The culture medium was replaced with DMEM/2% FCS 16 h after transfection, and the transfected cells were cultured for another 48 h. The recombinant proteins secreted into the medium were purified using anti–FLAG M2 affinity gel (Sigma-Aldrich), or anti–MFG-E8 antibody (clone 2422)–conjugated protein A–Sepharose 4FF beads (Amersham Biosciences). To examine the purity of the proteins, they were subjected to SDS-PAGE and Western blotting. For SDS-PAGE, proteins were separated by electrophoresis on a 10/20% polyacrylamide gradient gel and stained with Coomassie brilliant blue. For Western blotting, proteins were separated by electrophoresis, transferred to PVDF membranes, and detected with anti-FLAG antibody. We confirmed that the recombinant proteins alone did not stimulate the production of inflammatory cytokines including TNFα and IL-1β in macrophages.

    In Vitro Phagocytosis Assay and Measurement of Cytokines Production.

    For the preparation of bone marrow–derived macrophages (BMDMs), bone marrow cells were obtained by flushing the femurs from 8–10-wk-old C57BL/6 mice. The cells were treated with RBC lysis buffer (17 mM Tris-HCl, pH 7.5, 144 mM ammonium chloride, and 0.5% FCS) for 1 min at room temperature. Next, the cells were suspended in αMEM/10% FCS medium, and were plated at a density of 10 6 cells/ml in the presence of recombinant mouse M-CSF (a gift from A. Kudo, Tokyo Institute of Technology, Kanagawa, Japan). Cells were harvested on day 3, diluted 1:10 with the medium, and cultured for another 3 d at 37°C. On day 6, the cells were used for phagocytosis assay. 2.5 × 10 5 BMDMs were seeded in a 48-well cell culture cluster (Corning Inc.) and cultured overnight at 37°C. Cells were preincubated with or without D89E (1, 2, and 4 μg/ml) or E1E2PT protein (4 μg/ml) for 30 min. For the preparation of apoptotic cells, thymocytes from 4 to 8–wk-old CAD-deficient mice (6) were incubated with 10 μM dexamethasone at 37°C for 4 h. 2.5 × 10 6 of apoptotic thymocytes were added to BMDM cultures, and phagocytosis was allowed to proceed for 2 h. Cells not being engulfed were removed by washing with PBS, and the BMDMs were detached with 1 mM EDTA/PBS. Afterwards, the cells were stained with PE-conjugated anti–mouse CD11b, followed by TUNEL staining as described previously (18).

    Resident peritoneal macrophages were suspended in DMEM/10% FCS and 10 5 cells were seeded in a 96-well cell culture cluster. The cells were incubated 3 h at 37°C and used for the phagocytosis assay.

    For cytokine assay, 10 5 cells of thioglycollate-elicited peritoneal macrophages from C57BL/6 were seeded in a 96-well cell culture cluster. The macrophages were preincubated with or without D89E or E1E2PT protein for 30 min. Then, 2 × 10 6 cells of apoptotic thymocytes induced by UV irradiation were added to the macrophages for phagocytosis. After 2-h incubation, the macrophages were washed twice, and were stimulated with or without 1 μg/ml LPS for 20 h. For the measurement of TGF-β, the culture medium was changed to AIM-V (Invitrogen) 1 h after stimulation with or without LPS, and the cells were incubated for 20 h. Cytokine concentrations in the culture supernatants were measured using ELISA kits for IL-10, TNFα (BD Biosciences), and TGF-β (R&D Systems) according to manufacturer's protocols.

    Injection of Recombinant Proteins and Apoptotic Cells.

    The purified recombinant proteins were diluted with PBS containing 2.5% normal mouse serum obtained from C57BL/6 mice, and 300 μl of the solution was intravenously injected into 8-wk-old C57BL/6 female mice through the tail vein. In the case of apoptotic thymocyte injection, thymi from 4 to 6–wk-old C57BL/6 mice were removed and squeezed between glass slides. Thymocytes were then filtrated through a nylon mesh, and suspended in serum-free RPMI 1640 medium. The cells were irradiated with 40 J/m 2 UV light to induce apoptosis and were cultured in RPMI 1640 medium containing 1% normal mouse serum at 37°C for 20 h. Next, the cells were washed twice with PBS containing 1 mg/ml of mouse serum albumin and suspended in PBS containing 2.5% normal mouse serum. Recombinant MFG-E8 was added to apoptotic thymocytes 30 min before injection, and the cells were intravenously injected into mice. Immunizations were performed weekly for a total of four to six injections.

    Detection of Autoantibodies

    The serum levels of anticardiolipin antibody and anti-PS antibody were detected by ELISA. 96-well ELISA plates (Immulon 1B microtiter plate ThermoLabsystems) were coated with 10 μg/ml cardiolipin (CL) in methanol or 10 μg/ml l -α-phosphatidyl- l -serine, dioleoyl (Sigma-Aldrich) in ethanol. After blocking with 10% FCS, mice sera diluted 50 times with PBS were added and incubated for 1 h at room temperature. The mouse antibodies bound to the plate were detected using goat anti–mouse Ig conjugated to HRP (ICN Biomedicals) at a dilution of 1:2,000. The peroxidase activity was detected using o-phenylenediamine in the peroxidase detection kit (Sumitomo) as a substrate. The color reaction was read at 492 nm using a microplate reader (Titertek Instruments).

    Antinuclear antibody (ANA) was detected by indirect immunofluorescence and ELISA. For immunofluorescence, serum samples were diluted 50 times with PBS, and were added on glass slides coated with Hep-2 cells (MBL). The slides were incubated at 37°C in a humid chamber for 30 min. The antibodies bound to the slides were detected by Cy3-conjugated F(ab′)2 of goat anti–mouse IgG (Jackson ImmunoResearch Laboratories) diluted 100 times with PBS/10% normal goat serum. Slides were observed by fluorescence microscopy (model IX-70 Olympus).

    ANAs were also detected by using an ANA detection kit (MBL). Mouse sera diluted 100 times with the reaction buffer of the kit were added to a microcup coated with a mixture of human nuclear antigens (three types of RNP epitope [70k, RNP-A, and RNP-C], native Sm, native SS-A, recombinant SS-B, recombinant Scl-70, cenp-B, recombinant Jo-1, and γ phage DNA antigen) and incubated for 1 h at room temperature. The antibodies bound to the plate were detected using goat anti–mouse Ig's conjugated to HRP (ICN Biomedicals) at a dilution of 1:1,000. The peroxidase activity was detected using TMB as a substrate. The color reaction was read at 450 nm using a microplate reader (Titertek Instruments).

    Anti–double stranded (ds) DNA antibody was detected by using anti-dsDNA detection kit (MBL). Mouse sera diluted 100 times with the reaction buffer of the kit were added to a microcup coated with human dsDNA and incubated 1 h at room temperature. The antibodies bound to the microplate were detected using goat anti–mouse Ig's conjugated to HRP (ICN Biomedicals) at a dilution of 1:1,000. The peroxidase activity was detected as described in the previous paragraph. The color reaction was read at 450 nm using a microplate reader (Titertek Instruments).

    Immunohistochemistry

    20–36-wk-old mouse kidneys were fixed with 4% paraformaldehyde/4% sucrose in 0.1 M phosphate buffer, pH 7.2, and embedded in paraffin. 4-μm-thick sections were prepared and mounted on silanized slide glasses. For immunohistochemistry, sections were incubated for 60 min at room temperature in PBS containing 0.1% Triton X-100 and 10% normal goat serum and were stained for 60 min at room temperature with Cy3-conjugated F(ab′)2 of goat anti-mouse IgG (Jackson ImmunoResearch Laboratories) used at a dilution of 1:100. The sections were washed three times with PBS containing 0.1% Triton X-100 and observed by fluorescence microscopy.


    Abstract

    Alzheimer's disease (AD) is characterized by the accumulation in the brain of extracellular amyloid β (Aβ) plaques as well as intraneuronal inclusions (neurofibrillary tangles) consisting of total tau and phosphorylated tau. Also present are dystrophic neurites, loss of synapses, neuronal death, and gliosis. AD genetic studies have highlighted the importance of inflammation in this disease by identifying several risk associated immune response genes, including TREM2. TREM2 has been strongly implicated in basic microglia function including, phagocytosis, apoptosis, and the inflammatory response to Aβ in mouse brain and primary cells. These studies show that microglia are key players in the response to Aβ and in the accumulation of AD pathology. However, details are still missing about which apoptotic or inflammatory factors rely on TREM2 in their response to Aβ, especially in human cell lines. Given these previous findings our hypothesis is that TREM2 influences the response to Aβ toxicity by enhancing phagocytosis and inhibiting both the BCL-2 family of apoptotic proteins and pro-inflammatory cytokines. Aβ42 treatment of the human microglial cell line, HMC3 cells, was performed and TREM2 was overexpressed or silenced and the phagocytosis, apoptosis and inflammatory response were evaluated. Results indicate that a robust phagocytic response to Aβ after 24 h requires TREM2 in HMC3 cells. Also, TREM2 inhibits Aβ induced apoptosis by activating the Mcl-1/Bim complex. TREM2 is involved in activation of IP-10, MIP-1a, and IL-8, while it inhibits FGF-2, VEGF and GRO. Taken together, TREM2 plays a role in enhancing the microglial functional response to Aβ toxicity in HMC3 cells. This novel information suggests that therapeutic strategies that seek to activate TREM2 may not only enhance phagocytosis and inhibit apoptosis, but may also inhibit beneficial inflammatory factors, emphasizing the need to define TREM2-related inflammatory activity in not only mouse models of AD, but also in human AD.


    No-wash assays

    A very fast and highly accurate way to monitor stages in the phagocytosis pathway uses pHrodo indicators. pHrodo Deep Red, Red, and Green are conjugated to a range of particles for phagocytosis measurement with no quench or wash required.

    pHrodo dyes are essentially non-fluorescent at neutral pH and exhibit increasing signal with a red or green readout respectively as the pH decreases. The increase in fluorescent signal can be used to monitor progression in the phagocytic pathway.


    Clinical Trials Involving EVs and Mitochondria Transfer

    All the clinical trials concerning MSC-EVs and ATM can be found at www.clinicaltrails.gov. Although the majority of listed trials focus on the diagnostic properties of EVs, there are five trials testing the therapeutic applications of MSCs-EVs and two proposing the use of ATM (Table 2).

    A first trial is designed to test the anti-inflammatory properties of umbilical cord derived MSCs-EVs to prevent the destruction of pancreatic β-cell islets. The MSCs-EVs will be administered intravenously in two doses, the first dose of exosomes and, after seven days, the second dose of microvesicles (NCT02138331). Two other clinical trials using EVs will involve allogeneic MSCs-EVs. One of them will administer EVs enriched by miR-124 for the treatment of acute ischemic stroke (NCT03384433). The second clinical trial will attempt to treat lesions in patients affected by dystrophic epidermolysis bullosa (NCT04173650). The last clinical trial using EVs that is in the recruiting phase focuses on promoting the healing and recovery of refractory macular holes through direct injection of MSC exosomes to the site of the injury (NCT03437759). Finally, the only concluded trail to date used umbilical cord derived MSCs-EVs to inhibit the progression of chronic kidney disease in patients with grade III-IV CKD [64]. The study showed stabilization of the disease progression, as confirmed by stable levels of glomerular filtration rate, serum creatinine and blood urea in treated patients, and an increased level of anti-inflammatory factors (TGF-β1 and IL-10) in comparison with the matching placebo group.

    The first clinical trial using administration of isolated mitochondria for the treatment of myocardial IRI has also been concluded with positive results [65]. Mitochondria were isolated from non-ischemic skeletal muscles and injected in the myocardium of paediatric patients with myocardial IRI. No adverse effects were detected after AMT, and four out of five patients demonstrated an enhancement in ventricular function [65].

    Other clinical trials using AMT are focused on the improvement of infertility treatments (Table 2). Through autologous micro-injection of mitochondria prior to intra-cytoplasmic sperm injection, the patients’ oocyte quality was enhanced. In the first trial, concluded in 2017, mitochondria were isolated from autologous ovarian stem cells and directly injected into the oocytes themselves. To date, no results have been published. Embryo quality has been quantified through the pregnancy rate after treatment and morphological evaluation of the treated embryos. In the second clinical trial, which is still ongoing, mitochondria will be isolated from autologous bone marrow-MSCs and administrated immediately before intra-cytoplasmic sperm injection in the oocytes. Live birth rate, pregnancy rate, number of oocytes retrieved and fertility rate are going to be evaluated.


    Results

    Immature DCs Efficiently Phagocytose Apoptotic Cells.

    Based on previous observations that immature DCs are the cells responsible for capturing antigen (11), we predicted that apoptotic cells would be engulfed best by immature DCs. To test this hypothesis, we established a phagocytosis assay that allowed us to visually detect the uptake of apoptotic cells, and compare the phagocytic capacity of immature DCs, mature DCs, and macrophages. In brief, immature DCs were prepared by culturing a T cell–depleted fraction from peripheral blood in the presence of IL-4 and GM-CSF. Mature DCs were generated with the addition of MCM and these cells expressed the cell surface DC-restricted maturation marker CD83 (18, 19, 30). Macrophages were prepared by culturing a plastic adherent cell population in Teflon beakers for 3–9 d. As a source of apoptotic cells, we used influenza-infected monocytes (7) virus infection induces apoptotic death in these cells within 6–10 h (7, 25, 26). Monocytes were first infected with influenza virus as previously described (24), then dyed red using PKH-26 (Sigma Biosciences). After 6–8 h, the various APCs were dyed green using the fluorescent cell linker compound PKH67-GL (Sigma Biosciences) and cocultured with the apoptotic cells at a ratio of 1:1. After 2 h at 37°C, cocultures of cells were analyzed by FACScan ® analysis, allowing for quantification of phagocytic uptake as double positive cells. 80% of the macrophages, 50% of the immature DCs, and <10% of the mature DCs engulfed the apoptotic monocytes after 2 h of coculture (Fig. 1,A). The smear of double positive cells (PKH67-labeled APCs that engulfed the PKH26-labeled apoptotic cells) indicates that both apoptotic bodies and whole apoptotic cells served as ‘food' for the phagocytic cell (Fig. 1, iii, vi, and ix). Note that as the forward scatter of the APCs increased and the setting of the FACS ® shifted, the dying monocytes were excluded from the established region (Fig. 1, ii, v, and viii). Maximal uptake by all APC populations was achieved within 2–4 h and partially depended upon the source of apoptotic cell used (Fig. 1,B and data not shown). Given this kinetic data, we believe that macrophages and DCs engage and internalize dying cells while still displaying features of early apoptotic cell death. This data also demonstrates that it is the immature DC that preferentially acquires apoptotic material compared with the mature DC. The source of apoptotic cells was not critical, since we obtained similar results with UVB-irradiated HeLa cells (see Fig. 7, and data not shown).

    To confirm that this FACS ® assay was measuring phagocytosis, we carried out the assay at 4°C and in the presence of inhibitors of phagocytosis. Both low temperature (Fig. 2,A) and cytochalasin D, an inhibitor of cytoskeletal function, blocked uptake (Fig. 2,B). Phagocytosis by immature DCs also requires divalent cations as EDTA was inhibitory (Fig. 2,C). To visually confirm the uptake recorded by FACS ® , we prepared cytospins of the dyed cocultures. The frequency of uptake correlated with that measured on FACS ® (data not shown). We also performed immunofluorescence on cocultures of immature DCs labeled with anti– HLA-DR (DR) and apoptotic influenza-infected monocytes labeled with antiinfluenza nucleoprotein (NP) (Fig. 3). In the top panel an apoptotic cell is seen just prior to being engulfed by a DC (arrowhead). After phagocytosis, apoptotic cells were found in DR + vesicles (arrows), but not in the cytoplasm.

    Only Immature DCs Cross-present Antigen from the Apoptotic Cell on Class I MHC.

    We next correlated the phagocytic capability of macrophages and DCs with their ability to cross-present antigenic material derived from apoptotic cells. The cells were prepared from HLA-A2.1 + donors (18, 19), cocultured with HLA-A2.1 − influenza-infected monocytes for 12 h, and then loaded with Na 51 CrO4 for use as targets for influenza-specific CTLs (7, 24). Specific lysis indicates that the APCs cross-presented antigenic material derived from the apoptotic cell by forming specific peptide–MHC class I complexes on its surface (Fig. 4,A). As a direct comparison with the endogenous pathway for class I MHC presentation, the same APC populations were infected with live influenza virus and used as targets (Fig. 4 B).

    Although mature DCs were efficient targets when infected with influenza, they were unable to cross-present antigens, presumably because they had downregulated the ability to phagocytose the apoptotic monocytes. However, the immature DCs did cross-present antigens from apoptotic cells. Furthermore, if the immature DCs were cocultured with the apoptotic cells in the presence of MCM, a maturation stimulus, they were even better targets. This is possibly due to the upregulation of costimulator and adhesion molecules (13, 31), or to the increased stability of peptide–MHC I complexes. Given that maximal uptake of apoptotic cells by immature DCs occurs between 2 and 4 h (Fig. 1 D), we believe that cross-presentation of apoptotic material reflects the phagocytosis and processing of early apoptotic cells rather than secondary necrotic cells (see Materials and Methods). With respect to this issue, it is important to recognize that the influenza-infected monocytes require 24 h to undergo secondary necrosis (Albert, M.L., and N. Bhardwaj, unpublished data references 25, 26).

    Notably, macrophages that efficiently phagocytose apoptotic cells (Fig. 1,A) did not cross-present antigens to CTLs (Fig. 4 B). Presumably, the engulfed material is degraded, not cross-presented, on MHC I. This profound difference between the DC and macrophage populations is supported by our previous findings that macrophages do not cross-present antigens from apoptotic cells during the induction phase of a class I–restricted antigen-specific T cell response. In fact, when put into culture with DCs in a competition assay, they sequester the apoptotic material and abrogate the CTL response (7).

    Immature DCs Can Be Distinguished from Macrophages by Intracellular Expression of CD83 and a Unique Profile of Phagocytic Receptors.

    We investigated the possibility that immature DCs might phagocytose apoptotic cells via pathways distinct from macrophages. To clearly distinguish these cells, we characterized them phenotypically. Immature DCs are distinguished by the absence of both CD14, a macrophage restricted marker, and CD83, a maturation marker for DCs (30). We have extended the use of CD83, finding that immature DCs can be distinguished from both macrophages and mature DCs by their intracellular expression of CD83. Macrophages do not express CD83 intra- or extracellularly, whereas mature DCs express CD83 both intra- and extracellularly (Fig. 5).

    These APC populations were examined for surface expression of receptors involved in phagocytosing apoptotic material (Table 1). These include: αvβ3 and CD36, which act as coreceptors for engulfment of apoptotic neutrophils and lymphocytes by macrophages (32, 33) and CD14, which has been implicated in the uptake of apoptotic cells by macrophages (34). While studying the immature DC populations, we identified a discrepancy in the expression of the αv and β3 integrin chains and investigated the possibility that αv was binding an alternate β chain. Using antibodies that recognize combined epitopes of the αvβ3 and the αvβ5 heterodimers, we noted the selective expression of αvβ5 on immature DCs (Fig. 6,A). As is true for most receptors involved in antigen uptake (11, 12), the expression of CD36, αvβ5, and mannose receptor on DCs is downregulated with maturation (Fig. 6,B, Table 1).

    To evaluate whether this downregulation could be observed on the level of mRNA expression, we performed reverse transcriptase PCR using primers specific for β3, β5, and CD36 (Fig. 6 C). Immature DCs (lane 1) showed amplified DNA of the appropriate size for β3, β5, and CD36. In contrast, in mature DCs (lane 2), no β5 and much fewer CD36 sequences were seen, whereas β3 sequences were comparable to those in immature cells. These data, although not quantitative, are consistent with the levels of protein expression observed by FACS ® and suggest that phagocytic receptor expression in DCs may be regulated at a transcriptional level as mRNA expression of CD36 and β5 is downregulated during maturation.

    Αvβ5 and CD36 Mediate Phagocytosis of Apoptotic Cells in Immature DCs.

    To demonstrate a direct role for αvβ5 in the recognition of apoptotic cells by immature DCs, we performed the phagocytosis FACS ® assay in the presence of antibodies specific for αvβ5 (Fig. 7,A). In addition to the blocking observed using the mAb to αvβ5, blocking was also detected when using mAbs to αv, β5, and CD36. Blocking was not observed when isotype-matched mAbs were specific for β1, β3, or the transferrin receptor CD71. Note that control antibodies chosen recognized surface receptors present on the immature DCs (Fig. 7,A, Table 1). mAbs were tested in doses ranging from 10 to 80 μg/ml (data not shown). Maximal inhibition of phagocytosis of apoptotic cells was seen with mAbs specific for CD36, αv, and β5 at 50 μg/ml. The inhibition of phagocytosis of apoptotic cells by DCs was specific. We were unable to block the uptake of red fluorescent latex beads, a control particle, by DCs in the presence of these mAbs (Fig. 7 B). By histogram analysis, DCs phagocytose 1–6 particles per cell. mAbs to αvβ5 or αv did not alter the profile of these histogram plots (data not shown).

    Although some inhibition of phagocytosis was observed when using αvβ3 this may be due in part to transdominance and/or the effect on the pool of free αv (35). For example, anti-αvβ3 antibodies suppress the intracellular signaling of the α5β1 integrin (36). Alternatively, αvβ3 and αvβ5 may be working cooperatively in the immature DCs. We therefore tested combinations of anti-αvβ3 and anti-αvβ5 but did not observe an increase in the inhibition of phagocytosis. The low receptor density of αvβ3 on DCs (average mean fluorescence intensity of 7 ± 2 Table 1) also makes it unlikely that this integrin heterodimer is involved in the engulfment of apoptotic cells by immature DCs.

    Our data do not exclude a role for other receptors in the phagocytosis of apoptotic cells, e.g., the putative PS receptor or the lectin receptor (5). In fact, other receptors are probably involved, as blocking observed did not exceed 60% even when combinations of all relevant mAbs were tested (data not shown). CD14 is unlikely to be involved in the engulfment of apoptotic cells by DCs, as DCs do not express this receptor (Table 1). In macrophages, phagocytosis of apoptotic cells was inhibited by antibodies to αv, β3, αvβ3, and CD36 but not by antibodies to β1, β5, or αvβ5 (data not shown). This correlates with published data (6, 33).


    Results and discussion

    HS-induced plant cell death is morphologically distinct from apoptosis

    One of the earliest hallmarks of apoptosis is AVD [79, 80], followed by blebbing of the PM, nuclear segmentation and finally fragmentation of the cell into apoptotic bodies [17]. Since the ultimate goal of such cell dismantling is removal of the dying cell by phagocytes, the PM of the apoptotic cell should remain intact throughout the whole process, preventing spillage of the dying cell content. It is fairly obvious that formation of apoptotic bodies during plant cell death would serve no purpose, as plants lack phagocytes and a dead plant cell remains to be incapsulated within a rigid cell wall [25, 81]. However, death of plant cells caused by abiotic or biotic stress is accompanied by rupture of PM and/or vacuolar membrane, tonoplast, causing severe damage to the endomembrane systems and irreversible detachment of the PM from cell wall which is often referred to as the protoplast shrinkage [11, 82]. This phenomenon has been repeatedly confused with AVD, PM blebbing and formation of apoptotic bodies, and thus exploited to support existence of AL-PCD in plants [38, 39].

    To examine AVD, PM blebbing, nuclear segmentation and cellular fragmentation in plants, we analysed morphology of tobacco BY-2 cells under HS conditions that were previously described to induce AL-PCD [48, 51]. Cell culture was stained with a noncell-permeable Sytox Orange (SO) nucleic acid dye to visualize cells with compromised PM integrity and with the styryl dye FM4-64 to visualize cell membranes and then subjected to a pulse HS at 55 °C (Additional file 1: Video S1).

    Only SO-positive cells exhibited protoplast shrinkage, strongly indicating correlation between cell volume decrease and PM permeabilization (Fig. 1a). Notably, the treated cells became SO-positive at the earliest checked time point, i.e. within 10 min of the HS indicating rapid PM permeabilization. One prominent feature of SO-positive cells was formation of vesicle-like structures at the inner side of the PM (Fig. 1a, b). Such morphology is inconsistent with AVD and blebbing, which require intact PM furthermore, apoptotic PM blebs form outwards on the cell surface [45].

    Gross morphological changes in HS-treated plant cells do not match hallmarks of apoptosis. a FM4-64 and SO dyes were used to visualize all cells and cells with permeabilized PM, respectively. BY-2 cells were imaged under control conditions (No HS) or after a 10-min HS at 55 °C and were stained according to protocols i and ii, respectively (see “Sytox Orange and FM4-64 staining”). Scanning was performed within 1 h post-HS. The arrows indicate PM note the PM (red dotted line) being tightly pressed to the cell wall (white dotted line) under control conditions and detached under stress conditions. b Higher magnification of the areas indicated with arrows in a. c BY-2 cells expressing green fluorescent protein (GFP) fused to nuclear localization signal (NLS) were subjected to the same treatments as in a. Images represent maximum intensity projections of z-stack scans acquired 6 h post-HS. d Quantification of nuclear area in samples shown in c. Data from three independent experiments, with ≥ 142 cells counted per treatment. Student’s t test, *p < 0.005. DIC, differential interference contrast microscopy. n, nucleus. IQR, interquartile range. Scale bars, 20 μm (a, c) or 5 μm (b)

    The PM and shrunken protoplast were additionally imaged in the time interval from 15 min to 72 h after the pulse HS (Additional file 2: Figure S1). Nevertheless, we failed to detect PM blebbing or protoplast fragmentation into discrete bodies at any time point. Furthermore, although we did observe moderate nuclear condensation upon HS, it was not followed by segmentation of the nucleus (Fig. 1c, d). In summary, the gross morphological changes of cells undergoing HS-induced cell death do not resemble apoptosis.

    PM integrity is irreversibly compromised during or shortly after pulse HS

    Intact PM is a pivotal hallmark of apoptosis [10]. However, SO staining described above suggested permeabilization of PM in most BY-2 cells already within 10 min of the HS. To investigate dynamics of the PM permeabilization, we analysed cellular content leakage after two types of HS, at 55 °C or 85 °C which were reported to induce AL-PCD or necrosis, respectively [61, 70]. The leakage of cellular content was assessed using a fluorescein diacetate (FDA)-based fluorochromatic assay [83]. In brief, the non-fluorescent FDA molecules can passively diffuse into the living cells where their acetate groups are cleaved off by esterases. The resulting fluorescein molecules have poor membrane permeability and are retained in cells with an intact PM, but are released into extracellular space upon PM permeabilization.

    We imaged BY-2 cells loaded with FDA prior to the pulse HS at 55 °C or 85 °C (Fig. 2a). Both types of HS caused rapid (within 10 min) leakage of the dye into the extracellular space (Fig. 2a). We measured the amount of fluorescein accumulated in the extracellular space immediately after the HS and found that the rate of cellular content leakage in HS-treated cells was comparable to that occurring after severe disruption of plant cells caused by freeze-thaw in liquid nitrogen (Fig. 2b).

    Both 55 °C and 85 °C HS cause instant and irreversible PM permeabilization. a Green fluorescence in the BY-2 cells loaded with FDA. Fluorescein leaks into extracellular space within 10 min after HS at either 55 °C or 85 °C. Arrows indicate shrunken protoplasts in 55 °C HS-treated cells. b Similar fraction of fluorescein leaks out of cells after 10 min of 55 °C, 85 °C or liquid nitrogen (N2) treatment. c BY-2 cells stained with SO and FM4-64 to assess whether disruption of the PM integrity after pulse HS is transient or irreversible. Staining was performed for no HS, before HS or after HS treatments according to protocols i, ii and iii, respectively (see “Sytox Orange and FM4-64 staining”). The cultures were imaged within 1 h following HS. d Frequency of the SO-positive cells in the cultures shown in c. DIC, differential interference contrast microscopy. IQR, interquartile range. Scale bars in a and c, 50 μm. b Representative data from one out of three independent experiments. d Data from three independent experiments, with ≥ 115 cells counted per treatment. b, d One-way ANOVA with Dunnet’s test *p < 0.005

    To determine whether PM permeabilization was transient or permanent, we added SO and FM4-64 stains to the cell cultures either before or 30 min after HS. We speculated that if PM permeabilization upon HS was transient, cultures stained after HS would show a significantly lower frequency of SO staining as compared to cells stained before HS. However, SO staining before and after 55 °C or 85 °C HS showed no differences in the proportion of SO-positive cells (Fig. 2c, d), indicating that both treatments caused irreversible rapid permeabilization of PM typical for necrosis [10, 11, 84].

    The protoplast shrinkage during HS-induced cell death is ATP- and Ca 2+ -independent

    Dismantling of the apoptotic cells is ATP-dependent [42, 85]. Yet, loss of the PM integrity would lead to rapid depletion of intracellular ATP, rendering all energy-dependent processes defunct. To examine whether the HS-induced cell death requires ATP, we first imaged mitochondria after 55 °C and 85 °C HS. Already at the earliest checked time points 4–10 min after HS, under both temperatures, mitochondrial dye MitoTracker localized to aberrant structures similar to those observed after treatment with mitochondrial uncoupler and ATP synthesis inhibitor, protonophore carbonyl cyanide m-chlorophenylhydrazone (CCCP) [86], indicating disruption of mitochondrial membrane potential (MMP Fig. 3a). Furthermore, intracellular ATP content dropped dramatically after both HS treatments (Fig. 3b), most probably due to dissipation of the MMP and leakage of cytoplasmic content through the permeabilized PM.

    Protoplast shrinkage at 55 °C HS is an ATP- and Ca 2+ -independent process. a Mitochondria in BY-2 cells stained with MitoTracker Red and imaged 10 min after 55 °C or 85 °C HS, or after treatment with 48 μM CCCP under normal temperature. Severely damaged mitochondria were observed upon all three treatments. b Loss of intracellular ATP content upon HS. Snap freeze-thaw treatment in liquid nitrogen (N2) and CCCP treatment were used as positive controls for completely disrupted and uncoupled mitochondria, respectively. The experiment was repeated twice, each time using four biological replicates per treatment. c MitoTracker Red staining of BY-2 cells exposed to 55 °C in the presence or absence of 15 μM Cyclosporin A (CsA) reveals that inhibition of MPTP opening does not rescue mitochondria from severe damage and loss of MMP caused by HS. d MitoTracker Red localization in the cells pre-treated with 10 mM EGTA prior to the HS reveals that chelation of extracellular Ca 2+ does not rescue mitochondrial phenotype. e, f Dynamics of cell death (% SO-positive cells e) and protoplast shrinkage (f) in cells with normal and uncoupled (48 μM CCCP treatment) mitochondria. g, h Pre-treatment with 10 mM EGTA before HS does not affect dynamics of cell death (% SO-positive cells g) and protoplast shrinkage (h). Experiments shown in e–h were repeated three times, with ≥ 184 cells per treatment and time point. Each microscopy experiment was performed at least twice. Staining for no HS and 55 °C treatments were performed according to protocols i and ii, respectively (see “Sytox Orange and FM4-64 staining”). Scale bars, 20 μm (a) or 50 μm (c, d). IQR, interquartile range. b, eh One-way ANOVA with Dunnet’s test *p < 0.05

    In addition to ATP depletion, PM permeabilization would also cause entry of Ca 2+ into the cells, potentially followed by its accumulation in mitochondria. MMP-driven accumulation of Ca 2+ can trigger mitochondrial permeability transition (MPT) due to the opening of a nonspecific pore (mitochondrial permeability transition pore, MPTP) [87], which will cause arrest of ATP synthesis and production of reactive oxygen species ultimately resulting in necrotic cell death [88]. Although HS-induced plant cell death was previously suggested to be a Ca 2+ -dependent process [51], those experiments lacked controls for mitochondrial phenotype, respiration or ATP production.

    To test whether MPT plays a role in the observed mitochondrial phenotype, experiments were performed in the presence of cyclosporin A (CsA), an inhibitor of MPTP opening, or the Ca 2+ chelator ethylene glycol-bis (2-aminoethylether)-N,N,N′,N′-tetraacetic acid (EGTA). Neither CsA nor EGTA could alleviate the mitochondrial phenotype in cells subjected to the HS (Fig. 3c, d), indicating that mitochondrial malfunction was caused by the direct loss of the mitochondrial membrane integrity during the HS, independently on PM permeabilization.

    Next, we examined the importance of intracellular ATP for the AVD-like protoplast shrinkage. We found that pre-treatment of the cell cultures with CCCP prior to HS had no effect on the protoplast shrinkage (Additional file 2: Figure S2), demonstrating that the protoplast shrinkage does not require ATP. Time-resolved quantitative analysis of cell death (frequency of SO-positive cells) and protoplast shrinkage upon HS of cells with normal or uncoupled mitochondria revealed that both parameters were independent of the mitochondrial bioenergetic function (Fig. 3e, f). Pre-treatment with CsA or EGTA prior to HS did not alleviate the cell death rate either (Fig. 3g, h Additional file 2: Figure S2).

    The discrepancies between our observations and the previous studies could be caused by technical issues, primarily precision of the temperature measurement during HS. To examine this possibility, we compared frequency and phenotype of cell death after treatment at 40, 45, 50 and 55 °C. Although cell viability was inversely proportional to the temperature, the morphology of dead cells in all cases was identical to that observed at 55 °C (Fig. 4). Pre-treatment of cells with CCCP confirmed that at any of the checked temperatures the rate of cell death did not depend on mitochondrial activity (Fig. 4).

    HS at the temperature range 40–55 °C induces ATP- independent cell death. a SO staining of BY-2 cells heat-shocked for 10 min at 40, 45, 50 or 55 °C and imaged after 6 or 24 h. b Quantification of cell death (% SO-positive cells) in the samples showed that pre-treatment with CCCP provided no protection against cell death and protoplast shrinkage at any of the tested HS temperatures. On the contrary, after prolonged exposure (24 h), CCCP appeared to decrease HS tolerance of plant cells. Staining was performed for no HS and HS treatments according to protocols i and ii, respectively (see “Sytox Orange and FM4-64 staining”). IQR, interquartile range. Experiments were repeated three times, with ≥ 134 cells per treatment and time point. The data was subjected to one-way ANOVA with Bonferroni correction. *p < 0.05, ns, non-significant. Scale bars, 20 μm

    A recent study [89] proposed that HS at 55 °C caused ferroptosis in A. thaliana root hair cells. Therefore, we examined whether the cell death we observed in BY-2 cells under the same stress conditions could be classified as a ferroptosis. For this, BY-2 cells were treated with a ferroptosis inhibitor, Ferrostatin-1 (Fer-1), prior to HS at 55 °C and cell death rate was measured during 24 h after the stress. Treatment with Fer-1 did not alleviate the cell death (Fig. 5a, b data is shown for the first 12 h after HS). Furthermore, two independent experiments replicating conditions that were reported to induce ferroptosis in Arabidopsis root hair cells [89] did not confirm such type of cell death (Fig. 5c). Taken together, our results reject the notion that HS-induced cell death is a programmed process. On the contrary, they demonstrate that HS triggers rapid destruction of cellular components and passive decay of plant cells resembling accidental necrosis.

    HS-induced cell death response is not ferroptosis. a SO staining of BY-2 cell cultures demonstrates that pre-treatment with 1 μM Fer-1 does not affect HS-induced cell death. Staining for no HS and 55 °C treatments was performed according to protocols i and iii, respectively (see “Sytox Orange and FM4-64 staining”). b Quantification of cell death frequency in the samples illustrated in a. The chart shows representative results of three independent experiments, each including ≥ 280 cells per treatment and time point. c Pre-treatment with Fer-1 does not alleviate the HS-induced death of Arabidopsis thaliana root hair cells. Arrows indicate SO-positive nuclei. Three independent experiments demonstrated the same results. No quantification was performed in these experiments, since all cells were SO-positive. DIC, differential interference contrast. Scale bars, 20 μm

    Necrotic deaths caused by 55 °C or 85 °C display different cell morphologies due to a fixating effect of higher temperature

    The lack of protoplast shrinkage during cell death induced by 85 °C was used to classify it as necrosis [61, 70]. However, both 55 °C and 85 °C HS trigger instant and irreversible permeabilization of the PM, MMP dissipation, and drop in intracellular ATP content, which are hallmarks of necrosis [84]. Plausibly, morphological differences between necrotic cell deaths triggered by 55 °C and 85 °C could be explained by rapid protein denaturation occurring at 85 °C that would crosslink cellular components. Such “fixation” would prevent protoplast shrinkage. Consistent with this suggestion, high-temperature cell fixation protocols have been used as an alternative to chemical fixation [90].

    To test whether 85 °C HS acts as a fixative, we induced protoplast retraction from the cell wall by exposing stressed cell cultures to hypertonic conditions at 500 mM D-mannitol (Fig. 6a). The high osmotic pressure of the medium would induce dehydration and protoplast shrinkage in the non-fixed cells. As expected, the living cells treated with D-mannitol underwent typical plasmolysis manifested by reversible protoplast detachment from the cell wall. Cells exposed to 55 °C displayed irreversible protoplast shrinkage phenotype both with and without D-mannitol treatment. However, although cells treated at 85 °C HS exhibited visible signs of dehydration in the hypertonic solution, protoplasts of virtually all cells remained attached to the cell wall. Quantitative analysis revealed no significant changes in the protoplast area of cells treated at 85 °C HS under normal or high osmotic pressure (Fig. 6b). These data provide compelling evidence that the fixing effect of 85 °C HS prevents protoplast shrinkage. Thus, distinct phenotypes of cell deaths induced by HS at 55 °C and 85 °C do not reflect differences in the cell death execution mechanism.

    85 °C HS prevents protoplast shrinkage in necrotic cells by fixing cellular content. a Cells stained with FDA were exposed to 55 °C or 85 °C and mounted in the normal growth medium or hypertonic medium supplemented with 500 mM D-mannitol. As expected, hypertonic medium induced plasmolysis in the non-stressed cells (No HS) and had no effect on protoplasts shrunken after 55 °C HS. Importantly, high osmotic pressure failed to induce protoplast shrinkage in cells exposed to 85 °C HS, thus confirming that cells were fixed by the high-temperature treatment. b The extent of protoplast shrinkage upon HS followed by increased osmotic pressure was quantified as the percentage of the cell area (outlined by the white dotted line in a) occupied by the protoplast area (outlined by the red dotted line in a). Student’s t test, n = 3 replicates (each replicate representing ≥ 34 cells), *p < 0.0001, ns, non-significant. IQR, interquartile range. Scale bars, 20 μm


    Methods

    Recombinant sRAGE protein and Rage −/− mice were used for this study.

    Identification of sRAGE by flow cytometry. Alexa Fluor 660 dye-labeled sRAGE or bovine serum albumin was added to dexamethasone-treated thymocytes. Apoptotic cells were identified using a MitoProbe JC-1 assay kit (Molecular Probes). Flow cytometry was performed on a BD FACSCanto II (BD Biosciences).

    PIP strip assay. PIP strips on which the indicated phospholipids had been spotted, were purchased from Echelon Bioscience, and dot-blot experiments were carried out according to the manufacturer's protocol.

    Surface-plasmon resonance analysis (Biacore). The binding affinity of sRAGE to phosphatidylserine was analysed using a Biacore X100 (Biacore AB) in a single-cycle affinity model. KD was calculated using ka and kd values by a trivalent analyte model.

    FRET analysis. FRET analysis was performed as described previously (Liu et al, 2008).

    Confocal laser scanning microscopy. Alveolar macrophages were grown on coverslips and then preincubated with 5 μM NBD-phosphatidylserine liposome for 2 h, after which mRAGE was detected by indirect immunofluorescence using a rabbit polyclonal RAGE antibody (Abcam) with Alexa Fluor 647-conjugated goat rabbit IgG (Molecular Probes).

    Phagocytosis of apoptotic thymocytes. Phagocytosis was assayed by adding apoptotic thymocytes (2.5 × 10 6 cells/ml) with or without sRAGE (3 μg/ml). Phagocytosis was determined as a phagocytic index under microscopy, as described previously (Morimoto et al, 2006). Each condition was tested in duplicate, and the samples were analysed blindly and independently by two researchers (M.H. and K.M.).

    Rac1 activity assays. Rac1 activity was measured using an ELISA-based Rac1 Activation Assay Biochem Kit (G-LISA, Cytoskeleton).

    LPS-induced lung injury. LPS from Escherichia coli serotype 055:B5 was obtained from Sigma-Aldrich, and the induction of lung injury was performed as described previously (Yamada et al, 2004).