<figure><img src="image_1.png" /></figure>

HAL
open science

# Anticancer chemotherapy and radiotherapy trigger both non-cell-autonomous and cell-autonomous death

Isabelle Martins, Syed Qasim Raza, Laurent Voisin, Haithem Dakhli, Awatef Allouch, Frédéric Law, Dora Sabino, Dorine de Jong, Maxime Thoreau, Elodie Mintet, et al.

<figure><img src="image_2.png" /></figure>

## To cite this version:

Isabelle Martins, Syed Qasim Raza, Laurent Voisin, Haithem Dakhli, Awatef Allouch, et al. Anti-cancer chemotherapy and radiotherapy trigger both non-cell-autonomous and cell-autonomous death. Cell Death and Disease, 2018, 9 (7), pp.716. 10.1038/s41419-018-0747-y. hal-02356534

HAL Id: hal-02356534

https://hal.science/hal-02356534

Submitted on 13 Nov 2019

HAL is a multi-disciplinary open access archive for the deposit and dissemination of scientific research documents, whether they are published or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.

<figure><img src="image_3.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
ARTICLE

Open Access

# Anticancer chemotherapy and radiotherapy trigger both non-cell-autonomous and cell-autonomous death

Isabelle Martins<fnref n="1" />,<fnref n="2" />,³,⁴, Syed Qasim Raza<fnref n="1" />,<fnref n="2" />,³,⁴,⁵, Laurent Voisin<fnref n="1" />,<fnref n="2" />,³,⁴, Haithem Dakhli<fnref n="1" />,<fnref n="2" />,³,⁴, Awatef Allouch<fnref n="1" />,<fnref n="2" />,³,⁴, Frédéric Law<fnref n="1" />,<fnref n="2" />,³,⁴, Dora Sabino⁶, Dorine De Jong<fnref n="1" />,<fnref n="2" />,³,⁴, Maxime Thoreau<fnref n="1" />,<fnref n="2" />,³,⁴, Elodie Mintet<fnref n="1" />,<fnref n="2" />,³,⁴, Delphine Dugué<fnref n="2" />,³,⁴, Mauro Piacentini⁷,⁸, Marie-Lise Gougeon⁹, Fanny Jaulin⁶, Pascale Bertrand¹⁰, Catherine Brenner¹¹, David M. Ojcius¹², Guido Kroemer¹³,¹⁴,¹⁵,¹⁶,¹⁷,¹⁸, Nazanine Modjtahedi<fnref n="2" />,³,⁴, Eric Deutsch<fnref n="2" />,³,⁴ and Jean-Luc Perfettini¹⁰,¹²,³,⁴

## Abstract

Even though cell death modalities elicited by anticancer chemotherapy and radiotherapy have been extensively studied, the ability of anticancer treatments to induce non-cell-autonomous death has never been investigated. By means of multispectral imaging flow-cytometry-based technology, we analyzed the lethal fate of cancer cells that were treated with conventional anticancer agents and co-cultured with untreated cells, observing that anticancer agents can simultaneously trigger cell-autonomous and non-cell-autonomous death in treated and untreated cells. After ionizing radiation, oxaliplatin, or cisplatin treatment, fractions of treated cancer cell populations were eliminated through cell-autonomous death mechanisms, while other fractions of the treated cancer cells engulfed and killed neighboring cells through non-cell-autonomous processes, including cellular cannibalism. Under conditions of treatment with paclitaxel, non-cell-autonomous and cell-autonomous death were both detected in the treated cell population, while untreated neighboring cells exhibited features of apoptotic demise. The transcriptional activity of p53 tumor-suppressor protein contributed to the execution of cell-autonomous death, yet failed to affect the non-cell-autonomous death by cannibalism for the majority of tested anticancer agents, indicating that the induction of non-cell-autonomous death can occur under conditions in which cell-autonomous death was impaired. Altogether, these results reveal that chemotherapy and radiotherapy can induce both non-cell-autonomous and cell-autonomous death of cancer cells, highlighting the heterogeneity of cell death responses to anticancer treatments and the unsuspected potential contribution of non-cell-autonomous death to the global effects of anticancer treatment.

## Introduction

From the initial discovery of programmed cell death during animal development<fnref n="1" /> to the recent identification of entotic death during embryo implantation<fnref n="2" />, a cornucopia

of cell death modalities has been identified and shown to play a role in numerous physiological or pathological situations³, ⁴. Mainly studied as clonal cellular responses to lethal stress, cell death processes have been defined on the basis of their specific morphological features (e.g., apoptotic, autophagic, or necrotic), their metabolic and biochemical characteristics (e.g., the loss of mitochondrial transmembrane potential, the exposure of phosphatidylserine (PS) on the outer leaflet side, or the rupture of plasma membrane integrity), their enzymatic and

Correspondence: J.-L. Perfettini (perfettini@orange.fr)

Full list of author information is available at the end of the article

These authors contributed equally: Isabelle Martins, Syed Qasim Raza.

Edited by G. Raschella

<figure><img src="image_4.png" /></figure>

<footnote n="1">¹Cell Death and Aging Team, Gustave Roussy Cancer Campus, 114 rue Edouard Vaillant, F-94805 Villejuif, France</footnote>

<footnote n="2">²Laboratory of Molecular Radiotherapy, INSERM U1030, Gustave Roussy Cancer Campus, 114 rue Edouard Vaillant, F-94805 Villejuif, France</footnote>
catabolic activities (involving (or not) caspases, receptor-
interacting protein kinases (RIPKs), mixed lineage kinase
domain-like proteins, or cathepsins), and in relation to
their ability to elicit an inflammatory reaction or to sti-
mulate an immune response. A classification of cell death
modalities built on these criteria has been proposed⁵ and
led to the ordering of lethal processes into three distinct
types: type I cell death (or apoptosis), type II cell death (or
autophagic cell death), and type III cell death (or necro-
sis). All these processes, which are executed in a cell-
autonomous manner, can be induced in the targeted
stressed cells or at a distance, in the neighboring cells
(through bystander effects). These processes are known as
cell-autonomous death (CAD)⁶. Despite major progresses
that have been made in the field, the relative contribution
of both direct and bystander-signal-mediated killing
triggered by typical CAD remains poorly explored.

Cell death subroutines (such as mitotic death and cor-
nification) that do not or partially exhibit the typical
morphological and biochemical hallmarks of cell death
have been less studied and are listed in a poorly defined
subgroup of cell death modalities known as atypical cell
death⁵. In recent years, additional cell death mechanisms
(such as entosis or emperitosis) have been described and
associated with this neglected subgroup of cell death
modalities⁷, ⁸. Their examination revealed the existence of
cell death processes that are elicited after the engulfment
of live cells by neighboring live cells. These lethal pro-
cesses are also known as non-cell-autonomous death
(NCAD). The first step of NCAD programs, which start
with the interaction of two cellular partners through
membrane adhesion receptors (such as E- or P-cadherins)
or stress receptors (such as lipoprotein receptor-related
protein), requires the formation of adherent junctions
between interacting cells and the activation of signaling
pathways, which may involve small GTPases (such as
Rho⁹ and cell division cycle 42 (CDC42)¹⁰) and ROCK
kinases⁷, on both interacting cells. The modulation of
actomyosin contractility and the reorganization of the
actin cytoskeleton in “target” cells also favor their invasion
into host cells⁹, ¹¹. This process is distinct from cellular
cannibalism, which can also trigger NCAD through the
activation of specific signaling pathways (such as
phagocytosis-related signaling pathways that involve
CDC42, chemokine (C-X-C motif) ligand 1 (CXCL1) or
CXCL6) on host cells and leads to the active engulfment
of target cells¹⁰. Independently of the precise cell
engulfment process, engulfed cells are targeted by “host”
lysosomal enzymes (such as cathepsins and granzymes)
and eliminated through distinct lethal mechanisms that
may involve major modulators of typical cell death (such
as cytochrome c, caspases, or autophagy-related (ATG)
proteins). When NCAD leads to the apoptotic demise of
engulfed cells, the process is called phagoptosis¹²⁻¹⁴ or

emperipolesis¹⁵. This form of cell death has been pro-
posed to occur frequently in the body¹² and to control
erythrocyte, neutrophil, platelet, and T cell homeostasis¹²,
¹⁶, ¹⁷. The engulfment of natural killer cells by tumor cells
was also shown to induce emperitosis, which is a pro-
grammed cell-in-cell death process that requires caspase-
3 activation and leads to DNA fragmentation⁸. Inversely,
entosis, another form of NCAD initially described after
homotypic interactions between breast cancer cells, is a
cell-in-cell invasion mechanism that does not require the
activation of caspases to eliminate engulfed cells⁷. We
recently defined NCAD as type IV cell death⁶.

Despite intensive biological and pharmaceutical
research programs that have helped to better characterize
cellular and biochemical processes associated with antic-
ancer treatment, the ability of anticancer agents to
simultaneously induce CAD and NCAD has never been
investigated. Here, using novel multispectral imaging
flow-cytometry-based methodology, we show that cancer
cells respond to treatment with various anticancer agents
by undergoing simultaneously several death modalities
that can be either cell-autonomous or non-cell-
autonomous and are distinctly impacted by the tumor-
suppressive factor p53, revealing unsuspected hetero-
geneity of cell death responses to anticancer treatment.

Results

Ionizing radiation (IR) induces CAD of irradiated cancer
cells

Even though radiotherapy is one of the most frequent
anticancer treatments used in the clinic, the lethal
mechanisms responsible for the therapeutic effects of
radiotherapy are still largely unknown. Lethal processes
(such as apoptosis and mitotic catastrophe) that have
been detected in response to IR were never directly
implicated in treatment efficiency¹⁸, suggesting that
additional, uncharacterized cell death modalities may
contribute to the therapeutic effects of radiotherapy. To
precisely analyze the cellular mechanisms through which
cancer cells may simultaneously undergo direct and
bystander cell killing in response to IR, we designed a
novel cell death profiling assay based on co-culture of
untreated cancer cells that have been labeled with the red
fluorescent probe 5-(and-6)-(((4-chloromethyl)benzoyl)
amino) tetramethylrhodamine (CMTMR) and irradiated
isogenic cancer cells that have been labeled with the green
fluorescent probe 5-chloromethylfluorescein diacetate
(CMFDA). After different times of co-culture and expo-
sure to different doses of IR, irradiated CMFDA⁺ cells,
non-irradiated CMTMR⁺ cells, and the total cell popu-
lation (CMFDA⁺ and CMTMR⁺) were analyzed for PS
exposure, loss of plasma membrane integrity, and DNA
content (Fig. 1a). The simultaneous detection of the
above-indicated parameters allowed us to detect, through
<figure><img src="image_5.png" /></figure>

the use of multispectral imaging flow cytometry, the execution of at least four types of cell death (including several lethal pathways, such as apoptosis, mitotic death, pyroptosis, autophagic cell death, necrosis, necroptosis, entosis, emperitosis, and cellular cannibalism) on target cells and on neighboring cells (Fig. 1b), thus discriminating CAD from NCAD and direct cell killing from bystander lethal effects.

Thus CMFDA-labeled HCT116 human colon cancer cells (wild-type (WT) HCT116 (HCT116<sup>WT</sup>) cells, isogenic control HCT116 (HCT116<sup>+/+</sup>) cells, and HCT116<sup>p53R248W/+</sup> cells harboring the dominant-negative mutant of p53, p53R248W (Fig. 2a–h), or MCF7 human breast cancer cells (Fig. 2i–n) were irradiated with different doses (0, 4, 8, and 16 Gy), mixed after 24 h with

CMTMR-labeled isogenic cells (1:1 ratio), and cultured for the indicated times. PS exposure, plasma membrane integrity, and DNA content of each population were then determined by means of AnnexinV-BV786 (AV), DRAQ7 (D7), and Hoechst 33342 staining, respectively. Although no significant increase of apoptotic and necrotic cell death was observed in the untreated control cell population (as revealed by the detection of AnnexinV<sup>+</sup>DRAQ7<sup>-</sup> (AV<sup>+</sup>D7<sup>-</sup>), AnnexinV<sup>-</sup>DRAQ7<sup>+</sup> (AV<sup>-</sup>D7<sup>+</sup>), and AnnexinV<sup>+</sup>DRAQ7<sup>+</sup> (AV<sup>+</sup>D7<sup>+</sup>) cells (Fig. 2a, c–h)), a significant increase of AV<sup>+</sup>D7<sup>-</sup> cells and AV<sup>+</sup>D7<sup>+</sup> cells was detected in the total population of HCT116<sup>WT</sup> or HCT116<sup>+/+</sup> cells (as revealed by CMFDA<sup>+</sup> and CMTMR<sup>+</sup> cells) (Fig. 2b, c, f, o), in neighboring HCT116<sup>WT</sup> or HCT116<sup>+/+</sup> cells (CMTMR<sup>+</sup> cells)
<figure><img src="image_6.png" /></figure>

(Fig. 2b, d, g, p), and in irradiated HCT116<sup>WT</sup> or HCT116<sup>+/+</sup> cells (CMFDA<sup>+</sup> cells) (Fig. 2b, e, h, q). These processes were observed in a dose-dependent manner after 24 h (Fig. 2c-e) and 48 h (Fig. 2f-h) of co-culture, demonstrating that our methodology allows us to score the death of both non-irradiated and irradiated cells. These results were confirmed after 12 h (Fig. 2i-k) and 24 h (Fig. 2l-n) of co-culture of control or irradiated CMFDA-labeled MCF7 cells with untreated CMTMR-

labeled MCF7 cells. To characterize molecular mechanisms involved in the execution of these cell death processes, co-cultures of HCT116<sup>WT</sup> cells were performed in the presence of cell death modulators (Supplementary Figures 1a-e). Thus the ROCK-1 inhibitor Y27632 was used as an inhibitor of live cell engulfment following procedures of previously published reports<sup>7</sup>. The peptide derivatives benzyloxycarbonyl-Val-Ala-Asp(OMe)-fluoromethylketone (Z-VAD-fmk) and YVAD-cmk were used
(see figure on previous page)

**Fig. 2** Detection of γ-irradiation-elicited CAD modalities by quantitative imaging flow cytometry. **a–h** Detection and quantification of plasma membrane integrity loss (with DRAQ7) and PS exposure (with BV786-streptavidin/Annexin V biotin) were achieved for total cell populations (CMTMR<sup>+</sup> or CMFDA<sup>+</sup> HCT116<sup>WT</sup> cells (**a, o**), HCT116<sup>+/+</sup> cells (**c, f**), and MCF7 cells (**i, l**)), for the untreated (red) CMTMR<sup>+</sup> cell populations (HCT116<sup>WT</sup> cells (**a, b, o–q**), HCT116<sup>+/+</sup> cells (**c, d, f, g**), and MCF7 cells (**i, j, l, m**)), for untreated (green) CMFDA<sup>+</sup> cell populations (HCT116<sup>WT</sup> cells (**a, o, q**), HCT116<sup>+/+</sup> cells (**c, e, f, h**), and MCF7 cells (**i, k, l, n**)), and for treated (green) CMFDA<sup>+</sup> cell populations (HCT116<sup>WT</sup> cells (**b, o, q**), HCT116<sup>+/+</sup> cells (**c, e, f, h**), and MCF7 cells (**i, k, l, n**). CAD modalities were detected after 12-h (**i–k**), 24-h (**a–e**), or 48-h (**f–h**) co-culture of untreated (red) CMTMR-labeled cells with untreated (green) CMFDA-labeled cells (**a, c, f, i, l**) or of untreated (red) CMTMR-labeled cells with (green) CMFDA-labeled cells that have been irradiated with the indicated doses of γ-ionizing radiation (**b, d, e, g, j, k, m, n**). **o–q** Co-cultures of HCT116<sup>WT</sup> cells were also performed during 24 h in the presence or absence of the indicated pharmacological death effector inhibitors. CAD modalities were determined as previously described for the untreated (red) CMTMR<sup>+</sup> HCT116<sup>WT</sup> cells, for untreated (green) CMFDA<sup>+</sup> HCT116<sup>WT</sup> cells, for treated (green) CMFDA<sup>+</sup> HCT116<sup>WT</sup> cells, and for total cell population (CMTMR<sup>+</sup> or CMFDA<sup>+</sup> HCT116<sup>WT</sup> cells). Representative dot plots (**a, b**) and quantitative data (**c–q**) are shown (means ± SEM, *n* = 3). For **c–h**, asterisk (*) is used for the comparison of "HCT116<sup>+/+</sup>+Irr. HCT116<sup>+/+</sup>" with "HCT116<sup>+/+</sup>+0 Gy HCT116<sup>+/+</sup>" for AV<sup>+</sup>D7<sup>−</sup> and ampersand (&) is used for the comparison of "HCT116<sup>+/+</sup>+Irr. HCT116<sup>+/+</sup>" with "HCT116<sup>+/+</sup>+0 Gy HCT116<sup>+/+</sup>" for D7<sup>+</sup>. For **i–n**, asterisk (*) is used for the comparison of "MCF7+Irr. MCF7<sup>+</sup>" with "MCF7+0 Gy MCF7<sup>+</sup>" for AV<sup>+</sup>D7<sup>−</sup> and ampersand (&) for the comparison of "MCF7+Irr. MCF7<sup>+</sup>" with "MCF7+0 Gy MCF7<sup>+</sup>" for D7<sup>+</sup>. For **o–q**, asterisk (*) is used for comparison of "HCT116<sup>WT</sup>+4 Gy control (Co.) HCT116<sup>WT</sup>" with "HCT116<sup>WT</sup>+control (Co.) HCT116<sup>WT</sup>" for AV<sup>+</sup>D7<sup>−</sup>, hash (#) for the comparison of inhibitor-treated cells with respective control cells for AV<sup>+</sup>D7<sup>−</sup> and dollar symbol ($) for the comparison of inhibitor-treated cells with respective control cells for D7<sup>+</sup>. *, #, &p < 0.05; **, ##, &&p < 0.01; ***, ###, $$$, &&&&p < 0.001; and ****, ####, $$$$, &&&&&p < 0.0001

as pan-caspase or caspase-1 inhibitors, respectively¹⁹, ²⁰, and the RIPK1 inhibitor necrostatin-1 (NEC1) was employed as necroptosis inhibitor²¹. Bafilomycin A1 (BafA1) and the cyclin-dependent kinase inhibitor roscovitine (Rosco) were used to analyze the contribution of autophagic flux and mitotic progression, respectively²²⁻²⁴. We observed that the pan-caspase inhibitor Z-VAD-fmk reduced the exposure of PS on the plasma membrane of the total cell population of HCT116<sup>WT</sup> cells (Fig. 2o), of irradiated CMFDA<sup>+</sup> HCT116<sup>WT</sup> cells (Fig. 2q), and to a lower extent, those of non-irradiated CMTMR<sup>+</sup> HCT116<sup>WT</sup> cells (Fig. 2p). These results confirm the notion that irradiated CMFDA<sup>+</sup> cells die efficiently in the event of caspase activation²⁵. However, co-culture experiments performed with MCF7 cells that are deficient for caspase-3 revealed that both total (Fig. 2i, l) and irradiated cell population (Fig. 2j, n) also exhibited a significant increase of AV<sup>+</sup>D7<sup>−</sup> and AV<sup>+</sup>D7<sup>+</sup> cells (as compared to control cells), thus indicating that irradiated cancer cells can also die through caspase-3-independent death. In addition, the impairment of autophagic flux with BafA1 increased the frequency of dying cells (AV<sup>+</sup>D7<sup>−</sup>, AV<sup>+</sup>D7<sup>−</sup>, and AV<sup>+</sup>D7<sup>+</sup> cells) in the total cell population (CMFDA<sup>+</sup> and CMTMR<sup>+</sup> cells) (Fig. 2o), in non-irradiated CMTMR<sup>+</sup> cells (Fig. 2p), and in irradiated CMFDA<sup>+</sup> cells (Fig. 2q), pointing to autophagy as a pro-survival mechanism that contributes to rescue both non-irradiated and irradiated cells from death. The simultaneous analysis of the progression of non-irradiated and irradiated HCT116<sup>WT</sup> cells (Supplementary Figures 2a, 2b, and 2o-2s), HCT116<sup>+/+</sup> cells (Supplementary Figures 2c-2h), or MCF7 cells (Supplementary Figures 2i-2n) through their cell cycle compartment distribution showed that cell death induction was associated with the accumulation of irradiated cell populations in G2/M and 4N phases (Supplementary Figures 2a-2q). No alteration of

the cell cycle was detected in the non-irradiated CMTMR<sup>+</sup> cell populations (Supplementary Figures 2b, 2d, 2g, 2j, 2m, and 2p), implying that the cell cycle alterations were only detected in irradiated cells (Supplementary Figures 2b, 2e, 2h, 2k, 2n, and 2q). These results were confirmed by classical flow-cytometric analysis (Supplementary Figures 2r and 2s). Altogether, these results demonstrate that IR eliminates cancer cells mainly through direct cell killing.

## Paclitaxel induces CAD of both treated and neighboring cells

To further evaluate whether chemotherapeutic agents may also eliminate cancer cells by eliciting distinct cell death modalities, we examined the cell death features triggered by paclitaxel (PCT) (a taxane currently used in breast cancer treatment) and by oxaliplatin (OXA) and cisplatin (CDDP), two platinum salts that are frequently used to treat colorectal and non-small cell lung cancers, respectively. Thus CMFDA-labeled HCT116<sup>+/+</sup> cells (Fig. 3a–f), HCT116<sup>WT</sup> (Fig. 3m–o), or MCF7 cells (Fig. 3g–l) were treated with the indicated concentrations of PCT (Fig. 3), OXA, or CDDP (Fig. 4) during 24 h, then mixed at a 1:1 ratio with CMTMR-labeled cells, and cultured for 12, 24, or 48 h in the absence or presence of cell death inhibitors. Multispectral imaging flow-cytometric analysis revealed that, after the treatment with PCT, both untreated CMTMR<sup>+</sup> cells (Fig. 3b, e, h, k) and PCT-treated CMFDA<sup>+</sup> cells (Fig. 3c, f, i, l) were eliminated in a dose-dependent manner, by apoptosis (as compared to control cells (Fig. 3a–l)). Accordingly, the pan-caspase inhibitor Z-VAD-fmk inhibited the exposure of PS on the outer plasma membrane leaflet of both untreated CMTMR<sup>+</sup> cells and PCT-treated CMFDA<sup>+</sup> cells (Fig. 3m–o). Co-cultures of untreated CMTMR-labeled MCF7 cells with CMFDA-labeled MCF7 cells that
<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>

<figure><img src="image_15.png" /></figure>

<figure><img src="image_16.png" /></figure>

<figure><img src="image_17.png" /></figure>

<figure><img src="image_18.png" /></figure>

<figure><img src="image_19.png" /></figure>

<figure><img src="image_20.png" /></figure>

<figure><img src="image_21.png" /><figcaption>**Fig. 3** (See legend on next page.)</figcaption></figure>
<figure><img src="image_22.png" /><figcaption>**Fig. 4** (See legend on next page.)</figcaption></figure>
(see figure on previous page)

**Fig. 4** Quantitative imaging flow-cytometric detection of CAD modalities induced by oxaliplatin or cisplatin. The cell death profiling was performed after co-cultures of untreated (red) CMTMR-labeled HCT116<sup>WT</sup>, HCT116<sup>+/+</sup>, or MCF7 cells with untreated (green) CMFDA-labeled cells or with oxaliplatin (OXA)-treated or cisplatin (CDDP)-treated (green) CMFDA-labeled HCT116<sup>WT</sup>, HCT116<sup>+/+</sup>, or MCF7 cells. Co-cultures have been performed during 12 h (**d–f**) and 24 h (**a–c**, **g–l**) with the indicated concentrations of OXA or CDDP in the presence or absence of death effector inhibitors. As previously described, cells were labeled for the simultaneous detection of type I, II, or III cell death. Means ± SEM are indicated (n = 3). For **a–c**, asterisk (*) is used for the comparison of "HCT116<sup>+/+</sup>+OXA- or CDDP-treated HCT116<sup>+/+</sup>" with "HCT116<sup>+/+</sup>+0 nM HCT116<sup>+/+</sup>" for AV<sup>+</sup>D7<sup>-</sup>, hash (&amp;) is used for the comparison of "HCT116<sup>+/+</sup>+OXA- or CDDP-treated HCT116<sup>+/+</sup>" with "HCT116<sup>+/+</sup>+0 nM HCT116<sup>+/+</sup>" for D7<sup>+</sup>. For **d–i**, asterisk (*) is used for the comparison of "MCF7+OXA- or CDDP-treated MCF7" with "MCF7+0 nM MCF7" cells for AV<sup>+</sup>D7<sup>-</sup> and ampersand (&amp;) is used for the comparison of "MCF7+OXA- or CDDP-treated MCF7" with "MCF7+0 nM MCF7" for D7<sup>+</sup>. For **j–l**, asterisk (*) is used for comparison of "HCT116<sup>WT</sup>+OXA- or CDDP-treated control (Co.) HCT116<sup>WT</sup>" with "HCT116<sup>WT</sup>+control (Co.) HCT116<sup>WT</sup>" for AV<sup>+</sup>D7<sup>-</sup>, ampersand (&amp;) for the comparison of "HCT116<sup>WT</sup>+OXA- or CDDP-treated control (Co.) HCT116<sup>WT</sup>" with "HCT116<sup>WT</sup>+control (Co.) HCT116<sup>WT</sup>" for D7<sup>+</sup>, hash (#) for the comparison of inhibitor-treated cells with respective control cells for AV<sup>+</sup>D7<sup>-</sup>, and dollar symbol ($) for the comparison of inhibitor-treated cells with respective control cells for D7<sup>+</sup>. *, #, §, &amp; p < 0.05; **, ##, §§, &&& p < 0.01; ***, ###, $$$, &&&& p < 0.001; and ****, ####, $$$$, &&&&& p < 0.0001

as cytochrome c release, activating cleavage of caspase-3 and nuclear degradation (Fig. 7a–e). We also determined the impact of the pan-caspase inhibitor Z-VAD-fmk and the caspase-1 inhibitor YVAD-cmk on these apoptotic features. Fluorescence microscopy revealed that Z-VAD-fmk (but not YVAD-cmk) impaired release of cytochrome c and activation of caspase-3 in engulfed cells (Fig. 7c, d) but only partially inhibited nuclear degradation with such cells (Fig. 7e). These results confirm those obtained using multispectral flow imaging and clearly demonstrate that the death of engulfed cells requires the activation of both caspase-dependent and independent mechanisms. In addition, the vast majority of cannibal cells did not expose PS and did not exhibit loss of the integrity of their plasma membrane after anticancer treatments (Supplementary Figures 5a-5d) underscoring the fact that, after IR-, PCT-, OXA-, or CDDP-mediated cell engulfment, the internalized cells succumb to death without compromising the viability of the engulfing cells. Altogether, these results show that IR and major chemotherapeutic agents (such as PCT, OXA, and CDDP) can simultaneously induce CAD and NCAD in both treated and untreated cells.

## The transcriptional activity of p53 tumor-suppressor protein distinctly impacts the execution of CAD and NCAD elicited by anticancer treatment

Considering that p53 inactivation is a major cellular factor involved in the resistance of tumor cells to anticancer treatments<sup>26</sup>, we analyzed the impact of inhibition of p53 transcriptional activity on the induction of CAD and NCAD. Thus we examined the influence of a dominant-negative mutation at codon 248 in one allele of p53 (p53R248W) on the execution of CAD and NCAD elicited by IR, PCT, CDDP, or OXA treatment. In parallel to previous experiments (Figs. 2c–h, 3a–f, 4a–c, 5b, c, and 6b, c, h, i), CMFDA-labeled HCT116<sup>p53R248W/+</sup> cells were irradiated with different doses (Fig. 8a, c, e, g) or treated with the indicated concentrations of PCT, OXA, or CDDP

(Fig. 8a–h) for 24 h, then mixed at a 1:1 ratio with CMTMR-labeled cells, and co-cultured for 24 h (Fig. 8a, c, e, g) or 48 h (Fig. 8b, d, f, h). As described above, PS exposure, plasma membrane integrity, and DNA content of each cellular partner were then analyzed using multispectral imaging flow cytometry. As expected, we observed a significant inhibition of PCT-, CDDP-, and OXA-elicited apoptosis of irradiated or treated HCT116<sup>p53R248W/+</sup> cells (Fig. 8a, e). In addition, p53 inactivation impaired PCT-induced apoptosis of untreated HCT116<sup>p53R248W/+</sup> cells (Fig. 8c), as compared to control irradiated or treated HCT116<sup>+/+</sup> cells (Figs. 2c–h, 3a–f, 4a–c, 5b, c, and 6b, c, h, i). Although, p53 inactivation had no impact on IR-, PCT-, and CDDP-mediated cellular cannibalism, OXA-induced cellular cannibalism was repressed when transcriptional activity of p53 was inhibited (Fig. 8g, h), as compared to control irradiated or treated HCT116<sup>+/+</sup> cells (Fig. 5d and 6b, c, h, i). p53 transcriptional inactivation similarly impacted the progression through the cell cycles of IR-, PCT-, OXA-, and CDDP-treated HCT116<sup>p53R248W/+</sup> cells (Supplementary Figure 6), as compared to control cells (Figs. 2c–h, 3a–f, and 4a–c). Altogether, these results demonstrated that the transcription factor p53 distinctly modulates the induction of NCAD and emphasizes the conclusion that the stimulation of cannibalistic activity of p53-mutated cancer cells by anticancer chemotherapies (such as PCT and CDDP) or radiotherapy should be considered as a treatment option.

## Discussion

Using a novel quantitative multispectral imaging flow-cytometry-based platform and co-culture experiments, we could provide evidence suggesting that IR and chemotherapeutic agents (such as PCT, OXA, and CDDP) can simultaneously trigger CAD and NCAD in both treated and untreated cancer cell populations. Thus, for the first time, we observed that, after IR, a fraction of irradiated cancer cells is eliminated through CAD, while
<figure><img src="image_23.png" /><figcaption>**Fig. 5** (See legend on next page.)</figcaption></figure>

another fraction of the irradiated cancer cell population
engulfed and killed neighboring cells via both caspase-
dependent and -independent death mechanisms. Simil-
arily, OXA and CDDP triggered killing of some treated
cancer cells through cell-autonomous apoptosis and

cellular cannibalism. Interestingly, PCT not only simul-
taneously induced CAD and NCAD in distinct fractions of
the treated cell population but also caused the apoptosis
of untreated cells co-cultured with PCT-treated cells.
Consistent with prior reports¹⁸, ²⁷, these results reveal the
<figure><img src="image_24.png" /><figcaption>**Fig. 6** (See legend on next page.)</figcaption></figure>
(see figure on previous page)

**Fig. 6** Quantitative imaging flow-cytometric detection of NCAD modalities triggered by paclitaxel, oxaliplatin, or cisplatin. Simultaneously to the detection of CAD in Figs. 3 and 4, NCAD mechanisms were determined with the indicated concentrations of PCT, OXA, and CDDP and in the presence or absence of the indicated inhibitors after co-cultures of untreated (red) CMTMR-labeled cells (HCT116<sup>WT</sup> cells (**a**, **f**, **g**, **l**, **m**), HCT116<sup>+/+</sup> cells (**b**, **c**), or MCF7 cells (**d**, **e**) with untreated (green) CMFDA-labeled isogenic cells and after co-cultures of untreated (red) CMTMR-labeled cells (HCT116<sup>WT</sup> cells (**a**, **f**, **g**, **l**, **m**), HCT116<sup>+/+</sup> cells (**b**, **c**), or MCF7 cells (**d**, **e**) with PCT-, OXA-, or CDDP-treated (green) CMFDA-labeled isogenic cells. The detection of cell-in-cell structures (**a**–**f**, **h**–**l**) and target cell degradation (**g**, **m**) were performed as indicated in the legend of Fig. 5. Frequencies are shown (means ± SEM, *n* = 3). For **b**, **c**, asterisk (*) is used for the comparison of “HCT116<sup>+/+</sup>+PCT-treated HCT116<sup>+/+</sup>” with “HCT116<sup>+/+</sup>+0 nM HCT116<sup>+/+</sup>” for G(R), ampersand (&) is used for the comparison of “HCT116<sup>+/+</sup>+PCT-treated HCT116<sup>+/+</sup>” with “HCT116<sup>+/+</sup>+0 nM HCT116<sup>+/+</sup>” for R (G). For **d**, **e**, asterisk (*) is used for comparison of “MCF7+PCT-treated MCF7” with “MCF7+0 nM MCF7” cells for R(G). For **f**, asterisk (*) is used for the comparison of “MCF7+PCT-treated MCF7” with “MCF7+0 nM MCF7” cells for R(G). For **h**, asterisk (*) is used for the comparison of “HCT116<sup>WT</sup>+PCT-treated HCT116<sup>WT</sup>” with “HCT116<sup>WT</sup>+control (Co.) HCT116<sup>WT</sup>” for G(R), ampersand (&) for the comparison of “HCT116<sup>WT</sup>+PCT-treated HCT116<sup>WT</sup>” with “HCT116<sup>WT</sup>+control (Co.) HCT116<sup>WT</sup>” for R(G), hash (#) for the comparison of inhibitor-treated cells with control cells for G(R), and dollar symbol ($) for the comparison of inhibitor-treated cells with respective control cells for R(G). For **g**, **m**, hash (#) for the comparison of inhibitor-treated cells with respective control cells for R(G). For **h**, **i**, asterisk (*) is used for the comparison of “HCT116<sup>+/+</sup>+OXA- or CDDP-treated HCT116<sup>+/+</sup>” with “HCT116<sup>+/+</sup>+0 nM HCT116<sup>+/+</sup>” cells for G(R), and ampersand (&) is used for the comparison of “HCT116<sup>+/+</sup>+OXA- or CDDP-treated HCT116<sup>+/+</sup>” with “HCT116<sup>+/+</sup>+0 nM HCT116<sup>+/+</sup>” for R(G). For **j**, **k**, asterisk (*) is used for the comparison of “MCF7+OXA- or CDDP-treated MCF7” with “MCF7+0 nM MCF7” for G(R) and ampersand (&) is used for the comparison of “MCF7+OXA- or CDDP-treated MCF7” with “MCF7+0 nM MCF7” for R(G). For **l**, asterisk (*) is used for the comparison of “HCT116<sup>WT</sup>+OXA- or CDDP-treated HCT116<sup>WT</sup>” cells with “HCT116<sup>WT</sup>+control (Co.) HCT116<sup>WT</sup>” cells for G(R), ampersand (&) for the comparison of “HCT116<sup>WT</sup>+OXA- or CDDP-treated HCT116<sup>WT</sup>” with “HCT116<sup>WT</sup>+control (Co.) HCT116<sup>WT</sup>” cells for R(G), hash (#) for the comparison of inhibitor-treated cells with control cells for G(R) and dollar symbol ($) for comparison of inhibitor-treated cells with respective control cells for R(G). *, #, $, &p < 0.05$; **, ##, $$, &&p < 0.01$; *** or &&&p < 0.001$; and ****, $^{####}$, &&&&&p < 0.0001$

characterization defining the impact of each cell death modality on the global cancer cell fate in vitro and on tumor growth in vivo should help to unravel the exact contribution of CAD and NCAD to anticancer treatment.

Taken together, our results demonstrate the coexistence of CAD (such as type I, II, and III cell death modalities) with NCAD (as the proposed type IV cell death modality)⁶. Future studies aiming at the pharmacological or genetic manipulation of cell death effectors (such as proapoptotic BCL-2 family members, BAX and BAK) should contribute to mitigate the notion of “the point of no return,” which has been proposed as the central cellular event for cell fate decision, considering lethal signaling pathways as a linear cascade of molecular events coming into play in a cell-autonomous fashion⁵. Indeed, future analyses must contemplate the heterogeneity of cellular responses to anticancer agents and IR and admit the possibility that non-cell autonomous and bystander effects might play a major role in determining the efficacy of cancer treatments.

# Material and methods

## Chemicals, cell lines, and culture conditions

Unless otherwise indicated, chemicals and acetyl-Tyr-Val-Ala-Asp-chloromethylketone (Ac-YVAD-cmk) were purchased from Sigma-Aldrich. Antibiotics, media, and supplements for cell culture were obtained from Life Technologies. Z-VAD-fmk was from Bachem and recombinant mouse tumor necrosis factor-alpha from R&D systems. Human colon carcinoma WT HCT116 (HCT116<sup>WT</sup>) cells, isogenic dominant-negative p53R248W-expressing HCT116 (HCT116<sup>P53R258W/+</sup>) cells, WT p53 control HCT116 (HCT116<sup>+/+</sup>) cells,

human MCF7 breast cancer cells, and murine fibro-sarcoma cell line L929 were cultured in Dulbecco's modified Eagle's medium. HCT116<sup>P53R258W/+</sup> cells and HCT116<sup>+/+</sup> cells are isogenic pairs of cells obtained from WT HCT116 (HCT116<sup>WT</sup>). Jurkat T cells were maintained in RPMI medium. All the media were supplemented with 10% heat-inactivated fetal bovine serum (FBS), 10 mM HEPES buffers, 2 mM L-glutamine, 10 U/mL penicillin sodium, and 10 µg/mL streptomycin sulfate.

## Irradiation and treatment with chemotherapeutic agents

Cells were seeded in 6-well plates, 12-well plates, or 25 cm² flasks for 24 h. Then adherent cells were treated with different concentrations of PCT, OXA, or CDDP or irradiated at the indicated dose with gamma-ray irradiator IBL-637 (Cs¹³⁷, 1 Gy/min, gamma CIS-BioInternational, IBA, Saclay, France) during 24 h.

## CellTracker™ fluorescent probes labeling and co-culture experiments

Upon the removal of the culture medium, cancer cells were incubated with prewarmed medium containing 10 µM of CMFDA (green fluorescence) or CMTMR (red fluorescence) (Molecular Probes-Life Technologies) for 45 min at 37 °C. Thereafter, cancer cells were rinsed twice with prewarmed medium and incubated for 45 min at 37 °C. Stained cells were then detached with Tryspin-EDTA solution (Life Technologies), suspended in complete medium, and cultured for the indicated times for cell death profiling or confocal microscopic analysis. Untreated cancer cells were labeled with CMFDA (green fluorescence, CMFDA<sup>+</sup>) or CMTMR (red fluorescence, CMTMR<sup>+</sup>) and treated cancer cells with CMFDA (green
<figure><img src="image_25.png" /><figcaption>**Fig. 7** Detection of biochemical features of engulfed cells detected after OXA, CDDP-PCT or IR treatment. **a, b** Representative fluorescent images of cytochrome c release, cleavage of CASP3, and nuclear degradation in engulfed HCT116<sup>WT</sup> cells detected after 24-h co-culture of untreated HCT116<sup>WT</sup> cells with untreated (red) CMTMR-labeled HCT116<sup>WT</sup> cells (**a**—**all**, **b**—**b**III) and after 24-h co-culture of untreated HCT116<sup>WT</sup> cells with (red) CMTMR-labeled HCT116<sup>WT</sup> cells that have been irradiated with 4 Gy of γ-ionizing radiation in the presence or absence of 100 µM Z-VAD-fmk (ZVAD) or 100 µM Y-VAD-cmk (YVAD) (**a**IV—**a**VI, **b**IV—**b**VI) are shown. Frequencies of cells showing cytochrome c release (**c**), cleaved CASP3 (**d**), and nuclear degradation (**e**) are shown (means ± SEM, n = 3). For **c**–**e**, asterisk (*) is used for "HCT116<sup>WT</sup>+OXA-treated HCT116<sup>WT</sup>", "HCT116<sup>WT</sup>+CDDP-treated HCT116<sup>WT</sup>", "HCT116<sup>WT</sup>+PCT-treated HCT116<sup>WT</sup>", and "HCT116<sup>WT</sup>+Irradiated (IR) HCT116<sup>WT</sup>" with "HCT116<sup>WT</sup>+control (Co.) HCT116<sup>WT</sup>" for nuclear degradation and hash (#) is used for the comparison of inhibitor-treated cells with respective control cells for cytochrome c release, cleaved caspase-3, or nuclear degradation. \* \#p < 0.05; \*\* \#\#p < 0.01; \*\*\* \#\#\#p < 0.001; and \#\#\#\#p < 0.0001</figcaption></figure>
<figure><img src="image_26.png" /><figcaption>Fig. 8 CAD and NCAD modalities elicited by chemotherapies and radiotherapy are distinctly regulated by p53 transcriptional activity.</figcaption></figure>

Quantitative imaging flow-cytometric detection of CAD and NCAD modalities was performed after 24 h (a, c, e, g) and 48 h (b, d, f, h) co-cultures of untreated (red) CMTMR-labeled HCT116<sup>p53R248W/+</sup> cells with untreated (green) CMFDA-labeled HCT116<sup>p53R248W/+</sup> cells, or with OXA-, CDDP-, PCT- or IR-treated (green) CMFDA-labeled HCT116<sup>p53R248W/+</sup> cells. Concentrations of OXA, CDDP, or PCT or doses of γ-irradiation used are indicated. The frequencies of AV<sup>+</sup>D7<sup>-</sup> cells, AV<sup>-</sup>D7<sup>+</sup> cells, and AV<sup>+</sup>D7<sup>+</sup> cells were determined as previously described on total cell population (as revealed by CMTMR<sup>+</sup> or CMFDA<sup>+</sup> HCT116<sup>p53R248W/+</sup> cells (a, b)), on untreated (red) CMTMR<sup>+</sup> HCT116<sup>p53R248W/+</sup> cells (c, d), on untreated (green) CMFDA<sup>+</sup> HCT116 cells (e, f), and on treated (green) CMFDA<sup>+</sup> HCT116<sup>p53R248W/+</sup> (e, f). Means ± SEM are indicated (n = 3). For a–f, asterisk (*) is used for the comparison of "HCT116<sup>p53R248W/+</sup>+OXA- or CDDP- or PCT- or IR-treated HCT116<sup>p53R248W/+</sup> with "HCT116<sup>+/+</sup>+OXA- or CDDP- or PCT- or IR-treated HCT116<sup>+/+</sup>" (from Figs. 2c–e, 3a–f, and 4a–c) for AV<sup>+</sup>D7<sup>-</sup>, and ampersand (&) is used for the comparison of "HCT116<sup>R248W/+</sup>+OXA- or CDDP- or PCT- or IR-treated HCT116<sup>+/+</sup>" (from Figs. 2c–e, 3a–f, and 4a–c) for D7<sup>+</sup>. For Fig. 7g, h, asterisk (*) is used for the comparison of "HCT116<sup>p53R248W/+</sup>+OXA- or CDDP- or PCT- or IR-treated HCT116<sup>p53R248W/+</sup> with "HCT116<sup>+/+</sup>+OXA- or CDDP- or PCT- or IR-treated HCT116<sup>+/+</sup>" (from Fig. 5b, c) for G(R), and ampersand (&) is used for comparison of "HCT116<sup>p53R248W/+</sup>+OXA- or CDDP- or PCT- or IR-treated HCT116<sup>p53R248W/+</sup> with "HCT116<sup>+/+</sup>+OXA- or CDDP- or PCT- or IR-treated HCT116<sup>+/+</sup>" (from Fig. 5b, c) for R(G). \* &amp;p < 0.05; \*\* &amp;&amp;p < 0.01; \*\*\* p < 0.001; and \*\*\*\* &amp;&amp;&amp;p < 0.0001

fluorescence, CMFDA<sup>+</sup>). The following cell mixtures (at a 1/1 ratio) were performed: untreated CMTMR<sup>+</sup> cancer cells were mixed with untreated CMFDA<sup>+</sup> cancer cells or

untreated CMTMR<sup>+</sup> cancer cells were mixed with treated CMFDA<sup>+</sup> cancer cells. Then cells were co-cultured during the indicated times in the presence or absence of the
pharmacological inhibitor of ROCK, Y27632 (30 µM), the pan-caspase inhibitor, Z-VAD-fmk (ZVAD, 100 µM), the inhibitor of caspase-1, Ac-YVAD-cmk (YVAD, 100 µM), the necroptosis inhibitor, NEC1 (30 µM), the inhibitor of the vacuolar type H(+)-ATPase (V-ATPase) inhibiting autophagy, BafA1 (50 nM), and the inhibitor of Cdks with an anti-mitotic activity, Rosco (10 µM).

## Cell death profiling by quantitative flow imaging

After the indicated times of co-cultures, both detached and adherent cells were collected and stained with Hoechst 33342 (10 µg/mL) during 1 h at 37 °C in warmed complete medium. To detect PS exposure and plasma membrane permeability, labeled HCT116 cells were successively incubated with Biotin-AnnexinV (BD Pharmingen) as recommended by the manufacturer, 0.5 µg BV786-Streptavidin (BD Biosciences), and 3 µM DRAQ7 (BioSstatus) during 15 min at 25 °C. After washing with phosphate-buffered saline (PBS) solution, samples were immediately analyzed using an imaging flow cytometer FlowSight® (Amnis®, part of EMD Millipore). Data were acquired at a 20× magnification, using the INSPIRE software. The 405, 488, and 561 nm lasers were used for excitation. Brightfield, Annexin V-BV786, DRAQ7, CMFDA, CMTMR, and Hoechst 33342 stainings were detected using, respectively, channels for 420–480, 745–800, 642–745, 480–560, 595–642, and 430–505 nm. At least 1000 events of cells per sample were analyzed. Additional single-labeled controls were prepared to normalize fluorescent signal across different channels. Acquired data were analyzed using the IDEAS analysis software (v6.1; Merck-Millipore). Gating strategy was the following. Cells were gated for focused cells using the Gradient RMS feature. Cells were gated for single cells using the aspect ratio and area features. For the cannibalism detection, cells were gated in the double positive CMFDA+ and CMTMR+ staining. The imaging flow cytometer FlowSight® provides the images of each events and thus allows to remove from the analysis the artifacts (such as cellular fragments or debris). We determined the sample size required for analyzing the main types of cell death elicited by anticancer treatments (such as apoptosis, necrosis, and cellular cannibal) by calculating the statistical power of our method (statistical power = 0.8; α = 0.05; sampling ratio = 1). We found that the number of cellular events that we need to acquire for obtaining statistically significant results is ranging between 300 and 900 cellular events. Accordingly, 1000 cellular events per condition were analyzed in Figs. 2a, b, o–q, 3m–o, 4j–l, 5d, e, and 6f, g, l, m and Supplementary Figures 1e, 2o-2q, 3m-3o, 4j-4l, and 5a-5d. To confirm these results, we then analyzed 10,000 cellular events per condition and validated previous results obtained with imaging flow cytometer as shown in Figs. 2c–n, 3a–l, 4a–i, 5b, c, k, l, 6b–e,

h–k, and 8a–h. These results were also confirmed using conventional methods such as classical flow cytometry and microscopy (as shown in Fig. 3d–i).

## Flow cytometry and confocal fluorescent microscopy

To detect PS exposure, plasma membrane permeability, and cell cycle progression, cells were after co-culture sequentially labeled with specific fluorescent probes (such as fluorescein isothiocyanate (FITC)-conjugated AnnexinV, propidium iodide (PI), and Hoechst 33342) and analyzed by flow cytometry. Both detached and adherent cells were collected and stained with Hoechst 33345 (10 µg/ml) during 1 h at 37 °C in warmed complete medium. After washing with PBS, HCT116 cells were suspended in 1× binding buffer supplemented with FITC-conjugated Annexin V (BD Biosciences) and PI (1 µg/mL) (Sigma), as per the manufacturer's instructions. Samples were then analyzed using LSRII flow cytometer (Becton Dickinson) and the FlowJo software v10. For confocal fluorescence microscopy, CMFDA- and CMTMR-labeled HCT116 cells were fixed after co-culture in 3.7% paraformaldehyde–PBS for 15 min. Cells were either counterstained with Hoechst 33342 (Invitrogen) and analyzed for the detection of cell-in-cell structures or permeabilized in 0.1% sodium dodecyl sulfate in PBS and incubated with FBS for 20 min, as previously described³⁹. Primary antibodies for detection of cytochrome c (#6H2. B4, Thermo Fisher Scientific) or cleaved caspase-3 (Asp175) (#9661, Cell Signaling Technology) were used in PBS containing 1 mg/ml bovine serum albumin and revealed with goat anti-rabbit immunoglobulin G (IgG) conjugated to Alexa 488 fluorochromes or with rabbit anti-mouse IgG conjugated to Alexa 488 fluorochromes from Invitrogen. Then cells were counterstained with Hoechst 33342 (Invitrogen) and analyzed by confocal SPE microscope equipped with Apochromat 63× 1.3 NA and 63× 1.15 NA oil immersion objectives. The Leica Application Suite version 2.6 software was used (Leica Microsystems).

## Western blots

Total cellular proteins were extracted in lysis buffer (containing 1% NP40, 20 mmol/L HEPES, 10 mmol/L KCl, 1 mmol/L EDTA, 10% glycerol, protease, and phosphatase inhibitor tablets). Protein extracts (30 µg) were run on 4–12% NuPAGE® Novex® Bis-Tris gels (Life Technologies) and transferred at 4 °C onto Immobilon polyvinylidfluoride membranes (Thermo Scientific). After blocking, membranes were incubated at 4 °C overnight with primary antibodies specific for: caspase-3 (#9662), cleaved caspase-3 (Asp175) (#9661), Myosin Light Chain 2 (MLC2) (#3672), phospho-MLC2 (Ser19) (#3675), LC3 A/B (#4108), and p-(S)-CDKs Substrate (#9477), obtained from Cell Signaling Technology. Antibodies against
GAPDH (#MAB374) were purchased from Millipore. Horseradish peroxidase-conjugated goat anti-mouse or anti-rabbit (Southern Biotechnology) antibodies were then incubated for 1 h and revealed with the SuperSignal West Pico® reagent (Thermo Fisher Scientific) or the ECL™ Prime Western Blotting Detection System (GE Healthcare) using an ImageQuant LAS 4000 software-assisted imager (GE Healthcare).

## Statistical analyses

Each experiment has been repeated at least three times, yielding comparable results. Means and standard errors (SEM) of three independent biological replicates are shown. Data were analyzed by means of Prism v. 5.03 (GraphPad Software, La Jolla, CA, USA). Statistical significance was assessed by one-way analysis of variance tests. In all experiments, *p* values <0.05 were considered as statistically significant.

## Acknowledgements

This work was supported by funds from La Ligue contre le Cancer Comité du Val-de-Marne, Agence Nationale de la Recherche (ANR), Institut National du Cancer (INCA-2015-1-PLBIO-07-IGR-1), Laboratory of Excellence (LabEx) LERMIT, the SIRIC Stratified Oncology Cell DNA Repair and Tumor Immune Elimination (SOCRATE), NATIXIS, Electricité de France and Fondation Gustave Roussy (to J-L.P.), and from the Ligue contre le Cancer Comité de Charente-Maritime (équipe labelisée), Agence National de la Recherche (ANR) - Projets blancs, ANR under the frame of E-Rare-2, the ERA-Net for Research on Rare Diseases, Association pour la Recherche sur le Cancer (ARC), Cancéropôle Ile-de-France; Chancelerie des universités de Paris (Legs Poix), Fondation pour la Recherche Médicale (FRM), a donation by Elior; the European Commission (ArtForce); the European Research Council (ERC), Fondation Carrefour, Institut National du Cancer (INCa), Inserm (HTE), Institut Universitaire de France, LeDucq Foundation, the LabEx Immuno-Oncology, the RHU Torino Lumière, the Searave Foundation, the SIRIC Stratified Oncology Cell DNA Repair and Tumor Immune Elimination (SOCRATE), the SIRIC Cancer Research and Personalized Medicine (CARPEM), and the Paris Alliance of Cancer Research Institutes (PACRI) (to G.K.). I.M. is supported by the SIRIC Stratified Oncology Cell DNA Repair and Tumor Immune Elimination (SOCRATE) and INCA (INCA-DGOS-INSERM 6043). S.Q.R is funded by Higher Education Commission (Pakistan) and by the Laboratory of Excellence LERMIT with a grant from ANR (ANR-10-LABX-33) under the program "Investissements d'Avenir" ANR-11-IDEX-0003-01. E.M. and M.T. are also supported by the Laboratory of Excellence LERMIT with a grant from ANR (ANR-10-LABX-33) under the program "Investissements d'Avenir" ANR-11-IDEX-0003-01. L.V. is funded by a grant from INCA (INCA-2015-1-PLBIO-07-IGR-1). H.D and D.D are, respectively, recipients of PhD fellowships from LabEx LERMIT and Fondation Philantropia. Q.W. is supported by China Scholarship Council. The authors would like to thank C. Leteur, Y. Lecluse, P. Rameau, and S. Salome-Desnoulez for their technical supports.

## Author details

¹Cell Death and Aging Team, Gustave Roussy Cancer Campus, 114 rue Edouard Vaillant, F-94805 Villejuif, France. ²Laboratory of Molecular Radiotherapy, INSERM U1030, Gustave Roussy Cancer Campus, 114 rue Edouard Vaillant, F-94805 Villejuif, France. ³Gustave Roussy Cancer Campus, 114 rue Edouard Vaillant, F-94805 Villejuif, France. ⁴Université Paris Saclay, 114 rue Edouard Vaillant, F-94805 Villejuif, France. ⁵Institute of Biochemistry and Biotechnology, University of Veterinary and Animal Sciences, UVAS Syed Abdul Qadir Jilani (Out Fall) Road, Lahore 54200, Pakistan. ⁶INSERM U981, Gustave Roussy Cancer Campus, 114 rue Edouard Vaillant, F-94805 Villejuif, France. ⁷National Institute for Infectious Diseases "Lazzaro Spallanzani", Via Portuense 292, 00149 Rome, Italy. ⁸Department of Biology, University of Rome "Tor Vergata", Via della Ricerca Scientifica 1, 00133 Rome, Italy. ⁹Antiviral Immunity, Biotherapy and Vaccine Unit, Institut Pasteur, 25 rue du Dr. Roux, F-75015 Paris, France. ¹⁰DNA Repair and Aging Team, Commissariat à l'Energie Atomique, Direction

Recherche Fondamentale, Institut de Biologie François Jacob, INSERM UMR 967 CEA, Université Paris Diderot, Université Paris Saclay, 18, route du Panorama, F-92265 Fontenay-aux-Roses, France. ¹¹Laboratory of Signalling and Cardiovascular Pathophysiology, INSERM UMR-S 1180, Université Paris-Sud, Université Paris Saclay, 5 rue Jean-Baptiste Clément, F-92296 Châtenay-Malabry, France. ¹²Department of Biomedical Sciences, University of the Pacific, School of Dentistry, 155 Fifth Street, San Francisco, CA 94103, USA. ¹³Equipe 11 Labellisée par la Ligue Contre le Cancer, Centre de Recherche des Cordeliers, Paris, France. ¹⁴Cell Biology and Metabolomics Platforms, Gustave Roussy Cancer Campus, Villejuif, France. ¹⁵Université Paris Descartes, Sorbonne Paris Cité, Paris, France. ¹⁶Université Pierre et Marie Curie, Paris, France. ¹⁷Pôle de Biologie, Hôpital Européen Georges Pompidou, AP-HP, Paris, France.

¹⁸Karolinska Institute, Department of Women's and Children's Health, Karolinska University Hospital, Stockholm, Sweden

## Conflict of interest

The authors declare that they have no conflict of interest.

## Publisher's note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Supplementary Information** accompanies this paper at (https://doi.org/10.1038/s41419-018-0747-y).

Received: 12 April 2018 Revised: 11 May 2018 Accepted: 24 May 2018

Published online: 18 June 2018

## References

1. Fuchs, Y. & Steller, H. Programmed cell death in animal development and disease. *Cell* **147**, 742–758 (2011).

2. Li, Y., Sun, X. & Dey, S. K. Entosis allows timely elimination of the luminal epithelial barrier for embryo implantation. *Cell Rep.* **11**, 358–365 (2015).

3. Lockshin, R. A. & Zakeri, Z. Cell death in health and disease. *J. Cell. Mol. Med.* **11**, 1214–1224 (2007).

4. Eroglu, M. & Derry, W. B. Your neighbours matter - non-autonomous control of apoptosis in development and disease. *Cell Death Differ.* **23**, 1110–1118 (2016).

5. Kroemer, G. et al. Classification of cell death: recommendations of the Nomenclature Committee on Cell Death 2009. *Cell Death Differ.* **16**, 3–11 (2009).

6. Martins, I. et al. Entosis: The emerging face of non-cell-autonomous type IV programmed death. *Biomed. J.* **40**, 133–140 (2017).

7. Overholtzer, M. et al. A nonapoptotic cell death process, entosis, that occurs by cell-in-cell invasion. *Cell* **131**, 966–979 (2007).

8. Wang, S. et al. Internalization of NK cells into tumor cells requires ezrin and leads to programmed cell-in-cell death. *Cell Res.* **19**, 1350–1362 (2009).

9. Sun, Q. et al. Competition between human cells by entosis. *Cell Res.* **24**, 1299–1310 (2014).

10. Cano, C. E. et al. Homotypic cell cannibalism, a cell-death process regulated by the nuclear protein 1, opposes to metastasis in pancreatic cancer. *EMBO Mol. Med.* **4**, 964–979 (2012).

11. Sun, Q., Cibas, E. S., Huang, H., Hodgson, L. & Overholtzer, M. Induction of entosis by epithelial cadherin expression. *Cell Res.* **24**, 1288–1298 (2014).

12. Brown, G. C. & Neher, J. J. Eaten alive! Cell death by primary phagocytosis: 'phagoptosis'. *Trends Biochem. Sci.* **37**, 325–332 (2012).

13. Brown, G. C. & Neher, J. J. Microglial phagocytosis of live neurons. *Nat. Rev. Neurosci.* **15**, 209–216 (2014).

14. Brown, G. C., Vilalta, A. & Fricker, M. Phagoptosis - cell death by phagocytosis - plays central roles in physiology, host defense and pathology. *Curr. Mol. Med.* **15**, 842–851 (2015).

15. Sierro, F. et al. Suicidal emperipolesis: a process leading to cell-in-cell structures, T cell clearance and immune homeostasis. *Curr. Mol. Med.* **15**, 819–827 (2015).

16. Khandelwal, S., van Rooijen, N. & Saxena, R. K. Reduced expression of CD47 during murine red blood cell (RBC) senescence and its role in RBC clearance from the circulation. *Transfusion* **47**, 1725–1732 (2007).
17. Lagasse, E. & Weissman, I. L. bcl-2 inhibits apoptosis of neutrophils but not their engulfment by macrophages. *J. Exp. Med.* **179**, 1047–1052 (1994).

18. Abend, M. Reasons to reconsider the significance of apoptosis for cancer therapy. *Int. J. Radiat. Biol.* **79**, 927–941 (2003).

19. Slee, E. A., Zhu, H., Chow, S. C., MacFarlane, M., Nicholson, D. W. & Cohen, G. M. Benzyloxycarbonyl-Val-Ala-Asp (OMe) fluoromethylketone (Z-VAD.FMK) inhibits apoptosis by blocking the processing of CPP32. *Biochem. J.* **315**(Pt 1), 21–24 (1996).

20. Garcia-Calvo, M. et al. Inhibition of human caspases by peptide-based and macromolecular inhibitors. *J. Biol. Chem.* **273**, 32608–32613 (1998).

21. Degterev, A., Maki, J. L. & Yuan, J. Activity and specificity of necrostatin-1, small-molecule inhibitor of RIP1 kinase. *Cell Death Differ.* **20**, 366 (2013).

22. Yamamoto, A. et al. Bafilomycin A1 prevents maturation of autophagic vacuoles by inhibiting fusion between autophagosomes and lysosomes in rat hepatoma cell line, H-4-II-E cells. *Cell Struct. Funct.* **23**, 33–42 (1998).

23. Castedo, M., Perfettini, J. L., Roumier, T. & Kroemer, G. Cyclin-dependent kinase-1: linking apoptosis to cell cycle and mitotic catastrophe. *Cell Death Differ.* **9**, 1287–1293 (2002).

24. Cicenas, J. et al. Roscovitine in cancer and other diseases. *Ann. Transl. Med.* **3**, 135 (2015).

25. Lowe, S. W., Schmitt, E. M., Smith, S. W., Osborne, B. A. & Jacks, T. p53 is required for radiation-induced apoptosis in mouse thymocytes. *Nature* **362**, 847–849 (1993).

26. Hanahan, D. & Weinberg, R. A. Hallmarks of cancer: the next generation. *Cell* **144**, 646–674 (2011).

27. Stevens, J. B. et al. Heterogeneity of cell death. *Cytogenet. Genome Res.* **139**, 164–173 (2013).

28. Garg, A. D. et al. Molecular and translational classifications of DAMPs in immunogenic cell death. *Front Immunol.* **6**, 588 (2015).

29. Hoeppner, D. J., Hengartner, M. O. & Schnabel, R. Engulfment genes cooperate with ced-3 to promote cell death in *Caenorhabditis elegans*. *Nature* **412**, 202–206 (2001).

30. Reddien, P. W., Cameron, S. & Horvitz, H. R. Phagocytosis promotes programmed cell death in *C. elegans*. *Nature* **412**, 198–202 (2001).

31. Abdu, Y., Maniscalco, C., Heddleston, J. M., Chew, T. L. & Nance, J. Developmentally programmed germ cell remodelling by endodermal cell cannibalism. *Nat. Cell Biol.* **18**, 1302–1310 (2016).

32. Chautan, M., Chazal, G., Cecconi, F., Gruss, P. & Golstein, P. Interdigital cell death can occur through a necrotic and caspase-independent pathway. *Curr. Biol.* **9**, 967–970 (1999).

33. Lindsten, T. & Thompson, C. B. Cell death in the absence of Bax and Bak. *Cell Death Differ.* **13**, 1272–1276 (2006).

34. Denecker, G. et al. Caspase-14 protects against epidermal UVB photodamage and water loss. *Nat. Cell Biol.* **9**, 666–674 (2007).

35. Scheller, C. et al. Caspase inhibition in apoptotic T cells triggers necrotic cell death depending on the cell type and the proapoptotic stimulus. *J. Cell. Biochem.* **97**, 1350–1361 (2006).

36. Vercammen, D. et al. Inhibition of caspases increases the sensitivity of L929 cells to necrosis mediated by tumor necrosis factor. *J. Exp. Med.* **187**, 1477–1485 (1998).

37. Vercammen, D. et al. Dual signaling of the Fas receptor: initiation of both apoptotic and necrotic cell death pathways. *J. Exp. Med.* **188**, 919–930 (1998).

38. Vercammen, D., Vandenabeele, P., Declercq, W., Van de Craen, M., Grooten, J. & Fiers, W. Cytotoxicity in L929 murine fibrosarcoma cells after triggering of transfected human p75 tumour necrosis factor (TNF) receptor is mediated by endogenous murine TNF. *Cytokine* **7**, 463–470 (1995).

39. Seror, C. et al. Extracellular ATP acts on P2Y2 purinergic receptors to facilitate HIV-1 infection. *J. Exp. Med.* **208**, 1823–1834 (2011).