Wun2-mediated integrin recycling promotes apoptotic cell clearance in Drosophila melanogaster

Gao, Ning; Zheng, Qian; Wang, Yanzhe; Li, Xiaowen; Li, Zhi; Xiao, Hui

doi:10.1038/s41418-022-01039-3

Article
Published: 15 July 2022

Wun2-mediated integrin recycling promotes apoptotic cell clearance in Drosophila melanogaster

Ning Gao^1,2^na1,
Qian Zheng¹^na1,
Yanzhe Wang¹^na1,
Xiaowen Li¹,
Zhi Li¹ &
…
Hui Xiao ORCID: orcid.org/0000-0001-9305-4486¹

Cell Death & Differentiation volume 29, pages 2545–2561 (2022)Cite this article

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Abstract

Apoptotic cell (AC) clearance is a complex process in which phagocytes recognize, engulf, and digest ACs during organismal development and tissue homeostasis. Impaired efferocytosis results in developmental defects and autoimmune diseases. In the current study, we performed RNA-sequencing to systematically identify regulators involved in the phagocytosis of ACs by Drosophila melanogaster macrophage-like S2 cells, followed by targeted RNA interference screening. Wunen2 (Wun2), a homolog of mammalian lipid phosphate phosphatase (LPP), was deemed as required for efferocytosis both in vitro and in vivo. However, efferocytosis was independent of Wun2 phosphatase activity. Proteomic analysis further revealed that Rab11 and its effector Rip11 are interaction partners of Wun2. Therefore, Wun2 collaborates with Rip11 and Rab11 to mediate efficient recycling of the phagocytic receptor βν integrin subunit to the plasma membrane. The loss of Wun2 results in the routing of βv integrin subunit (Itgbn) into lysosomes, leading to its degradation. The deficiency of βv integrin subunit on the cell surface leads to aberrant and disorganized actin cytoskeleton, thereby influencing the formation of macrophage pseudopodia toward ACs and thus failure to engulf them. The findings of this study provide insights that clarify how phagocytes coordinate AC signals and adopt a precise mechanism for the maintenance of engulfment receptors at their cell membrane surface to regulate efferocytosis.

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Introduction

Apoptosis is a crucial type of programmed cell death controlled and executed via an evolutionarily conserved pathway. At the final stage of apoptosis, cells are swiftly removed by phagocytes [1, 2]. The clearance of dead cells, termed ‘efferocytosis,’ is critical for homeostasis, and failure to properly clear ACs is linked to developmental abnormalities, neurodegenerative diseases, chronic inflammation, and autoimmune diseases [3,4,5,6].

Over the past decades, the use of Drosophila species in genetics has continually provided information on vertebrates, facilitating the development of important therapeutic targets for defective cell clearance in organisms [7]. Several key proteins required for ACs clearance have been identified in D. melanogaster; for example, Draper (Drpr) [8, 9], Croquemort (Crq) [10, 11], and αPS3/βν integrin [12, 13]. However, the detailed molecular mechanisms and signals associated with efferocytosis have not been elucidated.

Wunen (wun) and Wunen2 (wun2) encode the D. melanogaster homologs of lipid phosphate phosphatases 1, 2, and 3, all comprising six transmembrane α-helices with the C- and N-termini facing the cytoplasmic side and the catalytic site facing the outside of cells or the luminal side of organellar membranes [14]. LPPs can dephosphorylate several phosphate esters, including sphingosine-1-phosphate (S1P), ceramide-1-phosphate (C1P), phosphatidic acid (PA), and lysophosphatidic acid (LPA) [15, 16]. Although Wun and Wun2 have redundant functions in somatic cells to direct primordial germ cells or pole cell migration and survival [15, 17, 18], their localization and function in the trachea are distinct [19]. Low expression of LPP3 in triple-negative breast cancer can hinder the recruitment of immune cells, resulting in the blockage of cancer cell clearance [20]. However, studies on LPPs in immunology are limited, and the possible association of LPPs with efferocytosis remains poorly understood.

Integrins are conserved transmembrane receptors consisting of α and β heterodimers [21]. These proteins connect the extracellular matrix to the cytoskeleton, allowing phagocytic cells to adjust membrane protrusions according to the shape and size of the ACs or microorganisms to be engulfed [22]. The integrin αPS3/βν heterodimer is involved in the phagocytosis of ACs and Staphylococcus aureus in D. melanogaster phagocytes [13]. During phagocytosis, cell surface integrins are endocytosed and undergo endosomal sorting, a process in which Rab family proteins and their effectors are involved [23,24,25,26]. Although the mechanisms for integrin recycling are well-established, those underlying its degradation are not well understood.

In the present study, we screened D. melanogaster macrophage-like lineage S2 cells for novel gene candidates required for the phagocytosis of ACs using whole transcriptome analysis (RNA-sequencing). The LPP Wun2 was identified, and the non-phosphatase domain of Wun2 could interact with the Rab11 effector Rip11 C2 domain and βν integrin. This interaction prevented the lysosomal degradation and transport of βν integrin from Rab11 recycling endosomes to the plasma membrane to promote ACs clearance. Our study reveals the critical role of Wun2 in the Rab11 recycling pathway to prevent βν integrin degradation and identifies the important function of Wun2 in efferocytosis, along with implications for the treatment of diseases that result from impaired efferocytosis.

Results

Screening of candidate genes required for efferocytosis in D. melanogaster macrophages

Crq is defined as an engulfment receptor; its expression can be regulated by the extent of cell apoptosis [11]. During the process of ACs clearance in S2 cells, the expression of crq was time-dependent (Fig. 1A) and dose-dependent (Fig. 1B). Therefore, we performed transcriptomic analysis of the S2 cells engulfing a single or double dose of ACs for 6 h; 652 genes were upregulated in a dose-dependent manner (Fig. 1C). Thirty-one of these genes yielded fragments per kilobase of transcript per million mapped reads (FPKM) values in the double dose incubation that were 1.5-fold higher than those in the single dose incubation (Fig. 1D and Supplementary Table 1). Individually silencing 17 of these genes resulted in an obvious reduction of efferocytosis efficiency in S2 cells (Fig. 1E); wun2 RNA interference (RNAi) cells exhibited the lowest phagocytosis rate; therefore, wun2 was selected for further analysis.

**Fig. 1: RNA-sequencing and RNAi screening of candidate genes involved in efferocytosis.**

wun2 is required for efferocytosis in S2 cells and D. melanogaster macrophages

In S2 cells, ACs incubation elevated the transcriptional level of wun2 for 8 h (Fig. 2A). However, wun2 RNAi led to a significant decrease in the phagocytosis ratio (Fig. 2B, C and Supplementary Fig. 1A). For the in vivo assays, we generated an eight-base pair-deleted wun2^KO mutant fly via CRISPR/Cas9; this frameshift mutation led to the early termination of Wun2 translation (Fig. 2D and Supplementary Fig. 2A), and both PCR and immunofluorescence staining confirmed wun2^KO as a null allele mutant (Supplementary Fig. 2B–D). The stage 13 wun2^KO embryo macrophages failed to engulf ACs efficiently (Fig. 2E and Supplementary Fig. 1B); the phagocytosis ratio (number of engulfed ACs compared to total ACs) decreased by 17.69% (Fig. 2F) and the average phagocytic index (average number of ACs per macrophage, PI) scores were lower than that in the wild-type (WT) embryo (Fig. 2G). The difference was observed in the crq-GAL4, UAS-GFP marker models; WT embryos showed ACs mostly inside macrophages, while in wun2^KO embryos, most ACs were outside GFP-positive macrophages (Supplementary Fig. 1B–F). The hemocyte-specific knockdown of wun2 using crq-GAL4 led to a phenotype similar to that of the wun2^KO mutant (Fig. 2E–G). Mutant macrophages that re-expressed wun2 driven by crq-GAL4 or hml-GAL4 appeared to be fully rescued as most contained numerous ACs (Fig. 2E–G and Supplementary Fig. 3C–E). Hemolectin (Hml) is specifically expressed in embryonic and larval hemocytes, especially in plasmatocytes and crystal cells [27]. We found that hml-GAL4 could drive UAS-LifeAct-GFP expression from stage 12 embryos to larvae (Supplementary Fig. 3A, B). In addition, the wun2 mammalian homolog LPP3 rescued the defective phenotypes in wun2^KO (Fig. 2E–G and Supplementary Fig. 3C–E), indicating their functional conservation. Hence, wun2 appears to be required for efferocytosis in D. melanogaster.

**Fig. 2: Macrophages of the *wun2* mutant show defective phagocytosis of ACs.**

In D. melanogaster, hemocytes originate from the mesoderm of the head and develop into macrophages to disperse throughout the embryo; Even with severe manipulations of macrophage function can have relatively mild effects on its dispersal [28]. In the wun2^KO mutant, macrophages developed normally and migrated throughout the embryos (Supplementary Fig. 4A, B and Videos 1 and 2). No differences were found in the number of macrophages at stage 13 between WT and wun2^KO mutant flies (Supplementary Fig. 4C). The first wave of cell apoptosis began at stage 11 of embryogenesis and lasted until stage 16. There was no obvious difference in the number of ACs at stage 11 but AC counts were significantly higher in stages 14–15 of wun2^KO embryos than in those of WT embryos (Supplementary Fig. 4A). And re-expression of wun2 or lpp3 in wun2^KO could rescue the AC increased phenotype (Supplementary Fig. 4D). Therefore, we inferred that the higher ACs observed at stages 14-15 was caused by insufficient ACs clearance.

wun is not required for efferocytosis

Previously, wun and wun2 served as redundant genes to guide primordial germ cells [18, 29]. However, wun was excluded from the conditional RNA-sequencing data. To verify whether wun might nonetheless affect ACs clearance, we analyzed the location of Wun and Wun2 in S2 cells. They showed markedly different subcellular distribution in S2 cells (Fig. 3A, B). In contrast with wun2, wun knockdown did not affect ACs clearance in S2 cells (Fig. 3C, D). Furthermore, wun RNAi-treated embryos showed a level of phagocytosis that was almost equal to that of the WT embryos (Fig. 3E–G). In addition, double RNAi of wun and wun2 did not exacerbated ACs clearance defects (Fig. 3E–G), suggesting the likelihood of their functional divergence in macrophages.

**Fig. 3: *wun* does not affect ACs clearance for macrophages.**

Wun2 is not dependent on phosphatase activity for efferocytosis

Previous structural characterization revealed that the Wun2 catalytic site is located at the C-terminus of the acidPPc domain [14]. As the conserved domains are important for Wun2 catalytic activity in germ cell migration and maintenance [18], catalytic residues H274K, H326K, or double mutants (DM) were generated; Y225W was mutated as a control (Fig. 4A) [30, 31]. Subcellular localization revealed that Wun2-GFP fusion proteins typically appeared as punctate structures in the internal membrane, and to a less extent in the plasma membrane (Supplementary Fig. 5A). None of these mutations affected Wun2 localization in S2 cells (Supplementary Fig. 5A). Wun2 and Wun2 (DM) formed a ring-like structure around engulfed ACs (Fig. 4B), suggesting their potential role in the endosomal system. We purified Wun2 and its mutant proteins from transfected S2 cells (Supplementary Fig. 5B). Biochemical assays of Wun2 and its mutant proteins on phosphate esters indicated that the WT and control mutant Wun2 (Y225W) could dephosphorylate PA, C1P, LPA, and S1P in vitro (Supplementary Fig. 5C). In contrast, H274K and H326K mutants exhibited varying degrees of inhibited catalytic activity; Wun2 (DM) exhibited no phosphatase activity regardless of the substrate (Supplementary Fig. 5C), which is consistent with a previous study [30]. Additionally, re-expression of wun2 (DM) in the wun2^KO mutant effectively rescued the ACs clearance defect (Fig. 4C–E), which indicated the participation of the catalytic site-null Wun2 in efferocytosis.

**Fig. 4: Wun2 catalytic activity is not required for efferocytosis.**

Wun2 interacts with Rab11 in regulating efferocytosis

To identify the potential role of Wun2 in efferocytosis, a Flag-tagged-Wun2 stable S2 cell line was established. The Wun2-Flag protein was enriched by immunoprecipitation (Supplementary Fig. 6A), followed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) (Supplementary Fig. 6B). Two-hundred ten proteins were identified as putative interaction partners of Wun2 (Supplementary Table 2). Based on the properties and distribution of Wun2 in cells, we selected a subset of endosome system proteins as strong potential interactors. RNAi suppression of Rab11 led to significantly reduced efferocytosis efficiency (Supplementary Fig. 6C). Endosomal recycling routes have been classified in the literature as the ‘fast recycling pathway,’ regulated by Rab4, and the ‘slow recycling pathway,’ regulated by Rab11 [32]. Co-expression of Wun2 with Rab4 or Rab11, indicated that Wun2 was generally localized in the ‘slow recycling pathway’ (Supplementary Fig. 6D, F), and subsequent co-immunoprecipitation (co-IP) indicated that Wun2 interacted with the small GTPase Rab11 (Fig. 5A). Rab proteins cycle between the GTP-loaded state (active) and GDP-loaded state (inactive) [33]. Co-IP results revealed that Wun2 formed a complex with the active form of Rab11(Q70L) (Fig. 5B) but not with the dominant-negative Rab11(S25N) (Fig. 5C). This was further confirmed by the co-localization analysis of Wun2 with Rab11 (WT, Q70L, or S25N) in HeLa cells (Fig. 5D, E) and S2 cells (Supplementary Fig. 6E, F).

**Fig. 5: Wun2 interacts with active Rab11 to regulate ACs clearance.**

To address whether physical binding between Rab11 and Wun2 resulted in physiological effects in vivo, we characterized the efferocytosis phenotype of rab11^j2D1 and found that it failed to efficiently engulf ACs, with a lower phagocytosis ratio (Fig. 5F, G) and PI (Fig. 5F, H). In contrast, single heterozygous wun2^+/− or rab11^j2D1+/− mutant embryos were not distinguishable from the WT embryos; only embryos carrying the wun2 and rab11 double heterozygous combination exhibited defects in ACs clearance (Fig. 5F–H). These results suggested a genetic interaction between wun2 and rab11 to promote efferocytosis.

Wun2 directly interacts with Rip11 and βv integrin to regulate efferocytosis

The immunoprecipitation assay of Wun2 and Rab11 showed that the interaction between these proteins was not very strong (Fig. 5A, B). The yeast two-hybrid assay performed to investigate whether Wun2 and Rab11 interacted with each other directly further revealed there was no interaction between these proteins, regardless of the forms of Rab11: WT, negative, or active (Fig. 6A). Hence, we hypothesized an indirect interaction between Wun2 and Rab11, likely through complex formation with other proteins. As the active Rab11 is reportedly capable of binding diverse effector proteins to regulate intracellular membrane traffic [34], we examined the possibility of a direct interaction between Wun2 and Rab11 effectors. Yeast two-hybrid analysis showed that Wun2 could directly interact with the Rab11 effector Rip11 (Fig. 6A). Co-localization and co-IP analyses revealed a strong interaction between Wun2 and Rip11 (Fig. 6B–E). Furthermore, the catalytic site-null Wun2 mutant did not affect the interaction with Rip11 (Supplementary Fig. 7A). To narrow down the potential regions required for Wun2 and Rip11 interaction, we generated a series of systematic truncations of Wun2 (Wun2-1, 2, 3, 4, and 5) and Rip11 (Rip11-1, 2, 3, 4, 5, and 6) (Supplementary Fig. 7B) and found that the binding of Wun2-1 to the C2 domain of Rip11 (Rip11-5) was essential (Supplementary Fig. 7C). However, replacement of the Wun2-1 residues 155–177 with Wun2 C-terminal 345–350 residues, i.e., Wun2-1 (RC), resulted in the complete loss of interaction with Rip11 (Supplementary Fig. 7D), suggesting that residues 155–177 of Wun2 are essential for its binding to the C2 domain of Rip11.

**Fig. 6: Wun2 directly interacts with Rip11 and βv integrin subunit to regulate ACs clearance.**

Previous studies have demonstrated that mammalian LPP3 interacts with a subset of integrins to mediate cell functions [35, 36]. βv and αPS3 (scb) integrin subunits function as heterodimers in the regulation of ACs clearance and phagocytosis of S. aureus in D. melanogaster [12, 13]. We therefore investigated whether Wun2 could interact with αPS3 and/or βv integrin subunits and found a strong interaction between Wun2 and βv but not between Wun2 and αPS3 (Fig. 6A, F). Furthermore, the catalytic site-null Wun2 (DM) could still interact with βv (Supplementary Fig. 7A); the noncatalytic domain in Wun2-1 (Supplementary Fig. 7C), especially the region spanning the 116–134 residues, was required for Wun2 and βv interaction (Supplementary Fig. 7E). Hence, both Rip11 and βv integrin interact with the different residues of the noncatalytic domain of Wun2.

The suppression of βv or αPS3 integrin subunits in S2 cells led to an obvious reduction in ACs clearance (Supplementary Fig. 8A, B) and phagocytosis of S. aureus (Supplementary Fig. 8C, D); the wun2-RNAi-treated S2 cells also exhibited a significant decrease in the phagocytosis of bacteria (Supplementary Fig. 8C, D). The null allele itgbn^MB02607 mutant embryo failed to efficiently engulf ACs, exhibiting a phagocytosis rate of approximately 50.15% ± 2.519% (Supplementary Fig. 8E, F) and a PI of 1.233 ± 0.06043 (Supplementary Fig. 8E, G). These results are consistent with those of previous studies [12, 13, 37]. The hemocyte-specific knockdown of rip11 or βv subunit using crq-GAL4 caused a significant inhibition of ACs clearance in embryos (Fig. 6G–I). Double RNAi of wun2 and rip11 or wun2 and itgbn in embryonic hemocytes exacerbated the ACs clearance defect (Fig. 6G–I). Heterozygous wun2/βv and βv^+/-;rab11^+/- embryos also exhibited similar defects in ACs clearance (Supplementary Fig. 8E–G). These results provide strong evidence of genetic interactions among wun2, rip11, rab11, and βv for ACs clearance in embryonic macrophages. Importantly, the defects displayed by wun2 and βv mutants could be rescued by the macrophage-specific expression of βv (Fig. 6J–L and Supplementary Fig. 8E–G).

Wun2 is required for efficient recycling of βv

As mentioned above, Wun2 interacted with the recycling pathway proteins Rip11 and Rab11. As so, we hypothesized that wun2 deficiency was likely to affect the recycling of βv integrin subunit to the macrophage surface. The wun2^KO mutant displayed relatively low levels of βv integrin protein in whole-cell lysates (Fig. 7A, B) but the βv mRNA expression remained unchanged (Supplementary Fig. 9A). Immunofluorescent staining of embryos indicated that most βv integrin was distributed in the plasma membrane of WT macrophages whereas an uneven and aberrant aggregated pattern was displayed in the cytoplasm of wun2^KO macrophages (Fig. 7C). Consistent with wun2^KO mutants, the depletion of rab11 induced relatively low levels of βv integrin (Supplementary Fig. 9B–E). In the wun2^KD S2 cells, the abundant βv integrin was transported to the lysosomes (Supplementary Fig. 9F, G). When wun2 RNAi cells were treated with the lysosomal acidification inhibitor chloroquine (CQ), the βv integrin levels were almost restored to that of the control cells (Fig. 7D, E). These results indicated that the loss of Wun2 resulted in routing βv integrin into the lysosomes, leading to its degradation.

**Fig. 7: Wun2 protects βv integrin from degradation by sorting the recycling pathway.**

The surface levels of transmembrane proteins are controlled by several processes, including protein synthesis, internalization, recycling, and degradation [38]. To determine whether the reduction of βv integrin in wun2-depleted cells results from inadequate recycling, we analyzed βv internalization and recycling by cell surface labeling with cleavable biotin [38, 39]. To determine βv internalization, the recycling pathways were inhibited with primaquine (PMQ). The βν integrin subunit was internalized with similar kinetics in wun2-knockdown (wun2^KD) and WT cells (Fig. 7F and Supplementary Fig. 9H). In contrast, the wun2^KD cells showed a significant decreased recycling rate of βν to the plasma membrane (Fig. 7G and Supplementary Fig. 9I). We then isolated macrophages from larvae and labeled them with the βv integrin subunit antibody on the cell surface; degradation and recycling pathway were inhibited using CQ and PMQ, respectively, and the βv integrin antibody-labeled macrophages were allowed to perform endocytosis; however, less punctate βv integrin subunit-labeled macrophages were detected in wun2^KO than in WT larvae, indicating that the βv integrin subunit is less distributed on wun2^KO macrophage surfaces (Fig. 7H, I). Collectively, these data demonstrated that Wun2 facilitates the recycling of internalized βv integrin to the cell surface; therefore, the loss of Wun2 inhibits the recycling of internalized βv integrin, resulting in the lysosomal degradation of the receptor.

Wun2 is an important regulator of integrin-mediated phagocytosis

Live cell tracking of efferocytosis could uncover multiple qualitative and quantitative features of coordinated clearance of apoptotic corpses [40]. Macrophages are highly motile cells that can dynamically alter their actin cytoskeleton to perform migration and phagocytosis. They use their actin-rich lamellipodia and filopodia to engulf ACs [41]. As the active state of integrin leads to the remodeling of the actin cytoskeleton [42], the insufficient expression of the βv integrin subunit in the wun2^KO mutant resulted in the disorganized actin cytoskeleton of macrophages, with smaller and fewer pseudopodia near the leading edge (Fig. 8A–D). In contrast, WT macrophages had rich branched F-actin meshes with large lamellipodial and filopodial protrusions (Fig. 8A–D). We injected fluorescent Annexin V into the ventral side of D. melanogaster embryos between the overlying epithelium and the central nervous system. In vivo imaging revealed that WT macrophages used their actin-rich lamellipodia and filopodia to envelop Annexin-V labeled ACs (Fig. 8B, E and Supplementary Video 3). The wun2 and itgbn mutant macrophages showed a significantly reduced pseudopodia structure. The lamellipodial protrusions of wun2 hemocytes were undynamic, thus affecting the ability of hemocytes to capture ACs (Fig. 8E, F and Supplementary Video 4).

**Fig. 8: Wun2 is important for F-actin arrangement at the protrusions to envelop ACs.**

Discussion

Removal of cellular corpses is important in both homeostasis and disease. Defective efferocytosis has been implicated in a variety of autoimmune and inflammatory diseases [4, 43,44,45]. Based on RNA-sequencing analysis, we attempted to identify novel genes that were involved in efferocytosis and their potential applicability in the discovery of novel treatments for diseases driven by defective efferocytosis. We identified a candidate gene, wun2, which acts as a regulator in macrophages to clear ACs. Wun2 participates in the Rab11-regulated recycling pathway, facilitating the recycling of the βv integrin subunit from endosomes to cell membrane surfaces, allowing the rearrangement of F-actin-rich lamellipodia and filopodia to envelop ACs (Fig. 8G).

The D. melanogaster LPPs Wun and Wun2 play a significant role in regulating the survival and migration of germ cells, and they have been reported to act redundantly in somatic cells and germ cells [17, 29,30,31, 46]. In a previous study using single-cell RNA-sequencing to map hemocytes across different inflammatory conditions in D. melanogaster larvae [47], we found that Wun2 were enriched in most hemocyte clusters. The mammalian homolog of Wunens (LPP3) is important in immune response; it allows for the efficient exportation of mature T cells from the thymus into circulation. The conditional deletion of LPP3 in epithelial cells or endothelial cells is sufficient to inhibit T cell egress [48]. However, the exact functions of LPP3 or Wunens in macrophages have not been studied. In contrast with germ cells and somatic cells, a previous study found that the localization and function of Wun and Wun2 differed in the trachea, which is consistent with the present results. Wun and Wun2 seem to play different roles in macrophages; while the loss of Wun2 leads to efferocytosis defects, Wun is not required for this process, supporting these LPPs function autonomously in the different tissues [19]. The phosphatase activities of Wunens are crucial for germ cell and caudal visceral mesoderm cell migration and survival [30, 31, 49]. In addition to its phosphatase activity, LPP3 exhibits noncatalytic effects at the cell surface [35, 36]. The transgenic expression of catalytic-site-null Wun2 in wun2^KO macrophages could rescue the defects in ACs clearance, demonstrating the diversity of Wun2 functions in the different cells. Therefore, we hypothesized efferocytosis may be independent of the Wun2 phosphatase activity in macrophages. However, various phosphatases synergize inside the organism, and whether other enzymes complement the catalytic function of Wun2 remains to be studied.

The subcellular localization of LPP3 is dynamic and cell-specific to the endoplasmic reticulum, endosomes, vesicular structure, or Golgi complex [50, 51]. Wun2 mainly localized to the intracellular membrane and interacted with GTP-loaded Rab11 in D. melanogaster macrophages. Rab11 is a GTPase localized to several intracellular membrane compartments, including endosomes and nascent phagosomes [52, 53]. Rab11 participates in the mobilization and recruitment of early endocytic compartments in macrophages to enhance phagocytosis and ACs clearance [53, 54]. Integrin subunits βv and αPS3 function as a heterodimer in regulating ACs clearance and S. aureus engulfment by phagocytes [12, 13]. Unlike many other cell surface receptors that undergo synchronized ligand-induced internalization and degradation, internalized integrin receptors that enter early endosomes can be recycled back to the plasma membrane directly from peripheral early endosomes via Rab4-regulated or Rab11-regulated pathways [23, 55, 56]. For example, the recycling of β1 integrin depends on SNX17 or SNX31 binding to sort integrin into recycling vesicles [57, 58]; VPS3 and VPS8 control β1 integrin trafficking [56]. The diverse functions of Rab11 are mediated by its effector proteins, which appear to operate as adaptor molecules to recruit various regulatory membrane transport proteins [59]. Rip11 is an effector for Rab11 that preferentially associates with the GTP-bound Rab11, regulating apical membrane trafficking via recycling endosomes in polarized epithelial cells [59]. Rip11 comprises a C2 domain, which is found in several proteins that serves as the protein–phospholipid or protein–protein binding modules involved in protein translocation to a specific region [60]. For example, the binding of the C2 domain to anionic phospholipids mediates the rapid translocation of protein kinases C to the plasma membrane [61]; the C2 domain of cytosolic phospholipase A2 is necessary and sufficient for its translocation to the nuclear envelope upon calcium ion influx [62]; the interaction of Rip11 C2 domain with neutral phospholipids is important for protein trafficking from recycling endosomes to the apical plasma membrane [59, 63]. The present study showed that the C2 domain is also important for the interaction of Rip11 with Wun2.

Mammalian LPP3 interacts with a subset of integrins to mediate cell–cell adhesion and functions [35, 36]; Wun2 is highly similar to LPP3, showing a strong interaction with integrin subunit βv. Accumulating evidence suggests that integrins connect the extracellular matrix to the cytoskeleton, allowing phagocytic cells to adjust F-actin-containing protrusions, including lamellipodia and filopodia [41, 42, 64]. Macrophages use two distinct modes of engulfment: lamellipodial and filopodial phagocytosis [41]. We concluded that Wun2 physically interacts with the Rip11 C2 domain and integrin subunit βv; Wun2 and Rip11 function in the Rab11 pathway to regulate the recycling of integrin subunit βv to the plasma membrane; Wun2 depletion increases βv integrin subunit transport to lysosome, resulting in its insufficient distribution on the cell surface of macrophages and thus to insufficient generation of lamellipodia and filopodia for engulfing ACs. However, the possible participation of Wun2 in modifying the endosomal membrane is unclear. Further research is required to determine whether Wun2 and Rip11 regulate the stability of integrin subunit βv to bind to endosomal membranes, or the budding and fission of endosomal lipid membranes, or bind to selective phospholipids to enrich these membranes.

Materials and methods

Cell culture and cell transfection

Drosophila melanogaster S2 cells were cultured in Sf-900TM II SFM (Gibco # 10902088) or Schneider’s Drosophila Medium (Gibco # 21720001) with 50 U/mL penicillin and 50 μg/mL streptomycin at 25 °C. S2 cells were then transfected with Fugene HD reagent (Promega # E2311) for transient expression or selected with 5 μg/mL puromycin to generate stable S2 cell lines. HeLa cells were grown at 37 °C and 5% CO₂ in Dulbecco’s modified Eagle medium (Gibco # 11965175) supplemented with 10% fetal bovine serum, 1% penicillin, and 1% streptomycin. Additionally, cells were transfected with Lipofectamine^TM 2000 (Thermo # 11668019). All the cell lines were confirmed by ATCC via short tandem repeat profiling and expanded at low passages and stored in liquid nitrogen after receipt. Cells were used within ten passages for experiments or resuscitated within 1 month.

Antibody generation

The anti-integrin subunit βv and anti-Wun2 antibodies were obtained by immunizing mice in our lab; the integrin βv immune protein fragment consisted of 1–360 amino acids and Wun2 was prepared by fusing the amino acid fragments 1–75 and 231–265, which were expressed in Escherichia coli as a glutathione S-transferase-fused protein and then purified. The generation and use of anti-Crq antibodies have been reported in a previous study [10].

RNA-sequencing analysis

Total RNA samples were extracted from S2 cells using the Direct-zol RNA miniprep kit (Zymoresearch #R2052) and reverse-transcribed using the Transcriptor First Strand cDNA Synthesis Kit (Roche #04896866001). RNA-sequencing was performed by Annoroad Corporation using a HiSeq 2500 platform (Illumina) for determining differential gene expression. Raw reads containing adapters, poly-N, and low-quality reads were filtered, and the effective data were mapped to the D. melanogaster reference genome using TopHat (version 2.0.12) [65]. After excluding ribosomal and transfer RNA, we estimated the gene expression based on FPKM. The p values were adjusted using the Benjamini and Hochberg method [66]. All RNA-seq data files are available from the NCBI SRA database (accession number PRJNA721223). The URLs is https://www.ncbi.nlm.nih.gov/Traces/study/?acc=PRJNA721223&o=acc_s%3Aa.

Double-stranded RNA production and RNAi treatment

Primer sequences used to generate specific double-stranded RNA (dsRNA) were obtained from the D. melanogaster RNAi Screening Center (DRSC) DRSC/TRiP Functional Genomics Resources (harvard.edu) and were predicted to have no off-targets and primers with T7 RNA polymerase binding sites at the 5′ end. Amplicons were obtained by PCR under the following conditions: 94 °C for 2 min; 30 cycles of 94 °C for 15 s, 56 °C for 30 s, and 68 °C for 45 s; and 10 min extension at 68 °C. dsRNA was generated using 1 µg of amplicon and a High Yield RNA Synthesis Kit (NEB #E2040S) for dsRNA synthesis. The dsRNA sequences are presented in Supplementary Table 3.

RNA preparation and quantitative real-time PCR

For quantitative analysis of gene expression, total RNA was extracted from S2 cells using the TRIzol reagent (Invitrogen #15596026) and the Direct-zol RNA miniprep kit (Zymoresearch #R2052). One microgram of RNA was reverse-transcribed with the Transcriptor First Strand cDNA Synthesis Kit according to the manufacturer’s instructions (Roche #04896866001). Quantitative real-time PCR was performed using cDNA, primers (Supplementary Table S3), and the SYBR Mix (Roche #04707516001) in ABI StepOne system (Perkin-Elmer). All experiments were performed in triplicate with mRNA samples extracted from at least three biological replicates. Relative gene expression was determined based on the cycle threshold value and normalized to that of the rpl32 gene.

In vitro S2 cell phagocytosis assay

As described in previous studies [67, 68], S2 cells were incubated with 0.25 μg/mL actinomycin D (Sigma #SBR00013) in a culture medium for 18 h, fixed in 10% formaldehyde, and resuspended in serum-free medium (SFM) at a concentration of 10⁷ cells/mL. Next, 1 mL ACs were treated with 5 μg/mL fluorescein-5-isothiocyanate (FITC, Thermo Fisher #F1906) and rotated for 1 h away from light to ensure even dispersion. The cells were then washed twice with SFM and resuspended in 1 mL SFM; FITC-labeled ACs were added to live S2 cells at a ratio of 10:1 and incubated for 6 h. Cells were counterstained with 25 mM CellTracker^TM Blue CMAC Dye (Thermo Fisher #C2110) for 1 h and then gently washed with phosphate-buffered saline (PBS); 0.4% trypan blue (Gibco, #15250061) was added to quench the fluorescence of non-phagocytic ACs. The proportion of S2 cells containing ingested ACs was determined by inverted fluorescence microscopy or flow cytometry, and the ratio of double-labeled live cells to all live cells was calculated as a percentage.

For the bacterial phagocytosis assays, the S. aureus strain RN4220 was transformed with the plasmid pRN11, which expresses the red fluorescent protein. The S. aureus cells were grown with constant aeration at 37 °C with B2 broth until they reached an optical density of 0.4–0.6 at 600 nm. The cells harvested from the B2 broth were centrifuged at 8000 × g for 10 min at 20 °C and then washed three times with deionized water and resuspended in 1 mL Sf-900TM II SFM medium. S2 Cells were plated at a density of 1 × 10⁶ cells/well in a 96-well plate and incubated for 6 h with 20 μL of the S. aureus culture. After this period, cells were mounted in Vectashield medium (Vector Laboratories) and observed on a confocal microscope (C2, Nikon instruments) using a 20× objective.

Fly strains and genetics

The following D. melanogaster strains were obtained from the Bloomington Stock Center: Rab11^j2D1, hml -GAL4, Itgbn^MB02607, wun2^TKO.GS01135, vasa–cas9, UAS-LifeAct-GFP, srpHemo-Moe-3×mCherry, UAS-GFP (IDs: 12148, 6395, 24022, 77295, 51323, 57326, and 35786). Crq-GAL4 (III) was a gift from Dr. Zongzhao Zhai. The following D. melanogaster RNAi strains were obtained from the Tsinghua Fly Center: rip11 (THU3832), βv integrin (THU0442), wun (THU0875), and wun2 (THU0869). The vectors pUAS-wun2, pUAS-βv integrin, pUAS-wun2 (DM), and pUAS-LPP3 were constructed in our lab. Briefly, selected genes were amplified by PCR and cloned into the Invitrogen pENTR/D-TOPO vector (Thermo Fisher #K242020). The Gateway™ LR Clonase™ II kit (Thermo Fisher #11791100) was used to move selected genes from vector pENTR/D-TOPO to pPWG following the manufacturer’s instructions. Transgenic lines were generated using the PhiC31-targeted integration system by Unihuaii Inc. All D. melanogaster stocks were maintained in incubators at 25 °C, unless otherwise indicated. The wun2^KO mutant line was generated by crossing the transgenic female wun2^TKO.GS01135 with vasa–cas9 males. vasa-cas9/+;sgRNA-wun2/+ flies were crossed with a Sp/CyO balancer strain. Based on heredity laws, we selected gene knockout flies.

Protein extraction and western blotting

Protein samples were extracted and homogenized on ice in lysis buffer (50 mM Tris-HCl [pH 7.4], 1% TritonX-100, 0.15 M NaCl, 1 mM EDTA) with protease inhibitors. Lysates were centrifuged at 10,000 × g for 30 min, and the supernatants were collected and quantified using the bicinchoninic acid assay. Equal protein amounts were separated using electrophoresis and transferred to polyvinylidene difluoride membranes, which were blocked and incubated overnight with primary and secondary antibodies coupled with peroxidase. The antibodies were used at the following dilutions: mouse anti-actin (CST), 1:1000; mouse anti-Flag (Sigma) 1:1000; mouse anti-HA (CST) 1:1000; mouse anti-GFP 1:1000, mouse anti-Wun2 1:500, mouse anti-βv integrin 1:500, and secondary antibodies 1:10 000 (all Jackson ImmunoResearch Laboratories). Enhanced chemiluminescence western blotting detection was performed according to the supplier’s protocol (Pierce). All experiments were repeated at least three times.

Immunoprecipitations and LC-MS/MS analysis

S2 cells were transfected with full-length wun2 expression plasmids, harvested, and lysed in 50 mM Tris HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, and 1% NP40 with complete EDTA-free protease inhibitor. Lysates were centrifuged at 10,000 × g for 30 min, and the supernatants were incubated with anti-Flag M2 Affinity Gel (Sigma #A2220) for 6 h at 4 °C. The resin was centrifuged at 1400 × g for 1 min and washed thrice with Tris-buffered saline; 50 μL of 2× sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer was added and boiled for 5 min. Supernatants were subsequently transferred for silver staining or LC-MS/MS analysis (Hoogen Biotech Co.).

Yeast two-hybrid assays

For binary yeast two-hybrid assays, transmembrane proteins using the split-ubiquitin system, which inserts genes of interest into Cub or Nub as vectors and transforms vectors into yeast THY.AP4 [69]. Yeast was obtained from a stock plate, added to 15 mL yeast peptone dextrose adenine (YPDA) medium, inoculated in a 50 mL sterile tube, and incubated in a shaker overnight at 280 rpm and 30 °C. Next, 300 mL of YPDA was inoculated with 15 mL of overnight culture in autoclaved flasks and incubated with shaking at 280 rpm and 30 °C for 3–4 h. Then, cells were harvested and washed with sterile water, resuspended in 1.8 mL of 0.1 M LiAc, transferred to a 2 mL Eppendorf tube, and centrifuged at 2000 × g and 4 °C for 30 s. The supernatant was discarded, and an appropriate amount of 0.1 M LiAc was added to the tube, which was incubated at 25 °C for 30 min. Sterilized PCR tubes were prepared with 10 µL of single-stranded DNA and 1 µg of plasmid DNA for each transformation mix, 70 µL of polyethylene glycol solution, 10 µL of 1 M LiAc, and 20 µL of competent yeast for each transformation. The mixture was stirred and incubated at 30 °C for 30 min, and cells were heat-shocked at 42 °C for 40 min. The yeast was then centrifuged at 2000 × g for 1 min, washed with 100 µL of sterile water, resuspended in 50 µL of sterile water, plated on synthetic-defined (SD)-Leu/Trp drop-out plates, and grown for 3–4 days at 30 °C. Finally, three representative colonies were selected and diluted for spot testing. The same concentration of yeast was spotted on selected SD-Ade/His/Trp/Leu plates.

Embryo immunostaining

Immunostaining was performed according to the method described in previous studies [70, 71]. Briefly, staged embryos were fixed in 4% paraformaldehyde and stained with Crq antibody, which specifically labels macrophages while Dcp-1 antibody labels ACs [70, 72]. Rabbit or mouse fluorescein-coupled secondary antibodies were used at a 1:1000 dilution. Stained embryos were mounted in Vectashield medium, observed using a confocal microscope (C2, Nikon Instruments), and photographed at 20× and 100× magnifications. Images were processed using Adobe Photoshop CS6 and ImageJ.

Phagocytosis ratio and phagocytic index

The phagocytosis ratio was quantified as the number of engulfed ACs compared to total ACs, and phagocytic index (average number of ACs per hemocyte, PI) was determined as previously described [71]. Phagocytosis ratio and PI were calculated from confocal images acquired from ten embryos in which three confocal image stacks (head, CNS and tail) of 6 sections through macrophages per embryo were taken.

Cell immunofluorescence assay

S2 or HeLa cells were plated on a cover glass and fixed in 4% paraformaldehyde for 5 min, incubated in 1% bovine serum albumin (BSA) in 0.1% PBS-Tween for 1 h, and then incubated with the antibody overnight at 4 °C in PBS containing 1% BSA and 0.1% Tween. Alexa Fluor 594-conjugated goat anti-rabbit antibody was used at a 1:500 dilution for 1 h. Hoechst 33342 was used to stain cell nuclei. The samples were imaged using a laser scanning confocal microscope (C2, Nikon Instruments).

Macrophage isolation and immunostaining

Isolation of macrophages from D. melanogaster pupae was performed as previously described [73, 74]. Briefly, late third instar larvae were cultured for an additional 1–2 h until the formation of white pupae; ten white pupae were then bled at the dorsal side into a cell culture medium. The cell suspension was transferred to a plate on top of a glass coverslip, and allowed to spread on coverslips for 1 h at 25 °C. For the endocytic trafficking of βv integrin subunits from the cell surface by surface labeling [38, 39], cells were washed with cold PBS and incubated with an anti-HA antibody (1:100) for 30 min on ice. The surface-bound antibody was allowed to internalize for 2 h at 25 °C in SFM medium with 1 µm lysosome inhibitor CQ and 0.6 µm PMQ. Thereafter, the samples were washed with cold PBS, and the remaining antibody at the cell surface was removed through two acid washes (0.2 M acetic acid, 0.5 M NaCl, both in PBS) for 2 min on ice. Subsequently, the cells were fixed, permeabilized, and stained, as described in the cell immunofluorescence assay.

Phosphate release assay

The malachite green phosphate assay kit (Sigma #MAK307) was used to analyze the enzymatic reaction [75, 76]. The assays were performed in 96-well plates at 25 °C. Briefly, 50 µL of the anti-Flag resin containing the immunocaptured proteins was added to each well, and the phospholipids LPA and C1P were dissolved in 50% and 100% ethanol, respectively, at a final concentration of 500 µM. PA in 100% ethanol was used at a final concentration of 70 µM; S1P was dissolved in methanol to a final concentration of 100 µM and supplemented with protein lysis buffer to a final volume of 80 µL. The enzymatic reaction was stopped with 20 µL of malachite green reagent at the indicated time point and the resulting absorbance was measured at 620 nm on a plate reader (BioTek ELX808) at a final volume of 100 µL. Each reaction was performed in triplicate and repeated thrice.

Integrin internalization

The internalization of integrins was determined by biotinylation [38, 39]. The following steps were performed on ice unless otherwise noted. S2 cells were serum-starved for 1 h, washed twice in cold PBS, labeled with 200 µg/mL Sulfo-NHS-LC-biotin (Thermo Fisher Scientific #A39257) for 30 min at 4 °C, and then incubated in regular growth medium at 25 °C in the presence of 0.6 µM PMQ (recycling pathway inhibitor) for 0, 5, 10, and 15 min to allow internalization. At the indicated times, cells were rapidly transferred to ice, and the medium was aspirated. The cells were washed twice with ice-cold PBS and incubated with 20 mM MesNa (in 50 mM Tris pH 8.6 and 100 mM NaCl) for 15 min to remove surface biotin, and 20 mM iodoacetamide (IAA) was added for 5 min to quench MesNa. Cells were lysed (75 mM Tris, 200 mM NaCl, 7.5 mM EDTA, 7.5 mM EGTA, 1.5% Triton X-100, 0.75% IgepalCA-630, and protease inhibitors), and lysates were centrifuged at 20,000 × g for 30 min at 4 °C. Supernatants were collected and analyzed using capture enzyme-linked immunosorbent assay (ELISA) or immunoprecipitation. Each reaction was performed in triplicates and repeated thrice.

Integrin recycling

For the integrin recycling assay, S2 cells were serum-starved for 1 h, washed twice in cold PBS, labeled with 200 µg/mL Sulfo-NHS-LC-biotin (Thermo Fisher Scientific #A39257) for 30 min at 4 °C, and then incubated in regular growth medium (without PMQ) at 25 °C for 30 min to allow integrin internalization. After this period, cells were washed twice with ice-cold PBS and incubated with 20 mM MesNa for 15 min to remove surface biotin; 20 mM iodoacetamide (IAA) was then added for 5 min to quench MesNa. The cells were incubated in regular medium at 25 °C for 0, 5, 10, and 15 min to allow for biotinylated-βv integrin subunit recycling to the plasma membrane. Following a second reduction with MesNa to remove the recycled biotinylated-receptors, IAA was added for 5 min to quench MesNa. Cells were collected and lysed and centrifuged at 20,000 × g for 30 min at 4 °C. Supernatants were collected and analyzed using capture ELISA or immunoprecipitation.

Capture ELISA

Capture ELISA was performed as previously described [38, 39]. Ninety-six-well plates were coated overnight with anti-βv integrin antibodies (diluted at 1:200 in 0.05 M Na₂CO₃, pH 9.6) at 4 °C. Unspecific binding was blocked with 5% BSA in PBS-T (0.05% Tween-20 in PBS) for 1 h at 25 °C. Fifty microliters of cell lysate were added for 8 h at 4 °C to capture βv integrin subunit. The cell lysates were then removed and washed with PBS-T twice. Plates were incubated with streptavidin-horseradish peroxidase (HRP) in 1% BSA in PBS-T for 1 h at 4 °C. Biotinylated βv integrin subunit was detected using a chromogenic reaction with o-phenylenediamine (OPD), and the plates were read at 490 nm.

Proximity labeling with TurboID

Enzyme-catalyzed proximity labeling (PL) is a popular approach for studying the spatial localization and interactions of proteins in living cells. TurboID, the promiscuous ligase, enables nontoxic biotin to label the neighboring proteins [77, 78]. Proteins biotinylated by TurboID were enriched with streptavidin beads and identified by western blotting. Briefly, exponentially grown S2 cells were co-transfected with TurboID-fused construct (bait) and protein vectors (prey); after 36 h, the cells were replaced with fresh culture medium containing 50 µM exogenous biotin for 1 h. The reaction was terminated by cooling cells to 4 °C and washing away excess biotin. Cell pellets were lysed in IP lysis buffer as described above, and centrifuged at 20,000 × g for 20 min at 4 °C, and the upper soluble fraction was retained. For enrichment of biotinylated proteins, 20 µL of streptavidin-coated magnetic beads (Thermo Fisher Scientific #88817) were washed twice with the IP lysis buffer and then incubated with lysates containing 2 mg protein for each sample on a rotator for 6 h at 4 °C. For quality control analysis, the beads were washed twice in 50 mM Tris-HCl (pH 7.5) and biotinylated proteins were eluted by boiling the beads in 40 µL of 2× protein loading buffer, run on SDS-PAGE for western blot analysis.

Live imaging of D. melanogaster embryos

Stage 13 embryos were dechorionated in bleach and mounted in Voltalef oil under a coverslip on a gas-permeable culture dish [79, 80]. In brief, embryos were collected from agar plates and washed in the cell strainer using water. Excess water was dried out and the embryos were dechorionated in 50% bleach for 2–3 min. Next, the embryos were extensively rinsed with water until they lost the bleach odor. The embryos were then attached to a coverslip coated with a strip of the heptane glue in the middle. The coverslip was placed in the dehydration chamber for approximately 5 min. After drying, the embryos were covered with halocarbon oil 700 (Sigma, #H8898) to avoid further dehydration. Next, 1 μL of Annexin V (Molecular Probes, #13203) was loaded into a needle, which was attached to a micromanipulator, and all the embryos were sequentially injected with a drop. Images were collected on a confocal microscope (C2; Nikon Instruments). To quantify the duration of ACs clearance, 45-min time-lapse videos were acquired with a z stack taken every 30 s. Time-lapse images were processed using ImageJ.

Statistical analysis

Statistical analysis was performed using GraphPad Prism software. One-way analysis of variance (ANOVA) or Student’s t test determined statistical significance at p < 0.05. In the plots, error bars represent the standard error of the mean (SEM), “*” stands for p < 0.05, “**” for p < 0.01, “***” for p < 0.001, and “****” for p < 0.0001. Ten embryos from each genotype were tested unless indicated otherwise.

Data availability

All data are provided in the manuscript and Supplementary files or are available from the corresponding authors upon reasonable request.

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Acknowledgements

We thank the Bloomington Drosophila Stock Center for the Drosophila melanogaster strains. Additionally, we thank Editage (www.editage.cn) for English language editing.

Funding

This study was partially supported by the National Natural Science Foundation Key Project (breeding program) of China (Grant no. 91954114 to HX), National Natural Science Foundation of China (Grant no. 31871387 to HX), National Natural Science Foundation of China Youth Program (Grant no. 31801164 to QZ), the program of Innovative Research Team for the Central Universities (Grant no. GK202001004 to HX), and the Fundamental Research Key Project Funds for the Central Universities (Grant no. GK202007009 to HX).

Author information

These authors contributed equally: Ning Gao, Qian Zheng, Yanzhe Wang

Authors and Affiliations

College of Life Sciences, Shaanxi Normal University, Xi’an, 710119, Shaanxi, China
Ning Gao, Qian Zheng, Yanzhe Wang, Xiaowen Li, Zhi Li & Hui Xiao
Medical College of Yan’an University, Yan’an, Shaanxi, 716000, China
Ning Gao

Authors

Ning Gao
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Qian Zheng
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Yanzhe Wang
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Xiaowen Li
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Zhi Li
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Hui Xiao
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Contributions

HX and NG conceived the study. NG and QZ performed most of the experiments. XWL, YZW, and ZL provided the materials and performed some experiments. HX, NG, and QZ wrote the manuscript.

Corresponding author

Correspondence to Hui Xiao.

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Competing interests

The authors declare no competing interests.

Ethics approval and consent to participate

All animal care and experiments were performed according to the “Guide for the Care and Use of Laboratory Animals,” which were approved by the Institutional Animal Care and Use Committee at Shaanxi Normal University, and all manipulations were conducted in accordance with established guidelines.

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original western blot

video 1. The dispersal of macrophages in wild-type embryo

video 2. The dispersal of macrophages in wun2 mutant embryo

video3. Time-lapse recordings of apoptotic cell clearance in wild-type embryo

video 4. Time-lapse recordings of apoptotic cell clearance in wun2 mutant embryo

Table 1

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Table 3

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Gao, N., Zheng, Q., Wang, Y. et al. Wun2-mediated integrin recycling promotes apoptotic cell clearance in Drosophila melanogaster. Cell Death Differ 29, 2545–2561 (2022). https://doi.org/10.1038/s41418-022-01039-3

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Received: 31 August 2021
Revised: 25 June 2022
Accepted: 04 July 2022
Published: 15 July 2022
Issue Date: December 2022
DOI: https://doi.org/10.1038/s41418-022-01039-3