Perilipin 1 Deficiency Prompts Lipolysis in Lipid Droplets and Aggravates the Pathogenesis of Persistent Immune Activation in Drosophila

Lipid droplets are monolayer organelles with a neutral lipid core surrounded by phospholipids [1]. These highly dynamic intracellular organelles are present in all eukaryotic and most prokaryotic cells [2]. LDs serve as a lipid buffer pool to coordinate the anabolism and catabolism of lipids and thus play an important role in maintaining the balance between energy expenditure and storage during biological processes. The immune response is a high-energy-cost process [3, 4]. Therefore, the precise regulation of the immunometabolic response is critical for host fitness [5, 6]. A large body of evidence has demonstrated the tight interaction between immunity and metabolism at the systemic level. However, at the organelle level, for example, the relationship between LDs and immune responses is still unclear.

The fruit fly, Drosophila melanogaster, is a productive animal model for studying immunometabolism, thanks to its powerful genetic manipulation and high degree of conservation for humans. In addition, the Drosophila fat body is analogous to the liver and adipocyte in mammals and contains a large number of lipid droplets. The fat body is also the central immune organ in flies. Hence, this makes the fat body an ideal place to dissect immunometabolic interactions [7, 8]. The immune deficiency (IMD) signaling pathway is a major innate immune signaling pathway in Drosophila which is homologous to the NF-κB/TNF signaling pathway in vertebrates [9]. IMD signaling is not only a dominant defense pathway against gram-negative bacterial infection but also reported to be involved in many non-immune processes [10‒12]. Briefly, recognition of bacterial peptidoglycan (PGN) by peptidoglycan recognition proteins (PGRPs) recruits the adaptor Imd and subsequently induces massive Relish/NF-κB-dependent transcription of antimicrobial peptides (AMPs) [9] which is also a primary hallmark of the Drosophila immune response. However, the relationship between the configuration of LDs and IMD signaling remains unclear.

As the major adipose organ in flies, fat body cells are full of LDs. LDs are dynamic, non-homogeneous intracellular organelles anchored to their surface by variable proteins [13, 14]. Our previous study has suggested that Plin1 (also called lipid storage droplet-1, Lsd-1), a constitutive protein of LDs, is transcriptionally downregulated in response to transient IMD signaling activation induced by bacterial infection [15]. Plin1 is involved in the regulation of LD morphogenesis by modulating protein flux on LDs [16, 17]. We found that enlarged LDs benefit flies against bacterial infection through antioxidative roles [15]. However, under long-term immune/inflammatory conditions, the function of LDs and the underlying mechanisms involving Plin1 require further investigation.

In this study, we induced persistent systemic innate immune activation by genetically overexpressing components of the IMD pathway in fly fat bodies. The morphology of LDs was tracked after long-term IMD activation. Plin1 was found to play a dual role in regulating LD remodeling and inflammatory pathogenesis at different stages of persistent immune activation conditions.

All flies were propagated at 25°C on standard cornmeal food (1 L food contains 77.7 g cornmeal, 32.19 g yeast, 10.6 g agar, 0.726 g CaCl2, 31.62 g sucrose, 63.2 g glucose, 2 g potassium sorbate, and 15 mL 5% Tegosept), 30–55% humidity with a 12 h/12 h light/dark cycle. Fly resources that were used in this study are as follows: w¹¹¹⁸ were used as wild-type controls if there was no additional indication. plin1³⁸, Bmm-GFP [18], UAS-plin1 RNAi, and ppl-GAL4 were kindly gifted from Dr. Xun Huang (Institute of Genetics and Developmental Biology, CAS). UAS-Bmm RNAi (V37880) was obtained from the Vienna Drosophila Resource Center. w¹¹¹⁸, GS106-GAL4, UAS-Rel.68, UAS-PGRP-LCx, and UAS-PGRP-LCa were obtained from the Bloomington Drosophila Stock Center. All flies used in this study were male.

For lifespan analysis, male flies were collected within 3 days after adult emergence and raised in the incubator (temperature: 25°C, humidity: 55%, 12:12 light:dark cycle). These flies were randomly divided into three separate vials in a number of 25 each for one replicate. At least three independent biological replicates were performed, with at least three biological replicates. Alive flies were counted and flipped to fresh food every 3 days.

For quantification of mRNA level, about 20 flies’ carcasses/fat body tissues were dissected in sterile PBS buffer on ice at indicated timepoints, immediately homogenized in TRIzol, and samples were stored at −80°C. RNA extraction was according to the manual of the commercial kit (Magen, Hipure Total RNA Plus Micro Kit). cDNA was synthesized by using the kit (abm, 5X All-In-One MasterMix) with 1 μg isolated RNA as templates in a 20 μL reaction system. Quantitative real-time PCR (qRT-PCR) was performed using an SYBR green kit (abm, EvaGreen Supermaster Mix) on an ABI 7,500 or ViiATM 7 thermocycler (Life Technology). At least four independent biological replicates per genotype were performed. Housekeeping gene rp49 is the reference gene for data normalization. Primer information for qRT-PCR is as follows: rp49 (F: AGA​TCG​TGA​AGA​AGC​GCA​CCA​AG, R: CAC​CAG​GAA​CTT​CTT​GAA​TCC​GG), Dpt (F: GGC​TTA​TCC​GAT​GCC​CGA​CG, R: TCT​GTA​GGT​GTA​GGT​GCT​TCC​C), plin1 (F: CAG​CGC​ATA​CCA​CTG​GTC​TAT, R: GCA​TTA​CCG​ATT​TGC​TTG​ACA​G)

Lipid droplet staining of flies’ fat bodies was performed as previously described [15]. In brief, adult male carcasses/fat body tissues were dissected and fixed in 4% fresh prepared paraformaldehyde (pH = 7.5) in PBS for 10 min on ice. After PBS washing, tissues were incubated in PBS containing 1 μg/mL of BODIPY 493/503 (Invitrogen) dye or 0.5 μg/mL Nile Red (Sigma) for 30 min on ice. DAPI (1 μg/μL, final concentration) was added to stain nuclei at the last 5 min of the staining process. The mounting medium (Vector, H-1,000) was used to mount tissues for microscopy analysis. The measurement of the average lipid droplet size was performed as previously described [15‒17]. Briefly, 30 fat body cells/nuclei of each genotype randomly selected from eight confocal images were used to analyze the lipid droplet size. The diameters of the three largest LDs per cell (or around per nucleus) were measured to quantify the relative size of LDs. To count the size distribution of lipid droplets, the average percentage of the indicated size range of lipid droplets per cell from 30 fat body cells was determined by using the “Analyze Particles” tool embedded in ImageJ software (https://imagej.nih.gov/ij/). To quantify the fluorescence intensity of GFP on the surface of lipid droplets, confocal images acquired from eight fat bodies were measured by ImageJ software.

Glyceride amounts were measured using a TG Quantification Kit (BIOSINO, TG kit). Briefly, for whole-body glyceride quantification, groups of 12 one-week-old male flies were collected and weighted, then immediately stored at −80°C for subsequent assay. Stored flies were homogenized in 200 μL lysis buffer (10 mm KH₂PO₄, 1 mm EDTA, pH = 7.4) with three fly body volumes of ceramic beads and inactivated in a water bath at 75°C for 15 min. The inactivated homogenate was homogenized again for 30 s and kept on ice ready for assay. For each glyceride measurement, 3 μL of homogenate was incubated with 250 μL reaction buffer at 37°C for 10 min. After removal of debris by centrifugation (2,000 rpm, 2 min), 150 μL of clear supernatant was used to perform a colorimetric assay in 96-well plate (Corning^® Costar) for absorbance reading at 505 nm. Glyceride level was normalized with fly weight in each homogenate (unit: nmoL/mg fly). For fat body glyceride quantification, 25 flies’ carcasses/fat body tissues were dissected and followed by an assay as described above.

RU486 induction was described as before [19]. Briefly, a 10 mg/mL stock solution of RU486 (mifepristone; Sigma) was dissolved in DMSO. Appropriate volumes of RU486 stock solution were diluted with water containing 20% ethanol to a final concentration of 50 μg/mL. 100 μL of the diluted RU486 solution was dipped onto the surface of fresh food in vials (diameter: 2 cm). The vials were then allowed to dry at room temperature for half a day or 4°C overnight. Flies were transferred to RU486-contained food and raised at 25°C, and fresh food was changed every 2 days.

We used dichlorofluorescein diacetate (DCFH-DA) labeling to detect ROS in the fat body. DCFH-DA (Beyotime, Reactive Oxygen Species Assay Kit) labeling of fresh dissected carcasses/fat body tissues was performed according to the manufacturer’s manual, which is based on the ROS-dependent oxidation of DCFH-DA to fluorescent molecule 2′-7′dichlorofluorescein (DCF). In brief, the tissues were incubated with PBS containing 20 μm DCFH-DA for 30 min at 30°C, washed with sterile PBS three times (3 min each) to remove free DCFH-DA that did not uptake by the cell, and then the flaky fat body cells attached to the inner carcass shell were immediately dissected out to mount and confocal image (Vector, H-1,000). It should be noted that the slices were confocal imaged using the exact same settings for control and experimental groups. The fluorescence intensity is proportional to the ROS levels, and the fluorescence intensity of DCF was quantified by using ImageJ software.

Adult male flies were collected within 3 days after eclosion and raised on standard fly food at 25°C. One week later, flies were randomly distributed in groups of 20 flies/vial and starved on 1% agar (dissolved in distilled water); new agar vial was changed every day. For the survival count, the deaths were scored every 2 h until all experimental files were dead.

About 10 one-week-old indicated flies were homogenized in 200 μL lysis buffer (50 mm Tris, 150 mm NaCl, 1% Triton X-100, pH7.4) with Protease and phosphatase inhibitor cocktail (Beyotime, P1048). After ice bath for 30 min, the supernatant was collected and mixed with protein loading buffer (Beyotime) and heated at 95°C for 7 min. Western blotting was performed following standard protocols. The following antibodies were used: rabbit anti-cleaved Caspase-3 (1:1,000, CST, 9,661), rabbit anti-tubulin (1:10,000, Beyotime, AF0001), and anti-rabbit IgG linked with HRP (1:10,000, Proteintech, SA00001-2).

The methods for Wolbachia [20], Microsporidian [21], and Nora virus detection [22] were described as before. For Wolbachia and Microsporidian detection, genomic DNA of indicated flies was extracted, and the PCR reactions were carried out in standard program. For Nora virus detection, total RNA from the indicated flies was extracted by TRIzol. The synthesized cDNA was used as template for the PCR reaction. Agarose gel electrophoresis was performed. The PCR reactions were carried out under the following conditions: 50°C for 30 min; 94°C for 2 min; 94°C for 30 s; 55°C for 30 s; and 68°C for 1 min for 30 cycles; 68°C for 5 min; and holding at 4°C. The information of primers was as follows: Wolbachia (16S rDNA F: TTG​TAG​CCT​GCT​ATG​GTA​TAA​CT, R: GAA​TAG​GTA​TGA​TTT​TCA​TGT; wsp F: GTCCAATARSTGATGARGAAAC, R: CYGCACCAAYAGYRCTRTAAA), Microsporidian (F: GAC​CTA​TTG​AGG​ACA​ATC​AGT​AGC, R: CTT​AGT​GAG​CTA​CGA​TTA​CTA​GGA), Nora virus (F: TGG​TAG​TAC​GCA​GGT​TGT​GGG​AAA, R: AAG​TGG​CAT​GCT​TGG​CTT​CTC​AAC)

E.coli (DH5α) was used in this study. A single colony was inoculated to 10 mL fresh LB medium and grown at 37°C with shaking. Grow the bacteria to an OD₆₀₀ of about 1.0. The bacterial culture was pelleted with sterile PBS buffer to the concentration of OD₆₀₀ = 1.0. We injected 50.6 nL of bacterial suspension into indicated flies with Nanoject II injector (Drummond). Infected flies about 20 per vial were maintained at 25°C. After indicated hours, 5 lived flies were collected and homogenized in sterile PBS. Add 100 μL liquid after dilution on LB plate and spread uniformly. The LB plate was incubated at 37°C. The CFU was counted, and the load was calculated.

LSM700 (Leica) and Olympus FV-1200 confocal laser scanning microscopy were used for imaging. Captured images were analyzed by implemented soft, respectively. ImageJ (https://imagej.nih.gov/ij/) was used for the analysis of fluorescence intensity and lipid droplets size.

All replicates are shown as the mean ± SD or mean with range. Statistical significance was determined using one-way ANOVA and t-test for pairwise comparisons. Log-rank test for survival curves comparison. All data processing was used with GraphPad Prism 7.0.

The sample size was determined according to the number of data points. Batches of the experiment were carried out to ensure repeatability and the use of enough animals for each data point.

Measures were taken to ensure randomization. Each experimental batch contained more animals than the number of data points, to ensure randomization and the accidental exclusion of animals. In vitro analyses were usually performed on a specimen from animals at each data point to ensure a minimum of three biological replicates.

Data collection and data analysis were routinely performed by different people to blind potential bias. All measurement data are expressed as mean ± SD to maximally show derivations unless otherwise specified.

The fat body is the central immune organ mediating the systemic immune response in Drosophila. IMD signaling pathway is a major innate immune signaling in flies, analogous to the NF-κB/TNF pathway in mammals. In order to investigate lipid metabolism under conditions of sustained immune activation, an RU486-inducible fat body-specific GAL4/UAS system was applied to induce chronic IMD signaling activation. After RU486 treatment, GS106-GAL4 constitutively drives the expression of two isoforms of IMD receptors, PGRP-LCx and PGRP-LCa. The expression level of an antimicrobial peptide, diptericin (Dpt), the target of IMD signaling, monitors the activity of IMD signaling. The resulting increase in Dpt expression was observed in the fat bodies of the flies along with the extension of drug feeding time (online suppl. Fig. S1A, B; for all online suppl. material, see https://doi.org/10.1159/000534099). Meanwhile, a continuous decrease in fat content was detected in whole bodies (online suppl. Fig. S1D–E). Consistent with our previous observation, there was a slight increase in the fat content of fat bodies at 12 h post-induction (hpi) (online suppl. Fig. S1F). This is reminiscent of the finding that transient IMD activation leads to an increase in fat body glyceride in the early phase of Escherichia coli (E. coli) injection [15]. However, when IMD activation was prolonged, the glyceride level of fat bodies started to gradually decrease after 24 hpi (online suppl. Fig. S1F). These results remind us that prolonged immune activation is a highly energetic process. Of note, all flies used in this study are free of Nora virus, Wolbachia, and Microsporidian(online suppl. Fig. S2A–C).

LDs serve as the major lipid buffering pools in fat body cells. The morphology of LDs was assessed after chronic IMD hyperactivation. After 1 week of induction, LDs became dramatically smaller, regardless of whether IMD receptors or the active form of Relish (Rel), the key transcription factor of IMD signaling, were overexpressed (Fig. 1a, b and online suppl. Fig S1C). The percentage of small LDs (<2 μm) in fat body cells increased to 78.7%, 82.9%, and 82.6% in flies ectopically expressing PGRP-LCx, PGRP-LCa, and Rel.68, respectively (Fig. 1c). Consistently, we also found that under the conditions of prolonged IMD activation, the glyceride content of fat bodies in flies decreased compared to controls at 1 week of age (Fig. 1d). Therefore, these results suggest that persistent immune activation is sufficient to cause a decrease in the size of LDs (Fig. 1a–c).

Plin1 is the key element in the regulation of lipid metabolism and LD size [16, 17]. We have previously reported that plin1 is downregulated in response to transient IMD signaling activation upon E. coli injection. In addition, plin1 depletion results in enlarged LDs, which benefit the host against ROS accumulation during bacterial infection [15]. Under conditions of sustained IMD signaling activation via overexpression of PGRP-LCx or PGRP-LCa in the fat body, the expression level of plin1 in the fat body decreased after drug feeding (Fig. 2a). As expected, further knockdown of plin1 in fat bodies reversed the reduction in LD size after 1 week of chronic IMD activation (Fig. 2b, c). The distribution of LD size was comparable to that before drug treatment (Fig. 2c and in contrast to Fig. 1c). Similar results were observed when we used another fat body-specific GAL4, ppl-GAL4, to drive UAS-Rel.68 to induce sustained IMD hyperactivation. Complete removal of Plin1 from ppl>Rel.68 flies using the null allele plin1³⁸ even made LDs bigger compared to their genetic controls at 1 week after eclosion (Fig. 2d). Notably, flies with constitutive IMD activation (ppl>Rel.68) were able to hatch without obvious developmental defects, although the level of apoptosis was slightly higher than in the genetic controls. Moreover, additional plin1 deficiency had no further effect on apoptosis (online suppl. Fig. S3). Taken together, these results indicate that Plin1 deficiency sufficiently ameliorates LD size reduction at the early stage (<1 week) of continuous IMD hyperactivation.

Immune activation is always tightly associated with ROS accumulation which is a major causative factor in inflammation-induced tissue damage. Given on the close connection between ROS levels and lipid metabolism [23], we monitored the ROS levels using fluorescent probe 2′,7′-dichlorofluorescein diacetate (DCFH-DA) in fat bodies. Indeed, ROS obviously accumulated in fat bodies 1 week after induction of IMD activation (Fig. 3a, b). plin1 deficiency, which made LDs bigger at the early stage of IMD hyperactivation (Fig. 2d), significantly reduced ROS accumulation in ppl>Rel.68 fat bodies (Fig. 3c, d). Thus, we just wondered whether plin1 deficiency might benefit the lifespan of flies with prolonged immune activation. Indeed, persistent IMD hyperactivation shortened the lifespan of the flies (Fig. 3e). To our surprise, plin1 deficiency could not extend the lifespan of flies with persistent IMD activation but instead accelerated mortality. This result seemed to be inconsistent with our previous observation that plin1 deficiency protects flies against transient bacterial infection [15]. Therefore, the underlying mechanisms need to be further investigated.

To better understand the function of Plin1-modulated LDs during long-term immune activation, we decided to monitor the morphology of LDs in ppl>Rel.68 and ppl>Rel.68; plin1³⁸/plin1³⁸ fat bodies for a long time. Strikingly, although plin1 deficiency indeed blocked LD size reduction at the early stage of IMD hyperactivation (the 1st week after eclosion) and even developed much larger LDs (Fig. 2b–d; Fig. 4a), these LDs sharply became as tiny as those of flies with only IMD activation since 2 weeks after eclosion (Fig. 4a, fatbody glyceride shown in online suppl. Fig.S4b). Statistical results confirmed that there were more large LDs (>4 μm) in ppl>Rel.68; plin1³⁸/plin1³⁸ fat bodies during the first week with IMD hyperactivation. Subsequently, after 2 weeks of continuous IMD activation, the distribution and the average size of LDs turned to the same level as in flies with or without plin1 deficiency (Fig. 4b, c). These results strongly suggest that larger LDs are preferentially broken down into tiny ones quickly during prolonged immune hyperactivation. To better assess the effect of plin1 deficiency on lipolysis, we performed a starvation experiment, in which flies were transferred to the 1% agar medium without any nutrition. The rate of fat depletion in plin1-deficient flies was much steeper than in their genetic controls, along with persistent IMD activation (Fig. 4d). Thus, high wasting and reduced lipid mobilization due to plin1 deficiency rendered flies with high-energy-cost immune activation much more sensitive to starvation (Fig. 4e). These results suggest that Plin1 likely has a dual function during different stages of chronic immune activation. Plin1deficiency leads to the formation of large LDs, which facilities ROS scavenging to benefit the host at the early stage of immune challenge. However, the resulting larger LDs due to plin1 deficiency prefer to dissociate quickly, which may lead to high wasting and produce excessive lipotoxicity at the later stage of immune hyperactivation.

A previous study suggested that Drosophila LDs lipolysis is predominantly mediated by Brummer (Bmm), the Drosophila homologue of mammalian adipose TAG lipase (ATGL) [24, 25]. To investigate whether Bmm plays a critical role in the reconfiguration of LDs during persistent immune activation, we further knocked down Bmm in the fat body of related flies. Depletion of Bmm sufficiently stopped LDs from minifying after 2 weeks of IMD hyperactivation, both in flies with and without plin1 deficiency (Fig. 5a, b). An allele containing GFP-tagged endogenous Bmm (Bmm-GFP) was then applied [18]. We found that Bmm proteins localized to LD membranes, particularly to the surface of small LDs at the 1-2w transition stage (around 10 days), suggesting their lipolysis function [24] (Fig. 5c). Plin1 deficiency significantly enhanced the recruitment of Bmm lipase to LDs surfaces during persistent activation of IMD signaling (Fig. 5c, d). It’s worth noting that Bmm-GFP signals were largely concentrated at the contact site between LDs, a sign of overactive lipolysis [24]. In addition, long-term immune activation is a physiological process with high energy consumption, which is extremely detrimental to the lifespan of an organism (Fig. 5e). However, knockdown of Bmm in fat bodies well rescued the shortened lifespan due to the long-term immune hyperactivation (Fig. 5e). Moreover, the observation that further Bmm knockdown significantly reduced ROS accumulation in the fat body of ppl>UAS-Rel68; plin1³⁸/plin1³⁸ flies after 1 week (Fig. 5f) further supports our conclusion. Therefore, these results demonstrated that hyperlipolysis of LDs due to plin1 deficiency during prolonged immune activation is likely to be mediated by recruitment of Bmm.

The immune response is a highly energy-consuming process. The LD is a major intracellular place for the mobilization of neutral lipids. Therefore, lipolysis or lipid storage in LDs efficiently coordinates energy expenditure to fuel biological processes in the host. On the other hand, the formation of LDs is helpful to restore cytoplasmic free fatty acid, which leads to ROS accumulation via lipid beta-oxidation [26]. Thus, the homeostasis of LDs is critical for host fitness. Recently, LDs have been reported to function during pathogen infection [27]. However, their dynamic reconfiguration and function in long-term inflammation are still unclear.

Considering that the growth of LDs by downregulating plin1 is an active host adaptation against transient bacterial infection [15]. In this study, we trace the morphogenesis of LDs during genetically induced persistent IMD hyperactivation. plin1 expression shows a steady decrease in response to immune activation, as in our previous report [15]. During sustained hyper-activation of IMD signaling by genetic manipulation, only progressively smaller LDs were observed. We speculate that the growth of LDs by autonomous downregulation of endogenous plin1 in response to IMD activation was unable to overcome the breakdown of LDs due to robust lipolysis in highly energy-consuming hyperinflammation. Interestingly and impressively, although further removing plin1 from flies with IMD overactivation did enlarge LDs at the early stage of hyper-inflammation (less than 1 week after induction of immune hyperactivation), the disintegration of LDs in these flies was somewhat faster than in flies with IMD hyper-activation alone from the second week of IMD induction onwards. Mechanistically, the protein translocation assay confirmed that plin1 deficiency increased the recruitment of the key rate-limiting lipase, Bmm/ATGL, to the surface of LDs under conditions of sustained IMD hyperactivation. A recent finding showing a higher rate of ATGL-mediated lipolysis in the adipose tissue of Plin1^−/− mice [28] also supports our notion that Bmm/ATGL-mediated lipolysis predominantly occurs in large LDs induced by plin1 deficiency. Indeed, the high level of wasting on enlarged LDs, combined with the high energy expenditure due to persistent immune hyperactivation, leads flies into a severe “metaflammatory trap”, a state in which the sustained inflammation and metabolic dysfunction are caught in a vicious circle of mutual reinforcement that threatens their lifespan (Fig. 6).

In addition, AMPs are the main effectors of the innate immune response in Drosophila. The rapid and robust production of manifold AMPs may be a potential metabolic burden that affects the physiological state of Drosophila. The synthesis of AMPs during immune activation is a point of energy expenditure and partially depends on the breakdown of intracellular lipids. However, the role of AMPs in the lifespan is still unclear and controversial. Recently, Hanson and B. Lemaitre found no major effect of individual AMPs on lifespan; only ΔAMP14 flies (flies lacking 14 AMP genes from seven families) had a reduced lifespan [29], suggesting that collective AMPs may be beneficial to the host. They also implied that the IMD pathway may affect lifespan independent of regulating AMP genes. Whether Bmm, which facilitates the breakdown of LDs, further aids AMP synthesis is an interesting question that requires more investigation. We did measure the AMPs capacities between ppl>Rel68 and ppl>Rel68, Bmm RNAi flies during aging. We found that further Bmm knockdown did not affect AMP synthesis since the killing efficiency of injected E.coli in these flies was comparable (online suppl. Fig. S5). Furthermore, in our case of persistent IMD activation, Bmm knockdown alleviated the shortened lifespan. Taken together, these findings suggest, at least partially, that lipotoxicity induced by the IMD/Bmm axis, rather than AMP burden, is the main cause of shortened lifespan under conditions of sustained immune activation.

However, Plin1 likely has a dual function in the regulation of LDs along with long-term hyper-inflammation. In the early stages of immune responses, downregulation of plin1 can enlarge LDs, preventing the release of free fatty acids and reducing lipotoxicity and ROS-induced oxidative stress [30‒32]. Meanwhile, sustained hyperinflammation requires high energy consumption. Enlarged LDs are preferable to rapid lipolysis. Instead, Plin1 mutation enhances the breakdown of these large LDs by facilitating Bmm recruitment at the later stage of immune hyperactivation. The high wasting caused by plin1 deficiency exacerbates the inflammatory pathogenesis.

Taken together, the results in this study strongly suggest that the reconfiguration of LDs plays a critical role in the resolution of inflammation. And Plin1, the anchor protein on the surfaces of LD, plays an important role in linking LD morphogenesis and immunometabolic responses. Since Plin1 controls the protein flux on the surfaces of LDs [16, 17], it is worthwhile to take a detailed look at the dynamic protein composition on the surfaces of LDs during different stages of persistent immune activation. However, this study provides new insights into the function of LDs in the immunometabolic response and promises to pave the way for the development of potential therapeutic targets for hyperinflammation.

We thank Dr. Xun Huang (Institute of Genetics and Developmental Biology, CAS) for providing stocks of plin1³⁸ and Bmm-GFP flies.

An ethics statement was not required for this study type; no human or vertebrate subjects or materials were used.

The authors have no conflicts of interest to declare.

This work was supported by grants from the National Key Research and Development Program of China (2022YFC2303504), the Strategic Priority Research Program of the Chinese Academy of Sciences (XDPB16), the National Natural Science Foundation of China (92157106, 32270917, 82003361), the Shanghai Municipal Science and Technology Major Project (2019SHZDZX02), and the Shanghai Committee of Science and Technology, China (22ZR1469900).

Conceptualization and investigation: Lei Wang, Jiaxin Lin, and Lei Pan; methodology and validation: Lei Wang, Jiaxin Lin, Kaiyan Yang, Weina Wang, Yan Lv, Xiangkang Zeng, and Lei Pan; formal analysis: Lei Wang, Jiaxin Lin, Junjing Yu, and Lei Pan; writing-original draft: Jiaxin Lin and Lei Pan; writing-review and editing and supervision: Lei Pan; and funding acquisition, Junjing Yu, Yaya Zhao, and Lei Pan.

Lei Wang and Jiaxin Lin contributed equally to this work.

All data generated or analyzed during this study are included in this article. Further enquiries can be directed to the corresponding author.