Targeting membrane contact sites to mediate lipid dynamics: innovative cancer therapies

Abstract

Membrane contact sites (MCS) are specialized regions where organelles are closely interconnected through membrane structures, facilitating the transfer and exchange of ions, lipids, and other molecules. This proximity enables a synergistic regulation of cellular homeostasis and functions. The formation and maintenance of these contact sites are governed by specific proteins that bring organelle membranes into close apposition, thereby enabling functional crosstalk between cellular compartments. In eukaryotic cells, lipids are primarily synthesized and metabolized within distinct organelles and must be transported through MCS to ensure proper cellular function. Consequently, MCS act as pivotal platforms for lipid synthesis and trafficking, particularly in cancer cells and immune cells within the tumor microenvironment, where dynamic alterations are critical for maintaining lipid homeostasis. This article provides a comprehensive analysis of how these cells exploit membrane contact sites to modulate lipid synthesis, metabolism, and transport, with a specific focus on how MCS-mediated lipid dynamics influence tumor progression. We also examine the differences in MCS and associated molecules across various cancer types, exploring novel therapeutic strategies targeting MCS-related lipid metabolism for the development of anticancer drugs, while also addressing the challenges involved.

Introduction

Most eukaryotic cells rely on membrane-bound organelles to compartmentalize cellular components, enabling distinct biochemical reactions to occur independently within enclosed spaces. The regions where these organelles physically interact are referred to as membrane contact sites (MCS) [1]. These sites facilitate the exchange of materials and signaling between organelles, thereby promoting the coordinated regulation of homeostasis and cellular functions.

Lipid metabolism plays a crucial role in tumorigenesis [2]. Lipids and their metabolites are integral to numerous metabolic and signaling pathways, influencing key cancer traits such as energy metabolism, growth, proliferation, apoptosis, metastasis, and the immune microenvironment [2, 3]. The synthesis and degradation of lipids are vital to how cancer cells adapt to stress and damage. Tumor cells, even within the same tumor, display unique metabolic traits but also exhibit significant adaptability. This flexibility enables them to thrive under adverse conditions and drive malignancy progression [4]. Moreover, the uptake and synthesis of lipids play a critical role in determining the energy sources and functions of both cancer cells and various cell types within the tumor microenvironment [4]. However, due to compensation from dietary lipids, therapeutic strategies that simply inhibit fatty acid (FA) or cholesterol biosynthesis may have limited effectiveness in clinical practice [5]. Consequently, more targeted and combinatorial strategies are needed to intervene in lipid absorption, synthesis, and lipolysis [6]. Lipid metabolic processes are largely confined to membrane-bound organelles and are facilitated by physical interactions at membrane contact sites, which are formed through combinations of membrane proteins and/or lipids on both sides, fostering communication between organelles [7, 8]. This interaction, in turn, influences the activation and function of immune cells within the tumor microenvironment [4]. Therefore, modulating tumor lipid metabolism by manipulating organelle contact may serve as both a novel predictive marker and a therapeutic target.

In this review, we examine the mechanisms through which these highly dynamic organelles orchestrate lipid uptake, synthesis, recycling, distribution, and catabolism, with a particular emphasis on how lipid dynamics at organelle contacts within the tumor microenvironment influence both cancer cells and immune cells to support malignant tumor progression. Additionally, we critically assess the heterogeneity of MCSs across different cancer types and mutations, as well as the efficacy and challenges associated with MCS-related anticancer therapies.

Structure and function of membrane contact sites

Membrane contact sites primarily occur between the membranes of different organelles, particularly between membrane-bound organelles such as the endoplasmic reticulum (ER), mitochondria, and Golgi apparatus. These organelles are tightly interconnected through specific protein complexes, creating narrow gaps (typically less than 30 nm) that facilitate efficient material exchange and signal transduction [9]. The fundamental components of these cellular membranes are lipids, and their composition varies across different organelle membranes and/or subdomains within those membranes. Consequently, the formation and dissolution of membrane contact sites are highly dynamic processes, rapidly adjusting in response to the cell's needs [9].

The molecules associated with membrane contact sites include tethering proteins, lipid transfer proteins, Ca2⁺ channel proteins, and signal transduction molecules [1]. Tethering proteins are essential for anchoring membrane contact sites to the membranes of different organelles, ensuring stable interactions between them. For instance, the ER protein seipin and the mitochondrial membrane protein Mitofusin 2 (MFN2) play key roles in this process. Lipid transfer proteins, such as members of the oxysterol-binding protein-related protein (ORP) family, ceramide transporter (CERT), Acyl-CoA: Diacylglycerol acyltransferase (DGAT), and Fatty acid transport protein 1 (FATP1), facilitate the transport of lipid molecules at membrane contact sites, which is crucial for material exchange and signal transduction. Ca2⁺ channel proteins, like Calsyntenin 3 (CLSTN3), are responsible for regulating Ca2⁺ transport and maintaining homeostasis at these sites.

Membrane contact sites as a platform for lipid metabolism in cancer cells

Due to their substantial energy demands for survival and proliferation, most cancer cells exhibit increased de novo fatty acid synthesis [10]. The pyruvate generated from FA breakdown produces ATP through oxidative phosphorylation (OXPHOS), providing twice the energy compared to carbohydrates [10]. Thus, lipids serve as a highly efficient energy source, playing a critical role in fueling ATP production in cancer cells and supporting rapid tumor cell proliferation. In light of this, we first explore the role of MCSs in cancer by investigating their involvement in cellular lipid synthesis, metabolism, and transport (Table 1).

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Storage of lipids

ER-LD Contacts

Lipid droplets (LDs) are key organelles for storing neutral lipids and originate from the ER. As LDs grow, they expand their lipid monolayer by acquiring phospholipids from the ER via non-vesicular transfer, facilitated by ER-LD contact sites [11]. Here, the ER protein Seipin mediates specific lipid rearrangements that drive LD outgrowth, coordinating with enzymes like triglycerides (TG) synthases at these sites [12]. Seipin also interacts with other key molecules at the ER-LD interface, affecting sphingolipid metabolism and LD morphology [13]. This function is conserved across species, as human Seipin can correct defects in yeast [14]. In human breast cancer cells, lipid droplet assembly factor 1 (LDAF1) forms complexes with seipin to enhance LD budding and growth [15]. Moreover, exhaustion of Seipin in HeLa, leads to increased expression of DFCP1, a Rab18 effector. DFCP1 interacts with the Rab18-ZW10 complex to mediate ER-LD contact and increase LD size [16] (Fig. 1a).

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In tumor cells, key molecules like the Oxysterol-binding protein (OSBP) and related proteins (ORPs) play crucial roles in lipid synthesis and exchange at ER-LD contact sites. These proteins possess a conserved OSBP-related domain (ORD) that binds and transfers sterols and other lipids, with most members featuring FFAT motifs for ER targeting and pleckstrin homology (PH) domains for other organelles, promoting membrane contact and lipid transfer [17]. In HeLa and Huh7 cells, ORP5 (an integral membrane protein of the ER) localizes to ER-LD contact sites during oleic acid loading. ORP5 deficiency enhances neutral lipid synthesis and increases LD size [18]. Additionally, glucose restriction can induce a lipid phase transition within LDs in HeLa cells, affecting interorganelle transport [19]. In colorectal cancer cells, ER-localized lysophosphatidylcholine acyltransferase 2 (LPCAT2) is recruited to the LD surface, resulting in LD accumulation, reduced immunogenic cell death, and resistance to chemotherapy [20] (Fig. 1b).

DGAT2, a key enzyme in TG synthesis, interacts with monoacylglycerol acyltransferase (MGAT)−2 at the ER-LD interface in rat hepatoma cells, promoting lipid droplet expansion [21]. Overexpression of DGAT1 or DGAT2 results in different LD populations, with DGAT2 increasing larger LDs due to structural connections between the ER and LDs [22]. FATP1 works with DGAT2 to enhance LD expansion, with their interaction crucial for triglyceride synthesis and LD growth [23]. Real-time imaging shows that FATP1 and DGAT2, located near the LD surface, are essential for LD expansion, emphasizing the need for their proximity at the ER-LD interface. The disruption of the fat storage-inducible transmembrane protein 2 (FIT2) in mice leads to ER stress and liver damage, highlighting its role in lipid metabolism [24, 25]. In HepG2 cells, a human hepatocellular carcinoma line, FIT2 works with ER tubule-forming proteins Rtn4 and REEP5 and cytoskeletal elements Septin7 (an isoform of Septin) to promote LD biogenesis, demonstrating its critical role in maintaining lipid homeostasis and cell health [26] (Fig. 1c).

Obesity and its metabolic dysregulation are well-known cancer risk factors. In adipocytes, ER membrane protein Calsyntenin 3 (CLSTN3)β at ER-LD sites inhibits lipid transfer between LDs, preventing LD fusion and expansion, promoting lipid degradation and fatty acid oxidation [27]. BAP31 interacts with Perilipin1 (PLIN1) on the LD surface and is integral to this process. BAP31 depletion inhibits PLIN1 and LD degradation, thereby promoting abnormal LD growth and adipocyte expansion. This expansion adversely affects insulin signaling and exacerbates inflammatory conditions in white adipose tissue (WAT) [28]. Rab18, essential in 3T3-L1 preadipocytes, regulates LD growth and maturation. Rab18 deficiency results in fewer mature LDs and increased ER stress. The Rab18-NRZ-SNARE complex tethers the ER to LDs, essential for LD growth [29]. In a 3D adipocyte model, lumican induces TNFα-mediated inflammation, mimicking obesity-related fibrosis. Inflammatory condition leads to ER fragmentation, increased ER-LD contacts, and altered Rab18 dynamics, highlighting complex cellular interactions in obesity-related conditions [30] (Fig. 1d).

Nuclear membrane-LD contacts

The nuclear envelope (NE) is considered a specialized extension of the ER since the outer nuclear membrane (NM) is continuous with the ER, facing the cytoplasm with similar morphology and composition. The saturation level of lipid acyl chains is crucial for maintaining the structure and function of the NE and the nuclear pore complex. Hypoxia intensifies the effects of saturated lipids, leading to NE hardening and potential rupture. However, lipid droplets can buffer these saturated lipids, thus preserving NE integrity [31]. In aging Cryptomeria hidradii nematodes, an accumulation of lipid droplets along the nuclear membrane is observed, where the triglyceride lipase ATGL-1 at the nuclear membrane helps manage nuclear lipid droplet abundance. Reduced ATGL-1 activity leads to excessive nuclear lipid accumulation, disrupting nuclear homeostasis [32].

In mammalian cells, lipid droplets are also present within the nucleus, but nuclear lipid droplets (nLDs) differ from cytoplasmic lipid droplets (cLDs) due to their isolation by dual nuclear membranes. This separation means nLDs do not interact with cytoplasmic organelles, and the lipids they store are not as readily utilized by cytoplasmic processes. Nuclear LD formation occurs through at least two pathways: one involves de novo formation at the inner nuclear membrane (INM), seen in yeast where LDs bud directly from the INM. The protein seipin, localized to the INM, is crucial for forming membrane bridges connecting INM to nuclear lipid droplets. The other pathway involves transforming lipoprotein precursors synthesized in the ER. In hepatocytes, nuclear LDs originate from ER luminal lipoprotein precursors, while in non-hepatocytes and budding yeast, nLDs form de novo at the INM [33]. In both hepatocytes and non-hepatocytes, nLDs often co-localize with PML nucleosomes, serving innate immune functions. In the hepatocellular carcinoma cell line Huh7, PML-II, known for its liquid–liquid phase separation properties, is essential for nLD formation; overexpression of PML-II boosts nLD formation, while its knockdown decreases nLDs [34]. This suggests that PML-II’s phase-separation tendency plays a significant role in nLD formation and impacts interactions between nLDs and PML nucleosomes.

LD-LD contacts

In mammalian cells, LDs can fuse and grow through lipid droplet-lipid droplet contacts, with cell death-induced DFFA-like effector C (CIDEC) playing a crucial role in this process in adipocytes and hepatocytes [35]. CIDEC features a CIDE-N domain that forms a homodimer and a CIDE-C domain important for LD targeting and enrichment [36]. Additionally, a polybasic RKKR motif in the linker between CIDE-N and CIDE-C domains acts as a regulatory motif for LD fusion. Deleting this linker or mutating the RKKR motif results in oversized LDs due to enhanced fusion activity reliant on CIDE-N domain interactions. This RKKR motif interacts with acidic phospholipids via electrostatic attraction, and its deletion disrupts these interactions, promoting larger LD formation [37]. Wu et al. identified Rab8a as a direct regulator of Fsp27 (CIDEC)-mediated LD fusion in mouse adipocytes. AS160, a GTPase-activating protein (GAP) for Rab8a, forms a ternary complex with Fsp27 and Rab8a to enhance LD fusion, while MSS4 inhibits this process through its interaction with Rab8a [38].The co-expression of PLIN1 and FSP27 (CIDEC) not only increases the average size of lipid droplets but also promotes the formation of unicompartmentalized adipocytes. PLIN1 may facilitate CIDEC-dependent lipid exchange and droplet fusion by influencing CIDEC oligomerization [39]. Moreover, inhibiting protein phosphatase, Mg2+/Mn2+ dependent 1D (PPM1D) in mature white adipocytes significantly reduces LD size by dephosphorylating Ser511 of PLIN1, further impacting LD dynamics [40] (Fig. 1e).

Synthesis and catabolism of lipids

ER-mitochondria contacts

The mitochondria-endoplasmic reticulum contact site (MERC) is a critical nexus between the endoplasmic reticulum and mitochondria, essential for various cellular functions and related pathologies. At these sites, mitochondrial-associated membranes (MAMs) play a key role in lipid metabolism and inter-organelle communication [41, 42]. MERC is instrumental in synthesizing various lipids like phosphatidylethanolamine (PE), TG, and cholesterol. In the ER, phosphatidic acid (PA) is transformed into phosphatidylserine (PS) by PSS1 and PSS2, which is then transferred to mitochondria and converted into PE by PS decarboxylase (PSD) in the inner mitochondrial membrane. PE is quickly moved back to the ER and converted to phosphatidylcholine (PC) by phosphatidylethanolamine N-methyltransferase (PEMT) [43]. Mfn2, a mitochondrial membrane protein, links ER membranes to mitochondria, crucial for mitochondrial metabolism and energy balance in mice, and facilitates PE synthesis by binding to PS [44]. Long-chain acyl-CoA synthetase 4 (ACSL4) also plays a key role in these processes by binding lipid fatty acids to coenzyme A and is involved in TG synthesis, marking a reliable protein for MAMs and directing polyunsaturated fatty acids (PUFAs) to the TG synthesis pathway in hepatocytes [45]. Additionally, steroidogenesis at MERC involves cholesterol import from the ER to mitochondria for conversion into pregnenolone, a precursor for all steroid hormones, with many hormones synthesized by ER-localized enzymes [46] (Fig. 2a).

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LD-mitochondria contacts

The metabolic relationship between mitochondria and LDs is essential for lipid oxidation processes [47]. Nutritional stress enhances LD-mitochondrial interactions in an AMPK-dependent manner, promoting the transfer of fatty acids from LD to mitochondria [48, 49]. Khaddaj et al. found that co-expression of the mitochondrial outer membrane protein, OM14, with the LD-targeting protein perilipin 3 (PLIN3), creates a close junction between LDs and mitochondria [50]. In brown adipocytes, perilipin 5 (Plin5) on LD envelopes fosters LD expansion by enhancing TG synthesis and attracting peridroplet mitochondria (PDM) to LDs via its C-terminal region. Interestingly, changes in mitochondrial fusion-fission dynamics are seen as a result rather than a cause of LD recruitment [51]. In hepatocellular carcinoma (HCC), when intracellular FAs surpass mitochondrial degradation capacity, they are redirected to LDs via aldo–keto reductase family 1 member C3 (AKR1C3). This LD metabolism alleviates cytosolic lipotoxicity and reactive oxygen species (ROS) production, protecting HCC cells from sorafenib-induced mitochondrial toxicity [52] (Fig. 2b).

ER-mitochondria-LD crosstalks

LDs form essential contacts with the ER to facilitate biogenesis and interact with mitochondria to enhance fatty acid β-oxidation. Mitochondria can transiently or permanently interact with ER membranes to support LD biogenesis by supplying ATP or molecules for TG synthesis. During starvation-induced autophagy, DGAT1-dependent LD biogenesis linked to mitochondria via the ER undergoes ATGL-mediated lipolysis, supplying FAs for mitochondrial energy production. This mechanism also traps FAs in a TAG form, preventing acyl-carnitine accumulation that could cause mitochondrial dysfunction [53]. DGAT2, another ER enzyme, localizes to the MAM and may induce LD formation from mitochondrial contact sites in COS7 cells [54]. In adipocytes, the mitochondrial protein MIGA2 connects mitochondria, LDs, and the ER to enhance TAG synthesis, improving lipid storage in LDs [55]. Combot et al. found that seipin regulates ER mitochondria MAMs by binding to calcium regulators like SERCA2, IP3R, and VDAC, managing intra-mitochondrial calcium and energy metabolism. Seipin's binding to lipid droplets increases during lipid loading, and its deficiency in adipose tissue reduces ATP production, leading to lipodystrophy [56].

In tumor cells, lipid metabolism is influenced by mitochondria-ER-LD interactions. Guyard V et al. showed that ORP5 and ORP8 control LD biogenesis at the phosphatidic acid-rich MAM subdomain in HeLa cells. Oleic acid treatment significantly recruits ORP5-labeled MAM to nascent and existing LDs, indicating their role in both LD biogenesis and maintenance. ORP5/8 are crucial for recruiting seipin to MAM-LD contacts, and their absence impairs LD biogenesis. ER-mitochondrial contact site integrity is essential for ORP5/8’s role in regulating seipin-mediated LD biogenesis [57]. In acute myeloid leukemia cells, mitochondrial OXPHOS controls LD degradation through autophagy. Inhibition of the mitochondrial electron transfer chain (ETC) regulates autophagy; ETC inhibition leads to LD accumulation and impacts tumor growth by disrupting mitochondria-ER contact sites (MERCS) [58] (Fig. 2c).

LD-peroxisome contacts

Peroxisomes are crucial for lipid catabolism, particularly influencing MUFA-induced longevity in Cryptomeria hidradii nematodes. Cis-MUFAs like oleic acid increase both LDs and peroxisomes, altering lipid homeostasis essential for longevity. VPS13D and RAB7 regulate lipid transport and lipolysis between LDs and peroxisomes, with VPS13D essential for peroxisome biogenesis in mammalian cells, and RAB7, typically an endosomal protein, also localizing to LDs to influence lipolysis [59] (Fig. 2d).

During fasting, peroxisomes play a significant role in lipolysis, facilitated by PEX5, which aids the translocation of adipose triglyceride lipase (ATGL) to LDs. Fasting increases contact between peroxisomes and LDs via KIFC3, promoting ATGL's spatial relocation. PEX5 escorts ATGL to the contact point between peroxisomes and LDs, crucial for efficient ATGL translocation and maintaining energetic homeostasis during fasting [60]. PEX19 farnesylation is vital for inserting UBXD8 into the ER and LD compartments, differentiating HP-anchored proteins targeting LDs from bilayer-spanning peroxisomal proteins. This shared targeting mechanism suggests a coordinated biogenesis of LDs and peroxisomes within the ER [61]. Additionally, the hereditary spastic paraplegia protein M1 Spastin, an AAA ATPase on LDs linking them to peroxisomes, recruits ESCRT-III proteins IST1 and CHMP1B to LDs. These proteins, through their MIT domains, facilitate LD transport to peroxisomal fatty acids, potentially altering LD membrane morphology [62]. This complex network of proteins highlights a sophisticated system of organelle interaction and lipid metabolism regulation (Fig. 2d).

Transport and recycling of lipids

LD-autophagosomes/lysosomes contacts

LDs play a crucial role in autophagosome formation by providing essential lipids and serving as a platform for assembly at their interfaces. In liver cancer cells, it's noted that both apolipoprotein B (ApoB) proteasome degradation and autophagy/lysosomal degradation occur around cLDs. cLDs facilitate this process by providing a site for amphiphilic ApoB, allowing proteasomes and autophagosomes to colocalize and coordinate degradation activities [63] (Fig. 3a).

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Lipophagy, the autophagic breakdown of LDs, involves transporting intracellular proteins and organelles encapsulated by LDs into autophagosomes, which are then delivered to lysosomes for degradation and recycling as an energy source, crucial during starvation [64]. Chronic starvation activates a non-classical autophagy mechanism, particularly in differentiated mouse 3T3-L1 adipocytes or Huh7 human hepatoma cells, where the E2 enzyme ATG3 localizes to oversized LDs, initiating the attachment of the microtubule-associated protein 1 light chain 3B (LC3B). This creates LC3B-lipidated LDs near clusters of LC3B membranes, indicating the LD surface as a platform for LC3B lipidation and subsequent autophagic processes [65]. Further research shows that prolonged starvation leads to ATG3 binding not only to LDs but also to Atg8 family proteins near LC3B-positive membranes, which undergo lysosome-mediated acidification. This suggests that LD-lipidated LC3B acts as a tether, linking autophagic lysosomes to LDs and facilitating their autophagic degradation [66]. This complex interaction underscores the vital role of LDs in regulating cellular energy balance and stress responses through autophagy (Fig. 3a).

TGs and LD structural proteins co-localize with autophagic compartments, and inhibiting autophagy results in increased TGs and LDs both in vivo and in vitro. There is also an inverse relationship where high intracellular lipid levels impair autophagic clearance, evidenced by reduced co-localization of LDs with lysosome-associated membrane protein type 1 (LAMP1), no autophagic up-regulation in lipid-rich cultured hepatocytes, and decreased association between autophagic vacuoles and LDs in starvation-responsive, high-fat diet-fed mice. This cycle traps hepatocytes in a harmful loop where reduced autophagy promotes lipid accumulation, which further inhibits autophagy, increasing lipid retention. Notably, lysosomal inhibition slightly increases LD co-localization with LAMP1 [67] (Fig. 3a).

ER-Golgi contacts

The ER-Golgi system, which includes the ER, Golgi apparatus, and ER-Golgi intermediate compartment (ERGIC), is crucial in mammalian cells not only for processing proteins but also for lipid synthesis and organelle biogenesis. A key function of this system is non-vesicular lipid exchange at ER-Golgi MCS, which alters lipid composition between the Golgi's regions, enriching them differently in (glyco)sphingolipids and cholesterol.

Central to this process is ceramide (Cer) transfer protein (CERT), which features a PH domain for Golgi targeting and a START domain for ceramide transfer between membranes. Between these domains is a FFAT motif that binds with ER-resident VAMP-associated proteins (VAP)-A and VAP-B, facilitating efficient ceramide transport from the ER to the Golgi [68]. In Hela cells, CERT's phosphorylation at serine 315 near the FFAT motif enhances its interaction with VAP proteins, with hyperosmotic stress triggering this phosphorylation, thereby increasing its tethering capabilities across the ER [69, 70]. The N-terminal PH domain of CERT targets phosphatidylinositol 4-phosphate (PI4P) on Golgi membranes, while the C-terminal START domain manages ceramide transfer [71]. Additionally, hyperphosphorylation of the serine repeat motif (SRM) adjacent to the PH domain in Hela can inhibit CERT’s transport capability, suggesting a regulatory mechanism controlling ceramide flux based on CERT's phosphorylation status [72]. This intricate regulation highlights the complex interaction between protein modifications and lipid metabolism within the ER-Golgi system (Fig. 3b).

OSBPs and ORPs are key PI4P effector proteins, functioning as lipid transfer proteins that exchange trans-Golgi PI4P for ER cholesterol, promoting Golgi PI4P homeostasis [73]. Specifically, the PH domains of ORP9 and ORP10 target PI4P to facilitate its transfer at the ER-the trans Golgi network (TGN) contact site in Hela cells [74]. The ORP9-ORP11 dimer localizes at the Golgi-late endosomal interface, inhibiting lipid transport. ORP9 is anchored at the TGN via a tandem α-helix and interactions with ORP10 and ORP11, extracting PI4P from the TGN to prevent overaccumulation and inhibit OSBP-mediated PI4P-driven cholesterol transport [75]. Furthermore, GRAM domain containing 1 s (GRAMD1s) transports excess cholesterol from the Golgi back to the ER, preventing its buildup in Hela. ORP9 deficiency leads to cholesterol accumulation at the Golgi, worsened by GRAMD1s depletion, causing significant cholesterol build-up in the plasma membrane and triggering chronic SREBP-2 pathway activation [76]. VAP interacts with both ceramide transfer protein for ceramide translocation and OSBP for cholesterol translocation from the ER in Hela. Under high cholesterol conditions, ER-localized SCAP interacts with the VAP-OSBP complex via the PI4P phosphatase Sac1, facilitating the trans-transport of ER cholesterol and Golgi PI4P at the ER-Golgi MCSs. This interaction promotes the carriers of the TGN to the cell surface (CARTS) biogenesis, aiding the transport of cellular materials from the trans-Golgi network to the cell surface [77]. Additionally, VAP and Sac1 at the ER-Golgi contact site significantly reduce CARTS, impacting the translocation of pancreatic cancer upregulation factors (PAUF) from the TGN to the cell surface [78]. These dynamics highlight the crucial role of regulated lipid transport at the ER-Golgi contact in maintaining cellular cholesterol distribution and overall homeostasis (Fig. 3b).

Golgi-endosome contacts

The Rab11-binding protein RELCH/KIAA1468 facilitates non-vesicular cholesterol translocation from the recycling endosome (RE) to the TGN in Hela cell line [79]. In COS7 wild-type cells, the Golgi-derived SEC14L2 vesicle facilitates the conversion of PI4P to phosphatidylinositol 3-phosphate (PI3P), playing a critical role in regulating endosome division. The knockdown of SEC14L2 leads to a notable accumulation of endosomes, characterized by an increase in PI4P levels accompanied by a decrease in PI3P. This imbalance impairs endosome division, resulting in a significant enlargement of nuclear endosome size. This study unveils a highly synergistic interplay between different membrane systems where the SEC14L2 vesicle, operating at membrane contact sites, promotes phosphatidylinositol conversion crucial for endoplasmic reticulum-associated endosome division [80] (Fig. 3c).

ER-endosomal contacts

Lipid countertransport mediated by ORP10 at the ER-endosomal MCSs plays a crucial role in regulating retrograde membrane transport. ORP9 and ORP10 form dimers and, along with VAP, establish the ER-endosomal MCS in a process dependent on PI4P ORP10 facilitates the transfer of PS from the endoplasmic reticulum to the endosomes, in exchange for PI4P. This lipid exchange allows for the recruitment of the PS-binding protein EHD1, which is essential for promoting endosomal fission. This mechanism highlights the complex interplay between lipid dynamics and membrane trafficking processes [81]. A deficiency in membrane-associated protein A6 (AnxA6) triggers Rab7 activation, enhancing cholesterol transfer from late endosomes to the ER via a Rab7 and StAR-associated lipid transfer domain-3 (StARD3)-dependent MCS formation in AnxA6's absence [82] (Fig. 3d).

ER-lysosomal contacts

The ER-lysosomal contact site is crucial for metabolite transport, such as cholesterol and lipids, between organelles, essential for cellular metabolism and homeostasis. SNX13, an ER-localized protein, negatively regulates lysosomal cholesterol export and contributes to ER-lysosomal membrane contact site formation. When Niemann-Pick C1 (NPC1) function is impaired, SNX13 knockdown redistributes lysosomal cholesterol, leading to triacylglycerol-rich lipid droplet accumulation and increased lysosomal bis(monoacylglycerol) phosphates [83]. SNX19, localized to ER-endosomal and lysosomal contact sites, interacts with endolysosomes through its PX structural domain and PI(3)P. However, its PXA and PXC domains inhibit its role in facilitating ER-lysosomal network sharing. Under high fatty acid conditions, SNX19 also associates with ER-LD contact sites, showcasing its versatility in mediating both ER-lysosomal and ER-LD tethering [84] (Fig. 4a).

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Cholesterol within lysosomes is sensed by ORP1L, which communicates with the Rab7-RILP-p150Glued complex via ER-lysosomal MCSs. In high cholesterol conditions, such as in Niemann-Pick type C, this signaling is disrupted, causing lysosomal accumulation at microtubule negative ends due to inhibited kinesin motor activity [85]. Lysosomal membrane damage prompts swift ER contact establishment, facilitated by ER proteins VAPA/B. Cholesterol-binding protein ORP1L, recruited to damaged lysosomes through VAP-ORP1L interaction, is crucial for cholesterol accumulation. PI4P production on lysosomes post-damage by PI4K2A is vital, and PI4K2A knockdown prevents ORP1L and cholesterol accumulation, leading to failed lysosomal membrane repair. Additionally, OSBP, recruited upon injury, is essential; its depletion results in PI4P build-up and cell death [86]. In HeLa and HEK293 cells, transferring lysosomal cholesterol to the ER requires ORP1L conjunction with VAP, PI4P, sterols, and NPC1 expression [87]. These dynamics highlight the complex protein and lipid interactions at the ER-lysosomal interface, crucial for maintaining cellular lipid balance and responding to stress and damage. (Fig. 4a).

ER-PM contacts

ER-PM contact sites are crucial for maintaining balance among phospholipids, sphingolipids, and sterols, and play a key role in cholesterol management, which is vital for cell signaling and structural integrity of cell membranes. Cholesterol is primarily concentrated in the PM but is regulated from the ER where its biosynthesis and uptake occur. At these sites, GRAMD1/Aster proteins, which bind PS and cholesterol, facilitate PM-to-ER cholesterol transport. GRAMD1, anchored in the ER, features a GRAM domain exposed to the cytosol and a StART-like domain, allowing it to detect increases in PM cholesterol and facilitate its transfer to the ER [88, 89]. Additionally, ORP2 significantly impacts these contact sites by tetramerizing and delivering cholesterol to the PM non-vesicularly, exchanging it for phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2). This mechanism is vital for maintaining lipid dynamics and membrane composition [90]. ORP5 and ORP8 also contribute to lipid transport at the ER-PM interface by translocating PS to the PM, where it is exchanged for PI4P, driven by a PI4P concentration gradient. This gradient is maintained by PI4KIIIα, which keeps PM PI4P levels high, and SAC1P at the ER, which hydrolyzes PI4P to keep its levels lower, ensuring continuous lipid transport across the contact site [90] (Fig. 4b).

NM-mitochondrial contacts

The formation of contact sites between mitochondria and the nucleus plays a crucial role in facilitating communication between these two organelles during events such as the mitochondrial retrograde response and under conditions of proliferative stimuli. The protein complex TSPO-PKA-ACDB3, mediated by specific translocators, is instrumental in tethering mitochondria to the nucleus during the MRR. This interaction promotes the nuclear stabilization of pro-survival transcription factors, such as NF-κB, and is also involved in regulating cholesterol transport between the nucleoplasm and the cytoplasm. In environments rich in ROS and Ca2+ signaling, cholesterol can function as a secondary messenger, playing a pivotal role in mitochondrial-nuclear communication, particularly in reverse transcription processes [91]. This intricate interplay underscores the dynamic regulatory mechanisms that govern cellular resilience and adaptation in response to internal and external cues (Fig. 4c).

Lysosomal-mitochondria contacts

An increased prevalence of lysosomal-mitochondrial MCSs has been identified in cells depleted of NPC1 or Gramd1b, dependent on the late endosomal sterol-binding protein STARD3. This interaction is believed to contribute to mitochondrial cholesterol accumulation observed in NPC1-deficient cells [92] (Fig. 4d).

The impact of lipids on the tumor microenvironment via membrane contact sites

Tumor cells not only adapt to their high energy demands through lipid metabolism reprogramming facilitated by membrane contact sites, but also modulate the tumor microenvironment through MCS-related signaling pathways (Table 1).

Regulation of T cell activation and cytotoxicity by lipids via membrane contact sites

In colorectal cancer (CRC), elevated cholesterol levels promote the interaction between molecules on the endoplasmic reticulum (ER) and mitochondrial membranes in CD8+ T cells. These interacting molecules include Fis1/Bap31, MFN2/cox4/HSP90B1, VAPB/PTPIP51, and VDAC1/IPR3/GRP75. This interaction results in an increased expression of mitophagy-related proteins, such as BNIP3, PINK1, and Parkin, which subsequently contribute to CD8+ T cell exhaustion in CRC [93]. Following acute viral infection, the expression of Sel1L increases in antigen-specific CD8+ T cells, promoting the formation of MERCS and the expansion of ER-associated mitochondria. This process enhances fatty acid metabolism and oxidative phosphorylation, which in turn exacerbates ER stress [94]. ER stress is negatively correlated with T cell function and persistence [95]. Therefore, targeting MERCS to alleviate ER stress may provide a novel pathway to regulate T cell immunity and improve the efficacy of T cell-based immunotherapies [96]. When antigen-presenting cells (APCs) and T cells make contact, it triggers the accumulation of SNX27 at their contact plasma membrane. Concurrently, the PtdIns-binding region within the FERM domain of SNX27 binds to PtdIns(3,4,5)P₃, facilitating the trafficking of SNX27 from the plasma membrane back to the endocytic recycling compartment (ERC) through interactions between the plasma membrane and endosomes [97]. This lipid-mediated dynamic reshaping of the T cell membrane structure enables the signaling, adhesion, and membrane transport processes essential for transitioning to the activated state [97, 98] (Fig. 5a).

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The impact of lipids on b cell maturation and antibody production via membrane contact sites

In obesity, B cells are activated within adipose tissue. The activation of the B cell receptor (BCR) triggers a signaling cascade that recruits lysosomes to the immune synapse at the plasma membrane. These lysosomes contain accessory molecules, including GILT, H2DM, proteases, and MHC class II. Upon fusion with the synaptic membrane, lysosomes release their contents, facilitating antigen processing and presentation [99]. It has also been reported that in mice fed a high-fat diet (HFD), B cells migrate to the liver, where they promote inflammatory responses by secreting pro-inflammatory cytokines [99]. An obesogenic diet further drives the infiltration of mature B cells into visceral adipose tissue, resulting in an increase in the secretion of pro-inflammatory IgG2c [100]. In preclinical models, n-3 polyunsaturated fatty acids (PUFAs) can inhibit lipid-related chronic inflammation by increasing B-1 cells and antigen-specific IgM levels [101]. IgM antibodies may neutralize the pro-inflammatory effects of oxidized low-density lipoprotein (OxLDL) and inhibit its uptake by macrophages [101]. These findings highlight the delicate balance between lipids and B cell subsets, which can exert either positive or negative influences, depending on the types of antibodies they secrete [102] (Fig. 5b).

Lipids affect the functions of dendritic cells and macrophages through membrane contact sites

Tumor-associated dendritic cells (DCs) exhibit impaired antigen cross-presentation, which diminishes the effectiveness of anti-tumor immune responses. Gabrilovich and colleagues identified that this impairment is mechanistically linked to the accumulation of lipid droplets (LDs) enriched with electrophilic oxidized truncated (ox-tr-) lipids in DCs. These LDs bind to the stress-induced peptide chaperone heat-shock protein 70 (HSP70), reducing its availability in lysosomes. This depletion causes peptide-MHC complexes (pMHC) to accumulate in late endosomes/lysosomes, rather than translocating to the cell surface. As a result, tumor-associated DCs fail to induce robust tumor-specific CD8+ T cell-mediated responses [103]. This dynamic involving HSP70 across cellular compartments underscores the critical interplay between lipid metabolism and immune function within the tumor microenvironment. Phagosome maturation is essential for immune defense. In macrophages and DCs, the interaction between Sec22b and ORP8 at endoplasmic reticulum-phagosome contact sites regulates the phospholipid composition of phagosomes, thereby influencing their maturation. Depletion of Sec22b leads to enhanced phagolysosome fusion and exacerbates the degradation of ingested antigens [104]. During the differentiation of tumor-associated macrophages (TAMs), increased caspase-1 activity cleaves PPARγ, producing a truncated form that translocates to the mitochondria, where it inhibits medium-chain acyl-CoA dehydrogenase (MCAD). This inhibition results in lipid droplet accumulation in TAMs and increased lactate secretion, which contributes to the tumor-promoting effects of TAMs [105] (Fig. 5c).

Membrane contact sites mediating lipid dynamics in virus assembly

Hepatitis C virus (HCV) plays a significant role in hepatocellular carcinoma, with its particle assembly initiating on LD surfaces. Overexpression of seipin results in a 60% increase in LD size and a reduction in their number, leading to a 34% decrease in the total outer LD surface area per cell. This disruption impairs HCV assembly and reduces virus production [106]. The HCV core capsid protein binds to LDs and facilitates the assembly of viral particles within the ER. Its amphipathic helical domain (D2) specifically interacts with triglycerides in the ER membrane, rather than directly with LDs. Triglycerides in lipid droplets facilitate the α-helix folding of D2, enabling the core protein to reposition to the ER, where HCV lipoviroparticles likely assemble. Inhibiting DGATs prevents D2 from binding to triglycerides, disrupts its folding process, and consequently inhibits viral assembly [107] (Fig. 5d).

Furthermore, Yue et al. discovered that the coronavirus ORF6 protein targets LDs and regulates ER-LD interactions by binding to BAP31 and USE1. Mitochondria-LD interactions are facilitated by the sorting and assembly machinery (SAM) complex, and ORF6 expression enhances LD biogenesis and lipolysis, thereby reprogramming lipid metabolism, which is critical for virus production [108].

Heterogeneity of membrane contact sites across different cancer types

Different types of tumors exhibit distinct metabolic properties, potentially related to their membrane contact sites. Studies on the ultrastructure of MERCs in surgical specimens of human astrocytic tumors have revealed significant differences between well-differentiated glioma cells and poorly differentiated glioma-like stem cells. The latter exhibit a near-complete absence of MERCs [109]. Numerous studies suggest that the expansion of membrane contact sites plays a crucial role in helping cells adapt to stress responses. For instance, it has been shown that poorly differentiated glioma stem cells primarily rely on oxidative phosphorylation, whereas well-differentiated glioblastoma (GB) cells are "MERCs-rich," with MERCs facilitating mitochondrial function and supporting their metabolic needs [109]. Additionally, some studies suggest that glioma stem-like cells (GSCs) possess shorter mitochondria, leading to reduced ER-mitochondria contacts. This reduction results in lower surface expression of sialylated glycans, making GSCs more susceptible to killing by cytotoxic T lymphocytes and NK cells compared to differentiated glioma cells (GDCs). Based on these findings, we propose that MERCs could serve as a diagnostic tool to predict the responsiveness of glioma patients to immunotherapy [110].

It is noteworthy that the expression of MCS molecules and their impact on survival vary across different tumors. For instance, Seipin, a key tethering molecule of MCSs, is positively correlated with prognosis in several tumors, including pancreatic and ovarian cancers. However, in glioblastoma (GBM), high expression of the Seipin (BSCL2) gene is associated with poorer patient survival [111]. Similarly, upregulation of RAB18, an MCS-related lipid transfer protein, correlates positively with overall survival in breast cancer [112], yet in gliomas, RAB18 inhibits tumor cell apoptosis and reduces sensitivity to temozolomide [113]. These findings suggest that brain tumors may be particularly dependent on MCSs and their associated molecules. Additionally, studies have shown that the expression of OSBPL5 is significantly higher in Kirsten rats arcomaviral oncogene homolog (KRAS)-mutant tumors compared to KRAS-WT tumors across 33 different cancer types [114]. Therefore, further research into the cancer-specific and organ-specific roles of MCSs could provide valuable insights into potential therapeutic opportunities (Fig. 6).

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Tumor markers of membrane contact sites mediating lipid dynamics and their therapeutic implications

Given the crucial role of MCSs in tumorigenesis and development, there is significant potential for the development of novel targeted therapies aimed at signaling pathways on MCSs. These therapies could inhibit tumor cell growth and proliferation by disrupting lipid metabolism and signal transduction, while simultaneously enhancing the anti-tumor activity of the immune system (Tables 2, and 3). In addition to the conventional pathology techniques, including immunohistochemistry (IHC), special staining, next-generation sequencing (NGS), real-time quantitative PCR (qRT-PCR), and so forth, the advent of novel technologies such as fluorescence probe imaging, Hyperspectral coherent Raman scattering (HS-CRS) microscopy, and cryo-electron tomography (cryo-ET) has facilitated significant advancements in the detection of these targets (Fig. 6).

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Targeting molecules at membrane contact sites in cancer

DGAT1/2

The membrane contact site molecules DGAT1 and DGAT2 significantly impact various tumors with their pro-carcinogenic roles. DGAT1, overly expressed in prostate cancer cells, reduces LD density, microtubule organizing centers, and microtubule stability when inhibited, suppressing cell migration and growth. Such inhibitors have successfully curbed prostate tumor growth and migration [115]. In contrast, DGAT2 promotes metastatic growth in gastric cancer models and enhances cell proliferation in breast cancer MCF-7 cells, but its inhibition via PF-06424439 decreases LD formation, causes G2/M cell cycle blockade, and reduces cell invasiveness, particularly when combined with radiotherapy [116, 117]. Additionally, using DGAT1 and DGAT2 inhibitors has shown potential in inhibiting tumor growth in colon cancer xenografts, decreasing immunosuppression, and enhancing anti-tumor immunity [118]. Inhibition of DGAT1 has also been linked to lipid droplet formation, suppressing tumor growth in clear cell RCC [119] (Figs. 6 and 7).

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Although DGAT inhibitors are being explored in clinical trials for treating obesity, diabetes, and other metabolic disorders, their antitumor effects in preclinical cancer models are promising. However, the side effects, particularly gastrointestinal adverse events, observed in completed trials, along with their systemic expression and suboptimal pharmacokinetic profiles, have been limitations in the use of DGAT1 and DGAT2 inhibitors in cancer patients [120]. Ervogastat (PF-06865571), a novel and specific DGAT2 inhibitor, has demonstrated safety and efficacy in reducing hepatic steatosis in early clinical trials, with a well-tolerated safety profile in a Phase II trial for non-alcoholic steatohepatitis (NASH), reducing hepatic fat by 58.80% compared to the baseline [121]. Further research in larger Phase II trials for NASH with fibrosis is ongoing, evaluating the inhibition of DGAT2 and ACC [122]. Despite these advances, DGAT’s role in oncogenesis varies across different tumor types. For instance, DGAT1 overexpression in AML cells led to increased LD accumulation, lipid peroxidation, and apoptosis [123]. In contrast, overexpression of DGAT2 in HCC cell lines Hep3B and Huh7 resulted in inhibited cell proliferation [124]. These findings underscore the complexity of DGAT's functions in tumors and highlight the need for thorough investigation into the appropriate indications for DGAT inhibitors in cancer therapy (Fig. 6 and 7).

FATP1

FATP1 plays a key role in tumor progression by influencing lipid metabolism in tumor and immune cells. In melanoma, aberrant FATP1 expression enhances lipid uptake, promoting invasion and growth. Inhibiting FATP1 with Lipofermata reduces melanoma growth and invasion [125]. In B-cell lymphomas, FATP1 facilitates energy production from palmitic acid, boosting cell proliferation. Interestingly, while CD37 normally inhibits FATP1, its knockdown increases fatty acid metabolism and cell viability, suggesting FATP1 as a therapeutic target [126]. In multiple myeloma, FATP1 enhances fatty acid uptake, impairing bone marrow CD8+ T cell function. Inhibiting FATP1 restores CD8+ T cell activity, offering a new strategy to enhance immunotherapy outcomes [127] (Figs. 6 and 7).

In breast cancer, FATP1 is downregulated, with higher levels associated with longer disease-free survival and reduced M2-type macrophage polarization [128]. Overexpression of FATP1 in macrophages increases inflammatory cytokine production and fatty acid uptake, suggesting it inhibits tumor-promoting macrophages [129]. Resveratrol, which activates PPARα and PPARγ, reduces Fatp1 expression, decreasing metabolite accumulation in macrophages treated with oleic acid [130]. These insights highlight FATP1's complex roles in cancer metabolism and immune regulation, providing several potential therapeutic targets (Figs. 6 and 7).

CERT

CERT is responsible for the non-vesicular transport of ceramide from the ER to the Golgi, where it is converted to sphingomyelin. This process is particularly crucial in the context of glioma proliferation. Research has identified nitric oxide as an inhibitor of ceramide transport from the ER to the Golgi, which subsequently inhibits glioma cell proliferation [131, 132]. This suggests that blocking CERT-mediated ceramide transfer could be a novel anticancer therapeutic strategy. Fleury et al. have developed a protein-lipid interaction assay specifically for identifying new inhibitors of the CERT-Cer interaction. Their research led to the discovery of seven antagonist compounds (hit1-7), which were found to be three orders of magnitude more potent than the reference antagonist HPA12 in inhibiting CERT [133]. Additionally, compounds resembling lemon-like substances have been shown to inhibit sphingomyelin biosynthesis by blocking the CERT protein-dependent extraction of ceramide from the ER [134]. This approach disrupts specific lipid metabolic pathways in cancer cells, potentially leading to novel treatments for cancers like glioma (Figs. 6 and 7).

Relevant lipid metabolism interactions of organelles as cancer diagnostic and therapeutic targets

LDs as cancer diagnostic and therapeutic targets

The presence of an increased number of LDs in cancer cells suggests potential uses of LD detection as a diagnostic and prognostic biomarker [135]. LDs are stable in physiological environments, possess controlled physicochemical properties, participate in the interactions of other organelles regulated by lipid metabolism, making them promising targets for therapy [136].

Common methods for detecting LDs in a range of tumors include Perilipin-2 immunohistochemistry and Nile Red staining in paraffin sections [135]. While Nile Red is a convenient marker for LDs, it suffers from drawbacks such as nonspecific labeling of other lipid-containing organelles like lysosomes. boron dipyrromethene (BODIPY) dyes are preferred for more selective LD staining due to better cell permeability and specificity compared to Nile Red, despite limitations such as limited photostability and spectral crosstalk [135]. A novel development in LD imaging is the creation of a modular fluorophore platform based on boronic acid salicylidene perylene-BAZY dye. These dyes are synthesized through the condensation of boronic acid with a salicylidenehydrazone (BASHY) ligand system, featuring advantageous photophysical properties for bioimaging. Experiments with these dyes have shown their efficacy in selectively staining LDs in Hela cells without affecting cell viability, making BASHYs a promising tool for future bioimaging applications [137]. Additionally, the development of nanostructures as imaging or delivery agents for LDs can address issues like poor water solubility, non-specific staining, and short intracellular retention times of traditional organic probes. Klymchenko and colleagues have synthesized solvent discoloration probes based on Nile Red and other compounds that enable visualization of dynamic changes in lipid membranes through polarity sensing, offering enhanced utility in bioimaging applications [138]. Liu et al. developed a novel fluorescent probe, Lipi-Bright, with a unique ring-fused molecular structure that enhances photostability significantly [139]. This probe exhibits exceptional specificity and ultra-high fluorescence brightness for staining LDs, making them much brighter than those stained with traditional probes like BODIPY 493/503 or Nile Red. The superior brightness of Lipi-Bright enables detailed, quantitative tracking of LD dynamics through in situ time-lapse fluorescence imaging. For example, it was observed that cytoplasmic LDs move significantly faster (37 ± 15 nm/s) than nuclear LDs (24 ± 4 nm/s), and their mobility varies with different stimuli [139]. This detailed tracking enhances our understanding of lipid metabolism and advances research in this area. Additionally, the development of nanomaterial-based probes marks a significant advancement in bioimaging. These probes amplify signals from labeled antibodies and address self-quenching issues common with traditional immunofluorescent dyes, making them invaluable for highly sensitive detection of disease biomarkers [140] (Fig. 6).

HS-CRS microscopy is an advanced technique used to analyze the composition of individual LDs within cells, offering detailed insights into the cellular lipidomic environment [141]. This method helps in identifying and characterizing various cellular states and pathologies. Clinically, the presence of a white opaque substance, often found with LDs, has been noted to obscure microvascular patterns more in adenomas than in carcinomas, making it a useful diagnostic marker for distinguishing between these lesions [142]. The incorporation of such innovative imaging techniques and novel probes significantly enhances research and clinical practices, opening new avenues for diagnosing and understanding diseases, especially those involving altered lipid metabolism (Fig. 6).

LDs in cancer cells offer unique opportunities for targeted drug delivery due to their lipid-rich nature, which can be exploited to improve therapeutic outcomes by leveraging the aberrant lipid metabolism often present in cancer cells [136]. Zhang et al. explored the use of curcumin, in conjunction with pyrrolidine-2 (RSC-3388), an inhibitor of cytoplasmic phospholipase A2α—a key enzyme in LD formation. Due to its lipophilic nature, curcumin tends to localize within lipid membranes and LDs. By inhibiting LD formation, pyrrolidine-2 enhances curcumin's effectiveness by preventing its sequestration in LDs, thus potentially increasing its bioavailability and therapeutic impact in glioblastoma cells [143]. Additionally, it has been reported that flufenamic acid and 5-β cholic acid, which are specific inhibitors targeting AKR1C3, induce cytosolic lipotoxicity and mitochondrial dysfunction by promoting autophagy-dependent LD catabolism in HCC cells [52] (Fig. 7).

Photodynamic therapy (PDT) is increasingly used to target LDs in cancer cells [144]. Jiang et al. developed TPECNPB, an amphiphilic pyridine compound designed to target LDs through electrostatic interactions, enhanced for cancer cell specificity with a borate group responsive to high H₂O₂ levels in hypoxic environments [145]. Xia et al. introduced BODSeI, a BODIPY-based photosensitizer with improved singlet oxygen yield for effective LD targeting in cancer cells, increasing therapeutic potential [146]. Sun et al. synthesized heteroaryl-bridged NIR AIEgens, noted for high-fidelity LD imaging, biocompatibility, photostability, and lipophilicity; TPET-Fu among these generates ROS under white light, effectively killing HeLa cells [147]. Tan et al. developed the fluorescent probe TTIE for LD-targeted image-guided PDT in renal carcinoma, using a one-step synthesis method for precise cancer targeting [148]. Dai et al. designed and synthesized a lipophilic Near-infrared (NIR) aggregation-induced emission (AIE) nanomaterial that target LDs in HepG2 cells, demonstrating the potential of nanoparticle-mediated PDT in treating hepatocellular carcinoma [149]. Additionally, coating LD surfaces with specific proteins enhances versatility, enabling them to interact with different cell types or organelles, shifting focus from traditional therapies to targeted organelle-based approaches, opening new clinical applications and treatment perspectives [136] (Fig. 7).

Mitochondria as cancer diagnostic and therapeutic targets

Mitochondria are dynamic and pivotal organelles that play a crucial role in both the diagnosis and treatment of cancer. Developing visual diagnostic methods to study the interactions between mitochondria and other organelles is essential for understanding their relationship with lipid metabolism. Wang et al. developed a near-infrared (NIR) AIE probe, NAP-Py-E, which targets both mitochondria and lipid droplets. Initially, the probe targets mitochondria, emitting a reddish fluorescence. Upon enzymatic hydrolysis within the mitochondria, NAP-Py-E is converted to NAP- Py, which then specifically accumulates in lipid droplets and emits green fluorescence [150]. Zhang et al. utilized C21H19N3O2, 7-(diethylamino) coumarin-3-vinyl-4-mwhich can reversibly migrate between mitochondria and lysosomes depending on changes in mitochondrial membrane potential [151]. In yeast, lipid transfer is facilitated by the ER-mitochondria encounter structure (ERMES). Utilizing quantitative live-cell imaging and cryo-electron microscopy, it has been observed that ERMES forms approximately 25 discrete bridge complexes, irregularly distributed across the contact sites. Each bridge features three synaptic-like mitochondrial lipid-binding protein domains arranged in a zigzag pattern, which delineates the lipid transfer route between the ER and mitochondria [41]. György Csordás and colleagues used electron tomography to study these interactions in rat liver mitochondria, identifying granules linking the outer mitochondrial membrane to presumed ER vesicles [42] (Fig. 6).

CPI-613 (Devimistat), a mitochondrial metabolism inhibitor, has shown notable efficacy in clinical trials across various cancers. For metastatic pancreatic cancer, a phase I trial reported an objective response rate (ORR) of 61% and median survival times of 19.9 months for overall survival (OS) and 9.9 months for progression-free survival (PFS) in 18 patients treated with the maximum tolerated dose [152]. Common severe side effects included hypokalemia, diarrhea, abdominal pain, and sensory neuropathy [152]. In acute myeloid leukemia (AML), combining CPI-613 with high-dose cytarabine and mitoxantrone achieved a 50% ORR and 6.7 months OS in a phase I study [153], with a phase II study showing an ORR of 44% and an OS of 5.9 months [154]. For advanced cholangiocarcinoma, CPI-613 combined with gemcitabine and cisplatin resulted in a 45% ORR and a 10-month median PFS [155]. However, it was ineffective in treating relapsed or refractory small cell lung cancer [156]. Additionally, nanomaterial-based strategies targeting mitochondrial functions have enhanced therapeutic outcomes in other preclinical cancer studies, underscoring the potential of targeting mitochondrial dynamics in cancer treatment [157,158,159] (Fig. 7).

Lysosomes as therapeutic targets

Since acidic sphingomyelinase (ASM) activity is typically lower in cancer cells compared to normal cells, resulting in elevated sphingomyelin levels, cancer cells are particularly vulnerable to ASM targeting within the lysosome. Targeting ASM can induce lysosomal membrane permeability (LMP) by promoting the accumulation of sphingomyelin, which in turn triggers tumor cell death. Effective ASM inhibition has been demonstrated with functional inhibitors such as zoledronic acid, chlorpromazine, and cationic amphiphilic drugs (CADs), highlighting their potential utility in inducing cancer cell apoptosis through this pathway [160].

Conclusions and prospects

Membrane contact sites play a crucial role in maintaining cellular lipid homeostasis, which is essential for normal cell function and for understanding the unique metabolic characteristics of cancer cells. While previous studies have explored targeting molecules or organelles associated with MCSs for cancer treatment, most have focused on promoting tumor cell proliferation and preventing apoptosis through metabolic reprogramming. Emerging research, however, reveals that MCSs also influence immune cell activation and function, as well as tumor-associated virus packaging within the tumor microenvironment, thus contributing to the malignant progression of tumors. Therefore, future drug development targeting signal crosstalk at MCSs in the tumor microenvironment may offer novel strategies for cancer treatment. In the era of precision medicine, it is also important to further investigate whether there are distinct differences in MCSs and their associated molecules across specific cancer types or mutations. The application of advanced technologies, such as cryo-electron tomography, super-resolution microscopy, optical tweezers, DNA origami, and lipid probes, will facilitate the discovery of new structures and functions of MCSs, leading to more precise treatment options [8].

Data availability

No datasets were generated or analysed during the current study.

Abbreviations

ACSL4:

Long-chain acyl-CoA synthetase 4

AKR1C3:

Aldo–keto reductase family 1 member C3

AnxA6:

A deficiency in membrane-associated protein A6

ASM:

Acidic sphingomyelinase

ATGL:

Adipose triglyceride lipase

BASHY:

Boronic acid with a salicylidenehydrazone

BODIPY:

Boron dipyrromethene

CAD:

Cationic amphiphilic drug

CARTS:

Carriers of the TGN to the cell surface

Cer:

Ceramide

CERT:

Cer transfer protein

CIDEC:

Cell death-induced DFFA-like effector C

cLDs:

Cytoplasmic lipid droplets

CLSTN3:

Calsyntenin 3

cryo-ET:

Cryo-electron tomography

DGAT:

Acyl-CoA: Diacylglycerol acyltransferase

ER:

Endoplasmic reticulum

ERGIC:

ER-Golgi intermediate compartment

ERMES:

ER-mitochondria encounter structure

ETC:

Electron transfer chain

FA:

Fatty acids

FATP1:

Fatty acid transport protein 1

FIT2:

Fat storage-inducible transmembrane protein 2

GRAMD1s:

GRAM domain containing 1 s

HCC:

Hepatocellular carcinoma

HCV:

Hepatitis C virus

HS-CRS:

Hyperspectral coherent Raman scattering

HSP70:

Heat-shock protein 70

IHC:

Immunohistochemistry

INM:

Inner nuclear membrane

KRAS:

Kirsten rats arcomaviral oncogene homolog

LAMP1:

Lysosome-associated membrane protein type 1

LC3B:

Light chain 3B

LD:

Lipid droplet

LDAF1:

Lipid droplet assembly factor 1

LMP:

Lysosomal membrane permeability

LPCAT2:

Lysophosphatidylcholine acyltransferase 2

MAM:

Mitochondrial-associated membrane

MCAD:

Medium-chain acyl-coenzyme A dehydrogenase

MCS:

Membrane contact sites

MERC:

Mitochondria-endoplasmic reticulum contact site

Mfn2:

Mitofusin 2

MGAT:

Monoacylglycerol acyltransferase

NASH:

Non-alcoholic steatohepatitis

NE:

Nuclear envelope

NGS:

Next-generation sequencing

nLDs:

Nuclear lipid droplets

NM:

Nuclear membrane

NPC1:

Niemann-Pick C1

ORD:

OSBP-related domain

ORP:

OSBP-related proteins

OSBP:

Oxysterol-binding protein

OXPHOS:

Oxidative phosphorylation

PA:

Phosphatidic acid

PAUF:

Pancreatic cancer upregulation factors

PC:

Phosphatidylcholine

PDM:

Peridroplet mitochondria

PDT:

Photodynamic therapy

PE:

Phosphatidylethanolamine

PEMT:

Phosphatidylethanolamine N-methyltransferase

PH:

Pleckstrin homology

PI3P:

Phosphatidylinositol 3-phosphate

PI4P:

Phosphatidylinositol 4-phosphate

PLIN1:

Perilipin 1

PLIN3:

Perilipin 3

Plin5:

Perilipin 5

PM:

Plasma membrane

pMHC:

Peptide-MHC complexes

PPM1D:

Hosphatase, Mg2 + /Mn2 + dependent 1D

PS:

Phosphatidylserine

PSD:

PS decarboxylase

PUFA:

Polyunsaturated fatty acids

qRT-PCR:

Real-time quantitative PCR

RCC:

Renal cell carcinoma

RE:

Recycling endosome

ROS:

Reactive oxygen species

SAM:

Sorting and assembly machinery

SIM:

Structured illumination microscopy

SRM:

Serine repeat motif

StARD3:

StAR-associated lipid transfer domain-3

TAM:

Tumor-associated macrophage

TG:

Triacylglycerol

TGN:

Trans Golgi network

VAP:

VAMP-associated protein

WAT:

White adipose tissue

References

Prinz WA, Toulmay A, Balla T. The functional universe of membrane contact sites. Nat Rev Mol Cell Biol. 2020;21(1):7–24.

Snaebjornsson MT, Janaki-Raman S, Schulze A. Greasing the Wheels of the Cancer Machine: The Role of Lipid Metabolism in Cancer. Cell Metab. 2020;31(1):62–76.

Röhrig F, Schulze A. The multifaceted roles of fatty acid synthesis in cancer. Nat Rev Cancer. 2016;16(11):732–49.

Pavlova NN, Zhu J, Thompson CB. The hallmarks of cancer metabolism: Still emerging. Cell Metab. 2022;34(3):355–77.

Bian X, Liu R, Meng Y, Xing D, Xu D, Lu Z. Lipid metabolism and cancer. J Exp Med. 2021;218(1):e20201606.

Martinez-Outschoorn UE, Peiris-Pagés M, Pestell RG, Sotgia F, Lisanti MP. Cancer metabolism: a therapeutic perspective. Nat Rev Clin Oncol. 2017;14(2):113.

Xu J, Huang X. Lipid Metabolism at Membrane Contacts: Dynamics and Functions Beyond Lipid Homeostasis. Front Cell Dev Biol. 2020;8:615856.

Sun S, Zhao G, Jia M, et al. Stay in touch with the endoplasmic reticulum. Sci China Life Sci. 2024;67(2):230–57.

Duckney PJ, Wang P, Hussey PJ. Membrane contact sites and cytoskeleton-membrane interactions in autophagy. FEBS Lett. 2022;596(17):2093–103.

Wu H, Chen W, Chen Z, Li X, Wang M. Novel tumor therapy strategies targeting endoplasmic reticulum-mitochondria signal pathways. Ageing Res Rev. 2023;88:101951.

Petan T. Lipid Droplets in Cancer. Rev Physiol Biochem Pharmacol. 2023;185:53–86. (8+3).

Olzmann JA, Carvalho P. Dynamics and functions of lipid droplets. Nat Rev Mol Cell Biol. 2019;20(3):137–55.

Choudhary V, El Atab O, Mizzon G, Prinz WA, Schneiter R. Seipin and Nem1 establish discrete ER subdomains to initiate yeast lipid droplet biogenesis. J Cell Biol. 2020;219(7):e201910177.

Su WC, Lin YH, Pagac M, Wang CW. Seipin negatively regulates sphingolipid production at the ER-LD contact site. J Cell Biol. 2019;218(11):3663–80.

Chung J, Wu X, Lambert TJ, Lai ZW, Walther TC, Farese RV Jr. LDAF1 and Seipin Form a Lipid Droplet Assembly Complex. Dev Cell. 2019;51(5):551-563.e7.

Li D, Zhao YG, Li D, et al. The ER-Localized Protein DFCP1 Modulates ER-Lipid Droplet Contact Formation. Cell Rep. 2019;27(2):343-358.e5.

Wang H, Ma Q, Qi Y, et al. ORP2 Delivers Cholesterol to the Plasma Membrane in Exchange for Phosphatidylinositol 4, 5-Bisphosphate (PI(4,5)P2). Mol Cell. 2019;73(3):458-473.e7.

Du X, Zhou L, Aw YC, et al. ORP5 localizes to ER-lipid droplet contacts and regulates the level of PI(4)P on lipid droplets. J Cell Biol. 2020;219(1):e201905162.

Rogers S, Gui L, Kovalenko A, et al. Triglyceride lipolysis triggers liquid crystalline phases in lipid droplets and alters the LD proteome. J Cell Biol. 2022;221(11):e202205053.

Cotte AK, Aires V, Fredon M, et al. Lysophosphatidylcholine acyltransferase 2-mediated lipid droplet production supports colorectal cancer chemoresistance. Nat Commun. 2018;9(1):322.

Jin Y, McFie PJ, Banman SL, Brandt C, Stone SJ. Diacylglycerol acyltransferase-2 (DGAT2) and monoacylglycerol acyltransferase-2 (MGAT2) interact to promote triacylglycerol synthesis. J Biol Chem. 2014;289(41):28237–48.

Wilfling F, Wang H, Haas JT, et al. Triacylglycerol synthesis enzymes mediate lipid droplet growth by relocalizing from the ER to lipid droplets. Dev Cell. 2013;24(4):384–99.

Xu N, Zhang SO, Cole RA, et al. The FATP1-DGAT2 complex facilitates lipid droplet expansion at the ER-lipid droplet interface. J Cell Biol. 2012;198(5):895–911.

Becuwe M, Bond LM, Pinto AFM, et al. FIT2 is an acyl-coenzyme A diphosphatase crucial for endoplasmic reticulum homeostasis. J Cell Biol. 2020;219(10):e202006111.

Bond LM, Ibrahim A, Lai ZW, et al. Fitm2 is required for ER homeostasis and normal function of murine liver. J Biol Chem. 2023;299(3):103022.

Chen F, Yan B, Ren J, et al. FIT2 organizes lipid droplet biogenesis with ER tubule-forming proteins and septins. J Cell Biol. 2021;220(5):e201907183.

Qian K, Tol MJ, Wu J, et al. CLSTN3β enforces adipocyte multilocularity to facilitate lipid utilization. Nature. 2023;613(7942):160–8.

Wei X, Li L, Zhao J, et al. BAP31 depletion inhibited adipogenesis, repressed lipolysis and promoted lipid droplets abnormal growth via attenuating Perilipin1 proteasomal degradation. Int J Biol Sci. 2023;19(6):1713–30.

Xu D, Li Y, Wu L, et al. Rab18 promotes lipid droplet (LD) growth by tethering the ER to LDs through SNARE and NRZ interactions. J Cell Biol. 2018;217(3):975–95.

López-Alcalá J, Soler-Vázquez MC, Tercero-Alcázar C, et al. Rab18 Drift in Lipid Droplet and Endoplasmic Reticulum Interactions of Adipocytes under Obesogenic Conditions. Int J Mol Sci. 2023;24(24):17177.

Romanauska A, Köhler A. Lipid saturation controls nuclear envelope function. Nat Cell Biol. 2023;25(9):1290–302.

Palikaras K, Mari M, Ploumi C, Princz A, Filippidis G, Tavernarakis N. Age-dependent nuclear lipid droplet deposition is a cellular hallmark of aging in Caenorhabditis elegans. Aging Cell. 2023;22(4):e13788.

Fujimoto T. Nuclear lipid droplets - how are they different from their cytoplasmic siblings?. J Cell Sci. 2022;135(5):jcs259253.

Fonin AV, Silonov SA, Shpironok OG, et al. The Role of Non-Specific Interactions in Canonical and ALT-Associated PML-Bodies Formation and Dynamics. Int J Mol Sci. 2021;22(11):5821.

Xu W, Wu L, Yu M, et al. Differential Roles of Cell Death-inducing DNA Fragmentation Factor-α-like Effector (CIDE) Proteins in Promoting Lipid Droplet Fusion and Growth in Subpopulations of Hepatocytes. J Biol Chem. 2016;291(9):4282–93.

Gong J, Sun Z, Wu L, et al. Fsp27 promotes lipid droplet growth by lipid exchange and transfer at lipid droplet contact sites. J Cell Biol. 2011;195(6):953–63.

Wang J, Yan C, Xu C, Chua BT, Li P, Chen FJ. Polybasic RKKR motif in the linker region of lipid droplet (LD)-associated protein CIDEC inhibits LD fusion activity by interacting with acidic phospholipids. J Biol Chem. 2018;293(50):19330–43.

Wu L, Xu D, Zhou L, et al. Rab8a-AS160-MSS4 regulatory circuit controls lipid droplet fusion and growth. Dev Cell. 2014;30(4):378–93.

Grahn TH, Zhang Y, Lee MJ, et al. FSP27 and PLIN1 interaction promotes the formation of large lipid droplets in human adipocytes. Biochem Biophys Res Commun. 2013;432(2):296–301.

Kamada R, Uno S, Kimura N, Yoshimura F, Tanino K, Sakaguchi K. Lipid Droplet Formation Is Regulated by Ser/Thr Phosphatase PPM1D via Dephosphorylation of Perilipin 1. Int J Mol Sci. 2022;23(19):12046.

Wozny MR, Di Luca A, Morado DR, et al. In situ architecture of the ER-mitochondria encounter structure. Nature. 2023;618(7963):188–92.

Csordás G, Renken C, Várnai P, et al. Structural and functional features and significance of the physical linkage between ER and mitochondria. J Cell Biol. 2006;174(7):915–21.

Li J, Xin Y, Li J, Chen H, Li H. Phosphatidylethanolamine N-methyltransferase: from Functions to Diseases. Aging Dis. 2023;14(3):879–91.

Hernández-Alvarez MI, Sebastián D, Vives S, et al. Deficient Endoplasmic Reticulum-Mitochondrial Phosphatidylserine Transfer Causes Liver Disease. Cell. 2019;177(4):881-895.e17.

Singh AB, Kan CFK, Kraemer FB, Sobel RA, Liu J. Liver-specific knockdown of long-chain acyl-CoA synthetase 4 reveals its key role in VLDL-TG metabolism and phospholipid synthesis in mice fed a high-fat diet. Am J Physiol Endocrinol Metab. 2019;316(5):E880–94.

Zaman M, Shutt TE. The Role of Impaired Mitochondrial Dynamics in MFN2-Mediated Pathology. Front Cell Dev Biol. 2022;10:858286.

Veliova M, Petcherski A, Liesa M, Shirihai OS. The biology of lipid droplet-bound mitochondria. Semin Cell Dev Biol. 2020;108:55–64.

Rambold AS, Cohen S, Lippincott-Schwartz J. Fatty acid trafficking in starved cells: regulation by lipid droplet lipolysis, autophagy, and mitochondrial fusion dynamics. Dev Cell. 2015;32(6):678–92.

Herms A, Bosch M, Reddy BJ, et al. AMPK activation promotes lipid droplet dispersion on detyrosinated microtubules to increase mitochondrial fatty acid oxidation. Nat Commun. 2015;6:7176.

Khaddaj R, Mari M, Cottier S, Reggiori F, Schneiter R. The surface of lipid droplets constitutes a barrier for endoplasmic reticulum-resident integral membrane proteins. J Cell Sci. 2022;135(5):jcs256206.

Benador IY, Veliova M, Mahdaviani K, et al. Mitochondria Bound to Lipid Droplets Have Unique Bioenergetics, Composition, and Dynamics that Support Lipid Droplet Expansion. Cell Metab. 2018;27(4):869-885.e6.

Wu C, Dai C, Li X, et al. AKR1C3-dependent lipid droplet formation confers hepatocellular carcinoma cell adaptability to targeted therapy. Theranostics. 2022;12(18):7681–98.

Nguyen TB, Louie SM, Daniele JR, et al. DGAT1-Dependent Lipid Droplet Biogenesis Protects Mitochondrial Function during Starvation-Induced Autophagy. Dev Cell. 2017;42(1):9-21.e5.

Stone SJ, Levin MC, Zhou P, Han J, Walther TC, Farese RV Jr. The endoplasmic reticulum enzyme DGAT2 is found in mitochondria-associated membranes and has a mitochondrial targeting signal that promotes its association with mitochondria. J Biol Chem. 2009;284(8):5352–61.

Freyre CAC, Rauher PC, Ejsing CS, Klemm RW. MIGA2 Links Mitochondria, the ER, and Lipid Droplets and Promotes De Novo Lipogenesis in Adipocytes. Mol Cell. 2019;76(5):811-825.e14.

Combot Y, Salo VT, Chadeuf G, et al. Seipin localizes at endoplasmic-reticulum-mitochondria contact sites to control mitochondrial calcium import and metabolism in adipocytes. Cell Rep. 2022;38(2):110213.

Guyard V, Monteiro-Cardoso VF, Omrane M, et al. ORP5 and ORP8 orchestrate lipid droplet biogenesis and maintenance at ER-mitochondria contact sites. J Cell Biol. 2022;221(9):e202112107.

Bosc C, Broin N, Fanjul M, et al. Autophagy regulates fatty acid availability for oxidative phosphorylation through mitochondria-endoplasmic reticulum contact sites. Nat Commun. 2020;11(1):4056.

Papsdorf K, Miklas JW, Hosseini A, et al. Lipid droplets and peroxisomes are co-regulated to drive lifespan extension in response to mono-unsaturated fatty acids [published correction appears in Nat Cell Biol. 2023 Oct;25(10):1547]. Nat Cell Biol. 2023;25(5):672–684.

Kong J, Ji Y, Jeon YG, et al. Spatiotemporal contact between peroxisomes and lipid droplets regulates fasting-induced lipolysis via PEX5. Nat Commun. 2020;11(1):578.

Schrul B, Kopito RR. Peroxin-dependent targeting of a lipid-droplet-destined membrane protein to ER subdomains. Nat Cell Biol. 2016;18(7):740–51.

Chang CL, Weigel AV, Ioannou MS, et al. Spastin tethers lipid droplets to peroxisomes and directs fatty acid trafficking through ESCRT-III. J Cell Biol. 2019;218(8):2583–99.

Ohsaki Y, Cheng J, Fujita A, Tokumoto T, Fujimoto T. Cytoplasmic lipid droplets are sites of convergence of proteasomal and autophagic degradation of apolipoprotein B. Mol Biol Cell. 2006;17(6):2674–83.

Wang CW. Lipid droplets, lipophagy, and beyond. Biochim Biophys Acta. 2016;1861(8 Pt B):793–805.

Omrane M, Ben M’Barek K, Santinho A, et al. LC3B is lipidated to large lipid droplets during prolonged starvation for noncanonical autophagy. Dev Cell. 2023;58(14):1266-1281.e7.

Omrane M, Melia TJ, Thiam AR. LC3 conjugation to lipid droplets. Autophagy. 2023;19(12):3251–3.

Singh R, Kaushik S, Wang Y, et al. Autophagy regulates lipid metabolism. Nature. 2009;458(7242):1131–5.

Kawano M, Kumagai K, Nishijima M, Hanada K. Efficient trafficking of ceramide from the endoplasmic reticulum to the Golgi apparatus requires a VAMP-associated protein-interacting FFAT motif of CERT. J Biol Chem. 2006;281(40):30279–88.

Kumagai K, Kawano-Kawada M, Hanada K. Phosphoregulation of the ceramide transport protein CERT at serine 315 in the interaction with VAMP-associated protein (VAP) for inter-organelle trafficking of ceramide in mammalian cells. J Biol Chem. 2014;289(15):10748–60.

Shimasaki K, Kumagai K, Sakai S, Yamaji T, Hanada K. Hyperosmotic Stress Induces Phosphorylation of CERT and Enhances Its Tethering throughout the Endoplasmic Reticulum. Int J Mol Sci. 2022;23(7):4025.

Prashek J, Bouyain S, Fu M, Li Y, Berkes D, Yao X. Interaction between the PH and START domains of ceramide transfer protein competes with phosphatidylinositol 4-phosphate binding by the PH domain. J Biol Chem. 2017;292(34):14217–28.

Sugiki T, Egawa D, Kumagai K, et al. Phosphoinositide binding by the PH domain in ceramide transfer protein (CERT) is inhibited by hyperphosphorylation of an adjacent serine-repeat motif. J Biol Chem. 2018;293(28):11206–17.

Doyle CP, Timple L, Hammond GRV. OSBP is a Major Determinant of Golgi Phosphatidylinositol 4-Phosphate Homeostasis. Contact (Thousand Oaks). 2024;7:25152564241232196.

He R, Liu F, Wang H, et al. ORP9 and ORP10 form a heterocomplex to transfer phosphatidylinositol 4-phosphate at ER-TGN contact sites. Cell Mol Life Sci. 2023;80(3):77.

Zhou Y, Li S, Mäyränpää MI, et al. OSBP-related protein 11 (ORP11) dimerizes with ORP9 and localizes at the Golgi-late endosome interface. Exp Cell Res. 2010;316(19):3304–16.

Naito T, Yang H, Koh DHZ, Mahajan D, Lu L, Saheki Y. Regulation of cellular cholesterol distribution via non-vesicular lipid transport at ER-Golgi contact sites. Nat Commun. 2023;14(1):5867.

Wakana Y, Hayashi K, Nemoto T, et al. The ER cholesterol sensor SCAP promotes CARTS biogenesis at ER-Golgi membrane contact sites. J Cell Biol. 2021;220(1):e202002150.

Wakana Y, Kotake R, Oyama N, et al. CARTS biogenesis requires VAP-lipid transfer protein complexes functioning at the endoplasmic reticulum-Golgi interface. Mol Biol Cell. 2015;26(25):4686–99.

Sobajima T, Yoshimura SI, Maeda T, Miyata H, Miyoshi E, Harada A. The Rab11-binding protein RELCH/KIAA1468 controls intracellular cholesterol distribution. J Cell Biol. 2018;217(5):1777–96.

Gong B, Guo Y, Ding S, et al. A Golgi-derived vesicle potentiates PtdIns4P to PtdIns3P conversion for endosome fission. Nat Cell Biol. 2021;23(7):782–95.

Kawasaki A, Sakai A, Nakanishi H, et al. PI4P/PS countertransport by ORP10 at ER-endosome membrane contact sites regulates endosome fission. J Cell Biol. 2022;221(1):e202103141.

Meneses-Salas E, García-Melero A, Kanerva K, et al. Annexin A6 modulates TBC1D15/Rab7/StARD3 axis to control endosomal cholesterol export in NPC1 cells. Cell Mol Life Sci. 2020;77(14):2839–57.

Lu A, Hsieh F, Sharma BR, Vaughn SR, Enrich C, Pfeffer SR. CRISPR screens for lipid regulators reveal a role for ER-bound SNX13 in lysosomal cholesterol export. J Cell Biol. 2022;221(2):e202105060.

Saric A, Freeman SA, Williamson CD, et al. SNX19 restricts endolysosome motility through contacts with the endoplasmic reticulum. Nat Commun. 2021;12(1):4552.

Rocha N, Kuijl C, van der Kant R, et al. Cholesterol sensor ORP1L contacts the ER protein VAP to control Rab7-RILP-p150 Glued and late endosome positioning. J Cell Biol. 2009;185(7):1209–25.

Radulovic M, Wenzel EM, Gilani S, et al. Cholesterol transfer via endoplasmic reticulum contacts mediates lysosome damage repair. EMBO J. 2022;41(24):e112677.

Zhao K, Ridgway ND. Oxysterol-Binding Protein-Related Protein 1L Regulates Cholesterol Egress from the Endo-Lysosomal System. Cell Rep. 2017;19(9):1807–18.

Naito T, Saheki Y. GRAMD1-mediated accessible cholesterol sensing and transport. Biochim Biophys Acta Mol Cell Biol Lipids. 2021;1866(8):158957.

Trinh MN, Brown MS, Goldstein JL, et al. Last step in the path of LDL cholesterol from lysosome to plasma membrane to ER is governed by phosphatidylserine. Proc Natl Acad Sci U S A. 2020;117(31):18521–9.

Kattan WE, Liu J, Montufar-Solis D, et al. Components of the phosphatidylserine endoplasmic reticulum to plasma membrane transport mechanism as targets for KRAS inhibition in pancreatic cancer. Proc Natl Acad Sci U S A. 2021;118(51):e2114126118.

Desai R, East DA, Hardy L, et al. Mitochondria form contact sites with the nucleus to couple prosurvival retrograde response. Sci Adv. 2020;6(51):eabc9955.

Höglinger D, Burgoyne T, Sanchez-Heras E, et al. NPC1 regulates ER contacts with endocytic organelles to mediate cholesterol egress. Nat Commun. 2019;10(1):4276.

Shuwen H, Yinhang W, Jing Z, et al. Cholesterol induction in CD8+ T cell exhaustion in colorectal cancer via the regulation of endoplasmic reticulum-mitochondria contact sites. Cancer Immunol Immunother. 2023;72(12):4441–56.

Correa-Medero LO, Jankowski SE, Hong HS, et al. ER-associated degradation adapter Sel1L is required for CD8+ T cell function and memory formation following acute viral infection. Cell Rep. 2024;43(5):114156.

Wang R, He S, Long J, et al. Emerging therapeutic frontiers in cancer: insights into posttranslational modifications of PD-1/PD-L1 and regulatory pathways. Exp Hematol Oncol. 2024;13(1):46.

Long J, Wang Y, Jiang X, et al. Nanomaterials Boost CAR-T Therapy for Solid Tumors. Adv Healthc Mater. 2024;13(20):e2304615.

Tello-Lafoz M, Ghai R, Collins B, Mérida I. A role for novel lipid interactions in the dynamic recruitment of SNX27 to the T-cell immune synapse. BioArchitecture. 2014;4(6):215–20.

Wang J, Wang Y, Jiang X, et al. Unleashing the power of immune checkpoints: Post-translational modification of novel molecules and clinical applications. Cancer Lett. 2024;588:216758.

Cabrera-Reyes F, Parra-Ruiz C, Yuseff MI, Zanlungo S. Alterations in Lysosome Homeostasis in Lipid-Related Disorders: Impact on Metabolic Tissues and Immune Cells. Front Cell Dev Biol. 2021;9:790568.

Winer DA, Winer S, Shen L, et al. B cells promote insulin resistance through modulation of T cells and production of pathogenic IgG antibodies. Nat Med. 2011;17(5):610–7.

Tsiantoulas D, Diehl CJ, Witztum JL, Binder CJ. B cells and humoral immunity in atherosclerosis. Circ Res. 2014;114(11):1743–56.

Shaikh SR, Haas KM, Beck MA, Teague H. The effects of diet-induced obesity on B cell function. Clin Exp Immunol. 2015;179(1):90–9.

Veglia F, Tyurin VA, Mohammadyani D, et al. Lipid bodies containing oxidatively truncated lipids block antigen cross-presentation by dendritic cells in cancer. Nat Commun. 2017;8(1):2122.

Criado Santos N, Bouvet S, Cruz Cobo M, et al. Sec22b regulates phagosome maturation by promoting ORP8-mediated lipid exchange at endoplasmic reticulum-phagosome contact sites. Commun Biol. 2023;6(1):1008.

Niu Z, Shi Q, Zhang W, et al. Caspase-1 cleaves PPARγ for potentiating the pro-tumor action of TAMs. Nat Commun. 2017;8(1):766.

Clément S, Fauvelle C, Branche E, et al. Role of seipin in lipid droplet morphology and hepatitis C virus life cycle. J Gen Virol. 2013;94(Pt 10):2208–14.

Ajjaji D, Ben M'barek K, Boson B, et al. Hepatitis C virus core protein uses triacylglycerols to fold onto the endoplasmic reticulum membrane. Traffic. 2022;23(1):63–80.

Yue M, Hu B, Li J, et al. Coronaviral ORF6 protein mediates inter-organelle contacts and modulates host cell lipid flux for virus production. EMBO J. 2023;42(13):e112542.

Grespi F, Vianello C, Cagnin S, Giacomello M, De Mario A. The Interplay of Microtubules with Mitochondria-ER Contact Sites (MERCs) in Glioblastoma. Biomolecules. 2022;12(4):567.

Bassoy EY, Kasahara A, Chiusolo V, et al. ER-mitochondria contacts control surface glycan expression and sensitivity to killer lymphocytes in glioma stem-like cells. EMBO J. 2017;36(11):1493–512.

Fan F, Zhang H, Dai Z, et al. A comprehensive prognostic signature for glioblastoma patients based on transcriptomics and single cell sequencing. Cell Oncol (Dordr). 2021;44(4):917–35.

Lin S, Cao C, Meng Y, et al. Comprehensive analysis of the value of RAB family genes in prognosis of breast invasive carcinoma. Biosci Rep. 2020;40(5):BSR20201103.

Yang K, Wang Z. Rab18 interacted with V-set and immunoglobulin domain-containing 4 (VSIG4) to involve in the apoptosis of glioma and the sensitivity to temozolomide. Bioengineered. 2021;12(1):1391–402.

Kattan WE, Chen W, Ma X, et al. Targeting plasma membrane phosphatidylserine content to inhibit oncogenic KRAS function. Life Sci Alliance. 2019;2(5):e201900431.

Nardi F, Franco OE, Fitchev P, et al. DGAT1 Inhibitor Suppresses Prostate Tumor Growth and Migration by Regulating Intracellular Lipids and Non-Centrosomal MTOC Protein GM130. Sci Rep. 2019;9(1):3035.

Wu H, Han Y, Rodriguez Sillke Y, et al. Lipid droplet-dependent fatty acid metabolism controls the immune suppressive phenotype of tumor-associated macrophages. EMBO Mol Med. 2019;11(11):e10698.

Ackerman D, Tumanov S, Qiu B, et al. Triglycerides Promote Lipid Homeostasis during Hypoxic Stress by Balancing Fatty Acid Saturation. Cell Rep. 2018;24(10):2596-2605.e5.

Deng B, Kong W, Shen X, et al. The role of DGAT1 and DGAT2 in regulating tumor cell growth and their potential clinical implications. J Transl Med. 2024;22(1):290.

Zhou J, Simon JM, Liao C, et al. An oncogenic JMJD6-DGAT1 axis tunes the epigenetic regulation of lipid droplet formation in clear cell renal cell carcinoma. Mol Cell. 2022;82(16):3030-3044.e8.

Loomba R, Morgan E, Watts L, et al. Novel antisense inhibition of diacylglycerol O-acyltransferase 2 for treatment of non-alcoholic fatty liver disease: a multicentre, double-blind, randomised, placebo-controlled phase 2 trial. Lancet Gastroenterol Hepatol. 2020;5(9):829–38.

Amin NB, Saxena AR, Somayaji V, Dullea R. Inhibition of Diacylglycerol Acyltransferase 2 Versus Diacylglycerol Acyltransferase 1: Potential Therapeutic Implications of Pharmacology. Clin Ther. 2023;45(1):55–70.

Amin NB, Darekar A, Anstee QM, et al. Efficacy and safety of an orally administered DGAT2 inhibitor alone or coadministered with a liver-targeted ACC inhibitor in adults with non-alcoholic steatohepatitis (NASH): rationale and design of the phase II, dose-ranging, dose-finding, randomised, placebo-controlled MIRNA (Metabolic Interventions to Resolve NASH with fibrosis) study. BMJ Open. 2022;12(3):e056159.

Nisticò C, Chiarella E. An Overview on Lipid Droplets Accumulation as Novel Target for Acute Myeloid Leukemia Therapy. Biomedicines. 2023;11(12):3186.

Li Y, Li T, Jin Y, Shen J. Dgat2 reduces hepatocellular carcinoma malignancy via downregulation of cell cycle-related gene expression. Biomed Pharmacother. 2019;115:108950.

Zhang M, Di Martino JS, Bowman RL, et al. Adipocyte-Derived Lipids Mediate Melanoma Progression via FATP Proteins. Cancer Discov. 2018;8(8):1006–25.

Peeters R, Cuenca-Escalona J, Zaal EA, et al. Fatty acid metabolism in aggressive B-cell lymphoma is inhibited by tetraspanin CD37. Nat Commun. 2022;13(1):5371.

Gudgeon N, Giles H, Bishop EL, et al. Uptake of long-chain fatty acids from the bone marrow suppresses CD8+ T-cell metabolism and function in multiple myeloma. Blood Adv. 2023;7(20):6035–47.

Liu Z, Gao Z, Li B, et al. Lipid-associated macrophages in the tumor-adipose microenvironment facilitate breast cancer progression. Oncoimmunology. 2022;11(1):2085432.

Nishiyama K, Fujita T, Fujimoto Y, Nakajima H, Takeuchi T, Azuma YT. Fatty acid transport protein 1 enhances the macrophage inflammatory response by coupling with ceramide and c-Jun N-terminal kinase signaling. Int Immunopharmacol. 2018;55:205–15.

Ye G, Gao H, Wang Z, et al. PPARα and PPARγ activation attenuates total free fatty acid and triglyceride accumulation in macrophages via the inhibition of Fatp1 expression. Cell Death Dis. 2019;10(2):39.

Viani P, Giussani P, Brioschi L, et al. Ceramide in nitric oxide inhibition of glioma cell growth. Evidence for the involvement of ceramide traffic. J Biol Chem. 2003;278(11):9592–9601.

Giussani P, Colleoni T, Brioschi L, et al. Ceramide traffic in C6 glioma cells: evidence for CERT-dependent and independent transport from ER to the Golgi apparatus. Biochim Biophys Acta. 2008;1781(1–2):40–51.

Fleury L, Faux C, Santos C, Ballereau S, Génisson Y, Ausseil F. Development of a CERT START Domain-Ceramide HTRF Binding Assay and Application to Pharmacological Studies and Screening. J Biomol Screen. 2015;20(6):779–87.

Hullin-Matsuda F, Tomishige N, Sakai S, et al. Limonoid compounds inhibit sphingomyelin biosynthesis by preventing CERT protein-dependent extraction of ceramides from the endoplasmic reticulum. J Biol Chem. 2012;287(29):24397–411.

Cruz ALS, Barreto EA, Fazolini NPB, Viola JPB, Bozza PT. Lipid droplets: platforms with multiple functions in cancer hallmarks. Cell Death Dis. 2020;11(2):105.

Jin Y, Tan Y, Wu J, Ren Z. Lipid droplets: a cellular organelle vital in cancer cells. Cell Death Discov. 2023;9(1):254.

Santos FM, Rosa JN, Candeias NR, et al. A Three-Component Assembly Promoted by Boronic Acids Delivers a Modular Fluorophore Platform (BASHY Dyes). Chemistry. 2016;22(5):1631–7.

Klymchenko AS. Fluorescent Probes for Lipid Membranes: From the Cell Surface to Organelles. Acc Chem Res. 2023;56(1):1–12.

Liu G, Zheng H, Zhou R, Ultrabright organic fluorescent probe for quantifying the dynamics of cytosolic, nuclear lipid droplets [published correction appears in Biosens Bioelectron., et al. 15;246:115908]. Biosens Bioelectron. 2024;2023(241):115707.

Yudhistira T, Da Silva EC, Combes A, Lehmann M, Reisch A, Klymchenko AS. Biotinylated Fluorescent Polymeric Nanoparticles for Enhanced Immunostaining. Small Methods. 2023;7(4):e2201452.

Zhang C, Li J, Lan L, Cheng JX. Quantification of Lipid Metabolism in Living Cells through the Dynamics of Lipid Droplets Measured by Stimulated Raman Scattering Imaging. Anal Chem. 2017;89(8):4502–7.

Yao K, Iwashita A, Tanabe H, et al. White opaque substance within superficial elevated gastric neoplasia as visualized by magnification endoscopy with narrow-band imaging: a new optical sign for differentiating between adenoma and carcinoma. Gastrointest Endosc. 2008;68(3):574–80.

Zhang I, Cui Y, Amiri A, Ding Y, Campbell RE, Maysinger D. Pharmacological inhibition of lipid droplet formation enhances the effectiveness of curcumin in glioblastoma. Eur J Pharm Biopharm. 2016;100:66–76.

Antunes P, Cruz A, Barbosa J, Bonifácio VDB, Pinto SN. Lipid Droplets in Cancer: From Composition and Role to Imaging and Therapeutics. Molecules. 2022;27(3):991.

Jiang G, Li C, Liu X, Chen Q, Li X, Gu X, Zhang P, Lai Q, Wang J. Lipid droplet-targetable fluorescence guided photodynamic therapy of cancer cells with an activatable AIE-active fluorescent probe for hydrogen peroxide. Adv Opt Mater. 2020;8:2001119.

Xia X, Wang R, Hu Y, et al. A Novel Photosensitizer for Lipid Droplet-Location Photodynamic Therapy. Front Chem. 2021;9:701771.

Sun Z, Shi S, Guan P, Liu B. Construction of heteroaryl-bridged NIR AIEgens for specific imaging of lipid droplets and its application in photodynamic therapy. Spectrochim Acta A Mol Biomol Spectrosc. 2022;272:120946.

Tan P , Zhuang W , Li S , et al. A lipid droplet targeted fluorescent probe for high-efficiency image-guided photodynamic therapy of renal cell carcinoma [published correction appears in Chem Commun (Camb). 2021 Jan 22;:]. Chem Commun (Camb). 2021;57(8):1046–1049.

Dai Y, Zhao X, Ji H, Zhang D, Zhang P, Xue K, Misal S, Zhu H, Qi Z. Multifunctional aggregation-induced emission nanoparticle for high-fidelity imaging of lipid droplets in living cells and its application in photodynamic therapy. Chem Eng J. 2021;410:128186.

Wang SH, Zhang YW, Wang XD, Zan Q, Yu X, Fan L. An esterase-sensitive AIEgen probe targeting mitochondria and lipid droplets for assessing cell viability. Spectrochim Acta A Mol Biomol Spectrosc. 2023;287(Pt 1):122122.

Zhang C, Shao H, Zhang J, et al. Long-term live-cell lipid droplet-targeted biosensor development for nanoscopic tracking of lipid droplet-mitochondria contact sites. Theranostics. 2021;11(16):7767–78.

Alistar A, Morris BB, Desnoyer R, et al. Safety and tolerability of the first-in-class agent CPI-613 in combination with modified FOLFIRINOX in patients with metastatic pancreatic cancer: a single-centre, open-label, dose-escalation, phase 1 trial. Lancet Oncol. 2017;18(6):770–8.

Pardee TS, Anderson RG, Pladna KM, et al. A Phase I Study of CPI-613 in Combination with High-Dose Cytarabine and Mitoxantrone for Relapsed or Refractory Acute Myeloid Leukemia. Clin Cancer Res. 2018;24(9):2060–73.

Anderson R, Miller LD, Isom S, et al. Phase II trial of cytarabine and mitoxantrone with devimistat in acute myeloid leukemia. Nat Commun. 2022;13(1):1673.

Mohan A, Griffith KA, Wuchu F, et al. Devimistat in Combination with Gemcitabine and Cisplatin in Biliary Tract Cancer: Preclinical Evaluation and Phase Ib Multicenter Clinical Trial (BilT-04). Clin Cancer Res. 2023;29(13):2394–400.

Lycan TW, Pardee TS, Petty WJ, et al. A Phase II Clinical Trial of CPI-613 in Patients with Relapsed or Refractory Small Cell Lung Carcinoma. PLoS One. 2016;11(10):e0164244.

Wang M, Zhang M, Hu X, et al. Lipid-functionalized gold nanorods with plug-to-direct mitochondria targeting ligand for synergetic photothermal-chemotherapy of tumor therapy. Eur J Pharm Biopharm. 2023;185:71–81.

Pawar A, Korake S, Pawar A, Kamble R. Delocalized Lipophilic Cation Triphenyl Phosphonium: Promising Molecule for Mitochondria Targeting. Curr Drug Deliv. 2023;20(9):1217–23.

Nguyen Cao TG, Truong Hoang Q, Kang JH, et al. Bioreducible exosomes encapsulating glycolysis inhibitors potentiate mitochondria-targeted sonodynamic cancer therapy via cancer-targeted drug release and cellular energy depletion. Biomaterials. 2023;301:122242.

Piao S, Amaravadi RK. Targeting the lysosome in cancer. Ann N Y Acad Sci. 2016;1371(1):45–54.