The diversity and breadth of cancer cell fatty acid metabolism

The extracellular pool of fatty acids consists of several sources. They include those in the plasma: adipocyte-derived, albumin-bound free fatty acids (or non-esterified fatty acids), and those contained in lipoprotein triacylglycerols and/or fatty acid esters (i.e., cholesteryl esters) and glycerophospholipids. These lipoprotein-contained fatty acids can be liberated by the actions of extracellular lipases, including lipoprotein lipase (LPL) and secreted phospholipase A₂ [42]. Likewise, there are stromal supplies which include local adipocytes, cancer-associated fibroblast-derived extracellular vesicles [43], and may consist of autophagy-lipophagy of stromal cells (analogous to the transfer of cancer-associated fibroblast-derived amino acids, etc. [44]). Finally, extracellular lysophospholipids can be taken up by cells; however, the mechanism by which these lipids cross the plasma membrane remains to be defined [45].

In general, extracellular sources of fatty acids are taken up by cells via two mechanisms:

To date, few studies have assessed the rate of long-chain free fatty uptake in cancer cells. One notable study reported that malignant prostate cancer tissue had higher fatty acid uptake rates compared to patient-matched benign tissue [52]. Interestingly, the same authors reported that near-complete ablation of SR-B2/CD36 mRNA reduced free fatty acid uptake by only 35% in PC-3 prostate cancer cells. This is consistent with knockdown of SR-B2/CD36 in SKOV3ip1 ovarian cancer cells which attenuated fatty acid uptake by ~ 40% [53]. In vivo, ablation of SR-B2/CD36 in the prostate tissue of Pten-deficient mice reduced fatty acid uptake by ~ 55%, while treating mice harboring PDXs of localized high-risk prostate cancer with a SR-B2/CD36 mAb reduced fatty acid uptake by 22% [52]. These studies demonstrate that SR-B2/CD36 plays a role in extracellular fatty acid uptake in cancer cells, but other mechanisms contribute to this process.

Despite this quantitatively “minor” role in fatty acid uptake, recent loss-of-function studies have clearly shown that SR-B2/CD36 is critical in prostate [52], ovarian [53], oral [54], cervical [55], gastric [56], breast [57, 58], and liver [59] cancer biology. For example, prostate-specific deletion of SR-B2/CD36 of cancer-susceptible Pten^−/− mice slowed cancer progression, while SR-B2/CD36 antibody therapy reduced cancer severity in patient-derived xenografts [52]. Likewise, SR-B2/CD36⁺ human oral carcinoma cells initiate metastasis in an SR-B2/CD36-dependent manner, and SR-B2/CD36 antibody therapy inhibited metastasis in preclinical models [54]. An array of signaling pathways that mediate these patterns have been proposed (see review [60]); however, the precise mechanism by which the reduced uptake of extracellular fatty acids by SR-B2/CD36 influences cancer cell biology remains a mystery.

Compared to SR-B2/CD36, the other proposed regulators of fatty acid uptake, such as FATPs and FABP plasma membrane [46], have received far less attention. This is likely due to what appears to be a widely held belief that SR-B2/CD36 is rate-limiting in fatty acid uptake or the only fatty acid transporter. Recently, Zhang and colleagues demonstrated that FATP1, which is overexpressed in melanoma, is required for fatty acid uptake and melanoma growth and invasion in vitro and in vivo [61]. What role the other five members of the FATP family play in cancer biology remains an area of opportunity.

The contribution of lipoprotein particles to the intracellular fatty acid pool remains poorly defined within cell biology. One key reason could be that triacylglycerol-rich chylomicrons and VLDL can provide fatty acids via at least two distinct pathways—extracellular lipolysis to liberate free fatty acids for uptake, or endocytosis [62]. Despite this, it is clear that cells, including U87-MG glioblastoma cells and MDA-MB-231 breast cancer cells, increase intracellular lipid levels and VLDL uptake in a time-dependent and dose-dependent manner [63, 64]. Triacylglycerol-rich VLDL can be endocytosed by VLDLr and protein levels of VLDLr are increased in hepatocellular carcinoma [65] and clear-cell renal cell carcinoma [66] compared to adjacent non-cancerous tissue. Knockdown of VLDLr reduced lipoprotein uptake and intracellular lipid levels in clear-cell RCC cells [66], and in MCF-7 and MDA-MB-231 breast cancer cells [64, 65]. To further complicate our understanding, it was recently demonstrated that VLDLr-mediated VLDL uptake requires LPL acting non-catalytically to facilitate endocytosis [64]. Triacylglycerol-rich lipoproteins are also metabolized by the cell surface protein LSR, which is highly expressed in breast cancer [67]. However, LSR’s role in breast cancer metabolism is complicated as it also regulates tight junctions and was recently identified to be capable of nuclear localization and DNA binding [67]. Likewise, LPR1, which is involved in lipoprotein transport, ligand uptake, and receptor-mediated endocytosis, also regulates cell surface protease activity and acts on many cell signaling pathways, and so its role in lipid-mediated changes in cancer biology is very complex [68]. Finally, the role of LDLr and its related proteins in cancer biology has rightly centered on its role in cholesterol homeostasis, and not fatty acid metabolism; however, it is conceivable that the glycerophospholipids and the fatty acid of cholesteryl esters contribute to intracellular fatty acid levels. To date, the contribution of this pathway to the intracellular fatty acid pool remains unknown.

A likely alternative pathway for the accumulation of extracellular fatty acid-based lipids is macropinocytosis. While this is a known mechanism for cancer cells to acquire extracellular proteins which then are processed by the endosomal-lysosomal pathway (see review [69]), it is conceivable, and has been hypothesized [69, 70], that lipids, including fatty acid-based lipids, are also endocytosed and contribute to the intracellular fatty acid pool. In fact, supplying mouse mPCE or human DU145 prostate cancer cells necrotic cell debris in glucose- and amino acid-restricted media completely restored lipid droplet content, suggesting membranes and lipids present in necrotic debris can maintain lipid stores [71].

The synthesis of new long-chain fatty acids from non-lipid substrates is another input to the intracellular free fatty acid pool. We will focus on the cytosolic pathway but acknowledge that mitochondria are capable of synthesizing predominantly short- and medium-chain fatty acids that can act as precursors for lipoic acid synthesis and protein lipoylation [72, 73]. The cytosolic synthesis of fatty acids starts with the export of mitochondrial citrate into the cytosol via the mitochondrial tricarboxylate transporter (encoded by SLC25A1) where it is converted into acetyl-CoA by ATP-citrate lyase (encoded by ACLY; Fig. 1). Extra-mitochondrial acetyl-CoA is also generated by acyl-coenzyme A synthetase short-chain family member 2 (ACSS2) [74] from acetate, which itself can be derived from a range of sources including extracellular acetate and the recently identified conversion of pyruvate into acetate by thiamine-dependent keto acid dehydrogenases as well as a ROS-coupled reaction [75].

Acetyl-CoA is converted to malonyl-CoA by acetyl-CoA carboxylase (ACC, encoded by ACACA and ACACB), with both acetyl-CoA (1 molecule) and malonyl-CoA (7 molecules) used by fatty acid synthase (FAS, encoded by FASN) to produce the 16 carbon saturated fatty acid palmitate. The production of one palmitate molecule requires 7 ATP and 14 molecules of NADPH and the molecular regulation of de novo fatty acid synthesis pathway has been comprehensively reviewed [76]. It should be noted that cytosolic acetyl-CoA is also a substrate for cholesterol synthesis (in fact, all 27 carbons in cholesterol are derived from acetyl-CoA [77]), and protein acetylation [78]. In contrast, malonyl-CoA is also a substrate for fatty acid elongation (see below).

Increased de novo fatty acid synthesis is a commonly observed feature of cancer cells [79, 80], and the enzymes ACLY, ACC, and FAS have been demonstrated as potential therapeutic targets. Recent examples include knockdown of ACLY impairing pancreatic tumor formation [81] and knockdown of FASN blocking tumor development in mTOR-driven liver cancer [82], while pharmacological inhibition of FAS reduced tumor growth in preclinical models of castration-resistant prostate cancer [83]. Similar observations have been reported in breast [84] and colon [85] cancer. As such, this is an area of ongoing drug development including the recently developed and characterized selective, irreversible, and potent FASN inhibitor IPI-9119 [83], as well as the recent report of a new mechanism to inhibit human ACLY [86].

De novo fatty acid synthesis has received significant attention; however, first principle questions remain. For example, why is the palmitate produced by de novo fatty acid synthesis required for cell viability? It is widely held that the increase in de novo production of palmitate is to meet demand for membrane synthesis of highly proliferative cancer cells. If this is so, one would assume that de novo synthesis of fatty acids is a greater contributor to bulk lipid synthesis than other pools. We and others have quantified the relative contribution of extracellular fatty acids and de novo synthesis of fatty acids to the cellular lipid pools in cancer cells. De novo synthesis of fatty acids from extracellular glucose contributes ~ 20–30% of cellular lipids, whereas glutamine contributes ~ 5% in H1299 and A549 non-small cell lung cancer cell lines [87], MCF-7 and MDA-231 cells [5] and a range of prostate cancer cell lines [4]; with the remainder (~ 65–75%) coming from extracellular fatty acids. These reports compliment observations made by the Nomura laboratory that showed that de novo synthesized palmitate, generated using extracellular isotopically-labelled glucose, are incorporated into a broad range of lipids including glycerophospholipids, glycerolipids, and sphingolipids but account for only a small fraction of the total levels of palmitate-containing lipids [88]. Specifically, the [¹³C]C16:0 FFA (m+16) pool, which represents newly synthesized palmitate from ¹³C-labelled glucose (incubated for 4 h), accounted for only ≤ 1.9% of the total free palmitate pool in five different cancer cell lines. Likewise, [¹³C]C16:0 was only a minor fraction of the total pool of other lipid pools; [¹³C]lysophosphatidate (m+19) was up to 14% of the total C16:0 lysophosphatidate pool, with similar patterns observed in the C16:0/C18:1 phosphatidate pool, C16:0/C18:1 diacylglycerol pool, C16:0/C18:1 phosphatidylserine, C16:0/C18:1 lysophosphatidylcholine, and C16:0/C18:1 lysophosphatidylethanolamine [88]. In general, de novo synthesized palmitate was not the majority source of C16:0 acyl chains for the broad range of lipids that were measured. Critically, these observations were made using serum-free conditions, and so likely represent the maximal contribution of de novo synthesized fatty acids to membrane synthesis as there were no competing extracellular lipids. The Rabinowitz laboratory also assessed the contribution of de novo palmitate synthesis in cells cultured in 25 mM ¹³C-labelled glucose, 4 mM ¹³C-labelled glutamine, and 10% dialyzed FBS for greater than five doublings (compared to 10 mM glucose, serum-free, 4 h ¹³C-glucose incubation for [88]). They reported that de novo synthesized palmitate generated in cells cultured in labeled glucose and labeled glutamine accounted for ~ 75–90% of the total cellular C16:0 pool, including fatty acyl chains from complex lipids [45]. The authors also reported the incorporation of these labeled non-lipid substrates into C18:0 and C18:1, including m+16 and m+18 isotopologues, indicating that a fraction of de novo synthesized palmitate is modified before being incorporation into lipids (discussed in greater detail in the “Modification of free fatty acids: fatty acid elongation” section). Overall, the majority of reports demonstrate that extracellular fatty acids contribute to the building of lipids to a greater extent than non-lipid substrates in cell culture. This pattern may or may not occur in vivo as it remains unclear what the fatty acid/lipid levels are in the tumor microenvironment, which themselves almost certainly differ in the primary tumor and metastatic tissues as well as at different sites of metastasis.

As de novo synthesized palmitate is not the major source for glycerophospholipid synthesis, it remains unclear why the activity of enzymes that produce palmitate de novo is critical for cell viability. It is conceivable that de novo synthesized palmitate, or its subsequently modified (i.e., elongated, desaturated) variants, are partitioned into specific lipids that are essential for cellular functions. This concept is supported by the observation that FAS inhibition sensitivity correlated with the incorporation of de novo-synthesized palmitate into (C16:0)lysophosphatidate, (C16:0/C18:1)DG, (C16:0/C18:1)PC, and (C16:0)LPC, rather than the free palmitate pool [88]. This suggests that the cell viability in FAS inhibitor sensitive cells is dependent upon the production of specific complex lipids. However, it remains to be determined whether this is also the case for other glycerophospholipids that incorporate elongated and/or desaturated de novo-synthesized palmitate. It is important to note that the inhibition of de novo fatty synthesis is robust primarily in tissue culture conditions where extracellular lipids are depleted, including low serum conditions, serum-free, or delipidated FBS [88‐90]. The ever-growing use of lipidomic analyses, in combination with stable isotopes, are likely to provide greater insight into membrane and other lipid pool composition and probe the biological function(s) of de novo-synthesized fatty acids.

One of the other aspects of palmitate metabolism that remains to be resolved, especially in terms of its requirement for cell viability, is the fact that palmitate supplementation of cell culture media leads to lipotoxicity and activation of apoptosis. This is consistently observed in a broad range of cell lines, including 3T3 fibroblasts [91], peripheral blood mononuclear cells [92], macrophages [93], and hepatocytes [94], as well as cancer cells lines [3, 4, 92, 94‐101]. We recently demonstrated that the higher rates of fatty acid oxidation in C4-2B prostate cancer cells and MCF-7 breast cancer cells protect from palmitate-induced apoptosis, and inhibition of mitochondrial fatty acid oxidation sensitized these cells and lead to increased cell death [3, 4]. Palmitate induced apoptosis in PC-3 prostate cancer cells and MDA-MB-231 breast cancer cells was prevented by pre-treatment of these cells with FAs (oleate or oleate:palmitate:linoleate mix), and this protective effect required DGAT-1–mediated triacylglycerol synthesis. More recently, palmitate-induced apoptosis was reported to require endoplasmic reticulum glycerol-3-phosphate acyltransferase activity and the formation of di-saturated glycerolipids [102]. Collectively, these observations point toward a scenario whereby intracellular palmitate levels, influenced by intracellular and extracellular sources, are tightly controlled and that insufficient or too much results in cell death.

While we have not discussed in great detail the role of de novo fatty acid synthesis in cancer beyond the relative contribution of palmitate to lipid synthesis, we believe that relative to other areas of tumor fatty acid metabolism, our understanding of this pathway in oncogenesis has not dramatically advanced since the excellent review of this pathway by Röhrig and Schulze [76]. As such, we have prioritized other facets of fatty acid metabolism and their emerging roles that have been reported in recent years.

Complex membrane lipids are also an input source for the intracellular fatty acid pool. Membranes are not a static cellular structure but constantly undergo remodeling via the removal of fatty acids and the addition of fatty acyl-CoAs to glycerophospholipids via the competing actions of phospholipases and acyltransferases (Fig. 2). Phospholipases vary in their structure and function with the acylhydrolases PLA₁, PLA₂, and PLB enzymes liberating free fatty acids from specific sites of the phospholipid (i.e., sn-1 or sn-2) producing lysophospholipids (also called monoacylglycerophospholipids [103])—see the review by Harayama and Riezman [26] for details of the chemical diversity of membrane lipids. Other family members include PLC and PLD that hydrolyze glycerophospholipids, but target head group phosphates and so do not liberate a fatty acid. The H-RAS-like suppressor (HRASLS) subfamily all possess in vitro PLA₁ and PLA₂ activities (producing fatty acids) as well as O-acyltransferase activities to remodel glycerophospholipid acyl chains [104]. Likewise, lysophospholipids can also be hydrolyzed to produce glycerophosphate and a fatty acid by the actions of the cytosolic serine hydrolases lysophospholipase A1 (LYPLA1) and lysophospholipase A2 (LYPLA2) [105]. Interestingly, LYPLAs also exhibit protein palmitoyl thioesterase (i.e., depalmitoylation) activity to produce palmitate, with targets including oncogenes HRAS and SRC [106].

A number of studies have explored the role of PLA₁, PLA₂, and PLB family of enzymes in cancer cell biology, with most centered on PLA₂-mediated release of arachidonic acid, which is then used as a substrate for eicosanoid synthesis [107, 108]. That said, the contribution of liberated fatty acids other than arachidonic acid to the free fatty acid pool is not described. In general, PLA₂ activity, including HRASLS expression, is lower in breast, ovarian, and other cancer cells compared to respective normal cells [104, 109]. Conversely, LYPLA1 plays a tumor-promotor role in non-small cell lung cancer cells [110], which is unlikely to be linked to the liberation of fatty acids from lysophospholipids contributing to the free fatty acid pool, but through changes in lysophospholipid levels which can regulate several signaling pathways including MAPK and ERK [105] or via depalmitoylation of the α-subunit of G-proteins and proto-oncogene H-Ras products [111].

Similar to membrane glycerophospholipids, sphingolipids can be hydrolyzed to release fatty acids. As an example, ceramides are substrates for the ceramidase family of enzymes, which produce a free fatty acid and a sphingosine molecule (Fig. 2) [112]. Ceramidases are classified by their optimal pH for catalytic activity, i.e., acid, neutral, alkaline. A recent review highlighted that acid ceramidases are commonly overexpressed in a range of cancer types [113]. Further, a role for neutral ceramidase in colon cancer biology has been demonstrated [114]; however, the role of the fatty acid that is liberated by ceramidases has not been investigated. This is likely due to the fact that this reaction also produces sphingosine which can be phosphorylated by sphingosine kinases to form sphingosine-1-phosphate, and can activate sphingosine-1-phosphate receptors to influence cancer cell biology (see review [115]).

Another intracellular source of fatty acids is neutral lipids stored in cytosolic lipid droplets, including triacylglycerols (TGs; 3 fatty acids attached to a glycerol backbone), sterol esters (1 fatty acid attached to 1 sterol), and 1-O-acylceramides (1 fatty acid attached to 1 ceramide) [116, 117]. The mobilization of lipid droplet-derived fatty acids occurs via the actions of cytosolic neutral lipases or lipophagy [118] (Fig. 3). The molecular regulation of lipolysis of lipid droplet-contained TG is complex and involves a combination of subcellular localization, post-translational modification (in particular phosphorylation), and protein-protein interaction [118]. In contrast, our understanding of the regulatory mechanisms of lipophagy and hydrolysis of sterol esters and 1-O-acylceramides remains underdeveloped.

Lipid droplet accumulation has been reported in a broad range of human cancers (see review [119]), in hypoxic cancer cells [41], and several studies have linked lipid droplet accumulation with more aggressive disease [119‐121]. In fact, lipid droplets are key managers of the storage and release of unsaturated fatty acids and play an important antioxidant role and protect cancer cells from stress associated with both nutrient excess and nutrient deprivation [42, 122]. As such, they have been proposed to play direct roles in many of the Hallmarks of Cancer (see review [123]).

The first step is TG hydrolysis, which releases one fatty acid and produces a diacylglycerol (DG). Several enzymes express TG hydrolase activity, including the adipose triglyceride lipase (ATGL, encoded by PNPLA2) and hormone-sensitive lipase (HSL, encoded by LIPE), as well as carboxylesterase 2 (CES2), alongside its well-described role in drug metabolism [124], and patatin-like phospholipase domain-containing protein 3 (PNPLA3), in human liver [125], but it remains unclear whether CES2 and PNPLA3 participate in TG hydrolysis in cancer cells.

ATGL targets the sn-2 position of TG to produce an sn-1,3 DG, that does not activate protein kinase C signaling (see below). There are many conflicting observations reported regarding the role of ATGL in cancer biology; note that this inconsistency in the role of ATGL is also observed in fatty liver [126, 127]. Some studies have shown that ATGL loss-of-function reduces cancer cell proliferation and invasion (see review [128]), whereas others have reported increased lung cancer cell proliferation and migration [129], or no change in colorectal, melanoma, lung, and liver cancer cell proliferation or in vivo tumor size [130, 131]. ATGL overexpression did suppress melanoma, lung, and liver cancer cell proliferation [131]. We and others have shown that ATGL protein levels, and TG levels, are increased in colon, breast, and prostate cancer cells in response to high levels of extracellular fatty acids [3, 4, 132] and co-culturing with adipocytes [133], which also increase cell proliferation. Importantly, we also showed that the increase in ATGL protein and intracellular TG levels increased the rate of mitochondrial oxidation of TG-derived fatty acids in breast and prostate cancer cells [3, 4]. This suggests that ATGL likely provides fatty acids that are key for mechanisms that support cell proliferation and invasion, and that tumors growing in lipid-rich environments (such as obesity) have enhanced fatty acid flux and thereby increased cell proliferation and invasion rates.

Complicating our understanding of the role of ATGL catalyzed TG hydrolysis in cancer cell biology is the fact that the activity of ATGL is regulated by protein-protein interactions. To date, ABHD5 (also known as CGI-58) is the only identified co-activator of ATGL [134]. Knockdown of ABHD5 reduced the rate of TG hydrolysis and increased TG level as expected in prostate cancer cells, but one study reported activated apoptotic signaling [135], whereas another showed induced epithelial-mesenchymal transition, leading to increased cell invasion and proliferation [136]. Interestingly, a recent study showed that cancer cell TG hydrolase activity in vitro is not activated by ABHD5 [131], while others have reported that ABHD5 possesses lysophosphatidate acyl transferase (LPAAT) activity, converting lysophosphatidate into phosphatidate [137]. As such, the influence of ABHD5 on cancer cell biology, affecting fatty acid mobilization, maybe through ATGL-dependent and independent mechanisms, but the impact on oncogenesis remains to be defined.

Several proteins act as co-suppressors of ATGL, including G0/G1 switch gene 2 (G0S2) [134], and hypoxia-inducible lipid droplet associated protein (HILPDA; also known as hypoxia-inducible gene 2, HIG2) [130, 132]. Knockdown of HILPDA increased lipid droplet-derived fatty acid mobilization, due to less suppression of ATGL activity, resulting in increased fatty acid oxidation and ROS production, and impaired the growth of HCT116 colon carcinoma and HeLa tumor xenograft growth [130]. Similar observations were reported in HILPDA loss-of-function studies in HCT116 colon carcinoma models [132]. Overall, reduced ATGL activity, via knockdown of ATGL or ABHD5, has similar effects on cancer cell proliferation and viability as increased ATGL activity, via overexpression of ATGL or knockdown of HILPDA. This suggests that too much or too little TG hydrolysis and fatty acid mobilization has deleterious effects on cancer cell viability, but it remains unclear by what mechanism(s) this occurs. It is possible that it may involve the prevention of lipotoxicity as well as PPAR-α signaling (see review [128]).

The next step in the lipolytic cascade is the hydrolysis of sn-1,3 DG into MG and the release of one fatty acid (Fig. 3). This reaction is catalyzed by HSL, which has broad substrate specificity including cholesteryl esters, TG, MG, and retinyl esters [138], and to date, little is known about the role of HSL in cancer. The final step in neutral lipolysis is catabolism of MG by monoglyceride lipase (encoded by MGLL) to produce a fatty acid and glycerol. Despite the MG pool being small relative to TG, DG, and glycerophospholipids levels, monoglyceride lipase exerts significant influence on cancer cell behavior [139, 140], and MG hydrolysis enzyme activity is more than 11-fold higher in cancerous lung tissues than in paired non-cancerous tissues [141]. Recently, ABHD6 has been reported to be the primary MG lipase in NSCLC and blockade of ABHD6 significantly reduced the migration, invasion, and in vivo tumor growth of NSCLC [141]. Importantly, the loss-of-function of monoglyceride lipase was partly rescued by exogenous palmitate supplementation, thereby demonstrating that the fatty acids liberated by MG hydrolysis are essential mediators of cancer cell migration [140].

Lipid droplets also store sterol ester, in particular cholesteryl ester, and 1-O-acylceramides, which have a fatty acyl attached that is liberated during hydrolysis and contributes to the intracellular fatty acid pool. The molecular mechanisms of cholesteryl ester hydrolysis are overall poorly understood, with many candidate enzymes being proposed to possess cholesteryl ester hydrolase activity. They include CES1, HSL, KIAA1363/NCEH1, and possibly CES3 [142‐144], but no consensus has been reached, let alone insight into their role in cancer. It will be challenging to determine the role that these cholesteryl ester hydrolase candidates play in fatty acid metabolism as many of these enzymes have an affinity for multiple lipid and non-lipid substrates (i.e., (pro)drugs and environmental toxicants) and that cholesteryl ester hydrolysis will influence cellular cholesterol levels as well as fatty acid levels [124, 143]. Finally, the molecular regulators of lipid droplet contained 1-O-acylceramide hydrolysis are unknown and so the role that 1-O-acylceramide breakdown in cell biology is yet to be described.

Lipid droplet-contained fatty acids can also be mobilized via lipophagy [145], which is the autophagic degradation of lipid droplets (Fig. 3). Some of the molecular mechanisms that facilitate lipophagy have been identified, and recent examples include RAB7 and ATG5 [146, 147], PNPLA5 [148], ATG14 and Ulk1 [149], MAP1S [150], and LAMP1 [151]. In general, lipophagy plays a tumor suppressor role as it leads to increased intracellular free fatty acid levels, which promotes cell death via ferroptosis, ROS production, and ER stress [149, 151]. However, it has also been proposed that autophagy can play a pro-tumor role in nutrient-deprived situations where mobilization of fatty acids via this pathway is used for subsequent catabolic and anabolic processing [145]. Currently, the contribution of lipid droplet-containing fatty acids to the intracellular fatty acid pool, relative to the other sources, let alone whether these are mobilized by lipolysis or lipophagy, is unknown.

Overall, cancer cells have many diverse sources of fatty acids that can supply the intracellular fatty acid pool. The stoichiometric relationships of the various supply lines remain to be defined. Still, they are likely to be heavily influenced by substrate availability, including low or high extracellular fatty acid and lipoprotein levels, as well as non-lipid precursors for de novo fatty acid synthesis. Further, the relationships between these pathways will also be influenced by extracellular cues, including hormonal stimulation and oxygen availability (see reviews [39, 40]).