Lipid Metabolic Alterations in KRAS Mutant Tumors: History
View Latest Version
Please note this is an old version of this entry, which may differ significantly from the current revision.
Contributor:
Georgia Konstantinidou

KRAS is one of the most commonly mutated genes, an event that leads to development of highly aggressive and resistant to any type of available therapy tumors. Mutated KRAS drives a complex network of lipid metabolic rearrangements to support the adaptation of cancer cells to harsh environmental conditions and ensure their survival. Because there has been only a little success in the continuous efforts of effectively targeting KRAS-driven tumors, it is of outmost importance to delineate the exact mechanisms of how they get rewired, leading to this distinctive phenotype.

  • KRAS-driven tumors
  • lipid metabolism
  • tumor vulnerabilities

1. Fatty Acid Uptake, Biosynthesis, and Degradation

The highly conserved process of FA synthesis incorporates carbons from acetyl-CoA into growing FAs, thus providing the cell with the required substrate for cell growth signaling, membrane building, and energy storage. In the 50′s, a seminal observation was made directly linking lipid biogenesis and cancer. Medes et al. described that tumor cells exhibit higher rates of lipogenesis and heavily rely on these de novo produced FAs in order to meet their increased metabolic needs [1].
Intriguingly, KRAS-driven cancers do not seem to follow this principle. Instead, these tumors scavenge serum FAs, with a preference for long-chain polyunsaturated FAs and lysophospholipids [2][3], while de novo lipid synthesis, although still present, seems to be dispensable. Pancreatic ductal adenocarcinomas (PDAC) and colorectal cancer (CRC) cells uptake FAs secreted from adipocytes, as a response to nutrient starvation, or from FAs secreted from cancer associated fibroblasts and inhibition of this process by blocking CD36 -a FA transporter on the cell surface reduces their metastatic capacity [4][5][6]. Interestingly, decreased REDD1 expression, a stress response gene which is associated with peroxisome proliferator activated receptor γ (PPARγ)/CD36 activation, predicts poor outcomes selectively in KRAS mutant but not KRAS wild-type human lung and pancreatic adenocarcinomas [7]. Notably, in the context of mutant KRAS, REDD1 loss is associated with suppressed FA synthesis, conferring resistance to acetyl-CoA carboxylase (ACC) inhibitors [7]. Therefore, CD36 is an attractive target for the treatment of KRAS mutant tumors.
The available data regarding whether de novo FA synthesis can be considered as a distinctive vulnerability for KRAS mutated tumors are, to say the least, divisive. The presence of multiple rate-limiting enzymes in the biosynthetic process creates many potential targets. The first step in lipid biosynthesis is catalyzed by ATP-citrate lyase (ACLY), a metabolic enzyme that cleaves cytoplasmic citrate, thus generating acetyl-CoA. Recently, Carrer et al. elegantly demonstrated in a Pdx1-Cre/KrasG12D mouse model that pancreas-specific ablation of ACLY suppressed acinar-to-ductal metaplasia, suggesting a role in early pancreatic tumorigenesis [8].
ACC is another gatekeeper in the FA synthesis process, responsible for malonyl-CoA production. In PDAC, ACC inhibitors blocked cancer cell proliferation and suppressed tumor growth by simultaneously downregulating the WNT and Hedgehog signaling pathways [9]. Moreover, inhibition of the ACC enzymes ACC1 and ACC2 in a preclinical model of KrasG12D-driven non-small cell lung cancer (NSCLC), led to reduced tumor growth [10]. However, as stated above, KRAS mutant tumors heavily rely on extracellularly derived lipids, therefore it is likely that ACC1/2 inhibition may not have robust antitumor effects. This is also confirmed by the authors finding that ACC1 knockout cell lines could survive only when supplied with exogenous palmitate [10].
Regarding fatty acid synthase (FASN), increased expression is often reported in many different cancer types and has in fact been linked to gemcitabine resistance in KRAS mutated PDAC [11]. The implicated signaling pathways vary in a tissue-specific manner. Ventura and colleagues proposed that pharmacological inhibition of FASN with TVB-3166 reduces cell proliferation and xenograft tumor growth of KRAS mutated NSCLC, but not CRC cell lines via, at least in part, inhibition of the PI3K–AKT–mTOR pathway [12]. This claim is in part supported by two other recent reports where it is shown that KRAS mutated NSCLCs exhibit increased FASN levels and its inhibition results in decreased tumor formation [13][14]. In PDAC, FASN inhibition led to decreased cancer cell proliferation and increased apoptosis [15]. To date, attempts of pharmacologically targeting FASN have been challenged by severe toxicity and compensatory mechanisms activated by cancer cells, namely upregulation of FA uptake [16]. One compound, TVB-2640, is currently in a phase 2 clinical trial in NSCLC patients bearing KRAS mutations (NCT03808558).
Once generated, FAs undergo sequential desaturation steps, a process performed mainly by stearoyl-CoA desaturase (SCD). However, accumulating evidence suggested that KRAS mutant cancer cells bypass sensitivity to SCD inhibition by scavenging unsaturated fatty acids from lysophospholipids [3]. Moreover, recent evidence showed that KRAS mutant cancer cells are also able to desaturate palmitate to sapienate via fatty acid desaturase 2 (FADS2), an event that confers additional independence to SCD-mediated FA desaturation [17].
For FA synthesis to take place unobstructed, vital secondary metabolites are required, which become available through glycolysis. Therefore, attenuating the glycolytic machinery could prove to be beneficial by restricting not only the energy supply, but also the availability of indispensable, for lipid generation, metabolites. In this direction, the importance of glucose transporters in Kras-driven lung adenocarcinomas has been investigated, revealing that in a KrasLSL-G12D/WT;Trp53flox/flox lung adenocarcinoma mouse model concomitant deletion of the glucose transporters Glut1 and Glut3, significantly decreases tumor progression [18]. On a similar note, pharmacologic inhibition of an alternative glucose transport system, the sodium-dependent glucose transporter Sglt2, delayed the development of lung adenocarcinomas [19]. Taking into account the fact that compounds for Sglt2 inhibition are already being used in the clinic to treat diabetes, it is realistic to say that targeting the lipid metabolism of cancer cells through the glycolytic pathway may become possible in the near future for the subset of patients with tumors that are heavily dependent on glycolysis.
Several key enzymes of the FA synthesis process undergo transcriptional regulation by sterol regulatory element-binding protein 1 (SREBP1), including FASN [20][21]. Mutant KRAS regulates SREBP1 mainly by upregulating the PI3K/Akt/mTOR signaling pathway [22] As expected, SREBP1 has been found upregulated in KRAS mutant PDAC patients and its expression correlates with poorer prognosis [23]. NSCLC cells also exhibit increased SREBP1 levels; however, a novel, non-canonical role has recently been attributed to the gene, as it was shown that its ablation inhibits tumor growth, not by affecting lipogenesis, but by altering mitochondrial function and impairing oxidative phosphorylation [24]. Similarly, in CRC cells, genetic inhibition of either SREBP1 or SREBP2 leads to reduced tumor initiation and growth in vitro and in vivo, accompanied by decreased oxidative phosphorylation and glycolysis levels [25]. Hence, it would undoubtedly be of interest to investigate whether SREBP, apart from its well-established role in FA and cholesterol synthesis, is also directly regulating genes involved in mitochondrial function in a setting of KRAS-driven tumorigenesis.
On the other side of the spectrum, FA degradation by β-oxidation (FAO) is responsible for energy supply and NADPH production. For years, the synthesis and degradation of FAs were considered as two mutually exclusive cellular processes. FAO is tightly regulated by carnitine palmitoyltransferase 1 (CPT1), an enzyme that couples FAs with carnitine to enable their translocation to the mitochondria. Increased FAO levels directly correlate with accelerated progression and therapy resistance in KRAS-driven tumors. This phenotype can be established via multiple mechanisms. In KRAS mutant NSCLC, FAO is facilitated by the upregulation of long-chain acyl-CoA synthetase 3 (ACSL3), the enzyme responsible for activating free FAs, allowing tumor cells to meet the increased ATP demand [2]. Moreover, loss of REDD1 in KRAS mutant NSCLC not only increases FAO, but also allows cancer cells to cope with increased reactive oxygen species (ROS) by boosting ROS detoxifying systems [7]. Interestingly, both ACSL3 upregulation and REDD1 loss exhibit enhanced uptake of lysophospholipids and lipid storage. Notably, in a KrasG12D-driven PDAC mouse model, inhibition of FAO reduces the tumor-initiating potential of a dormant, oncogene ablation-resistant, population of tumor cells that is often linked to relapse [26]. In colorectal cancer, elevated FAO is associated with extracellular FA-mediated AMPK activation, an event that strongly favors FAs as the main mitochondrial substrate for energy production [4].

2. Autophagy

The role of autophagy in cancer initiation and progression, as well as therapeutic targeting has been extensively described in many excellent reviews [27][28], but the general consensus is that its complex function acts in favor of the tumors. In KRAS-driven malignancies, the autophagic machinery is commonly upregulated [29]. However, tumor initiation and progression are affected differently, with the former being inhibited and the latter promoted by autophagy. Indeed, in a KrasG12D-driven PDAC mouse model, genetic ablation of autophagy results in increased tumor initiation, however, these tumors are lacking the ability to progress into adenocarcinomas, thus leading to improved survival irrespective of the p53 tumor suppressor status [30]. In KrasG12D-driven NSCLC, loss of autophagy via Atg7 deletion before lung tumor onset did not affect tumor growth, while blocked tumor progression in mice with preexisting lung cancer [31].
In PDAC cells, an interesting crosstalk has been unveiled between pancreatic stellate cells (PSCs) and cancer cells. In a nutrient-deprived context, PDAC cells stimulate autophagy in PSCs, and this leads to secretion of alanine. Alanine is then uptaken by PDAC cells and used as alternative fuel for the TCA cycle, thus sustaining mitochondrial oxygen consumption and lipid synthesis and promoting growth during tumor initiation, but not progression [32]. The complexity of the interplay between tumor cells and the TME has also been elegantly shown in a recent work revealing that, upon induction of autophagy as a response to oxidative stress, KRASG12D protein is secreted from PDAC cells and uptaken by macrophages. This event further stimulates FAO, promotes pro-tumor macrophage M2 polarization and facilitates tumor progression [33]. The above results suggested that inhibition of autophagy could be highly effective in suppressing PDAC growth. However, this was not the case as clinical trials observed limited to no efficacy of hydroxychloroquine, an orally administered FDA-approved autophagy inhibitor [34].
When it comes to the control of lipid metabolism, autophagy seems to be critical for maintaining fat and glycogen stores in mice both during fasting and fed state. Indeed, systemic Atg7 ablation leads to depletion of white adipose tissue in mice [31]. Moreover, inhibition of autophagy in CRC cells thwarts their ability to use FAs and hinders their proliferation, suggesting that mutant KRAS controls lipid fate by modulating the autophagic machinery [4].

3. The ATX-LPA Signaling Axis

Lysophosphatidic acid (LPA) is a phospholipid consisting of a glycerol backbone, a phosphate group and a FA chain that can vary in length and saturation [35]. LPA acts through LPA receptors, which are G-protein-coupled receptors, thus initiating signaling cascades involved in key cellular processes such as proliferation, survival, cytoskeletal changes, and calcium signaling [36][37]. LPA is produced mainly through two different pathways. Firstly, LPA can be generated via conversion of lysophosphatidylcholine (LPC) by autotaxin (ATX), a secreted glycoprotein that has lysophospholipase D (lysoPLD) activity [38]. Alternatively, phosphatidic acid (PA) can get deacylated by phospholipase A1 or A2 (PLA1, PLA2) [39].
Numerous in vitro and in vivo studies suggest that alterations in the levels of LPA or its receptors are involved in tumorigenesis. By far the largest body of research, with regard to the LPA signaling axis, focuses on LPA1 and LPA3 receptors. It has been shown that in PDAC cells treated long-term with cisplatin, the activity of LPA1 and LPA receptors increased cell motility and invasion capacity, as well as MMP-2 activation [40]. These advantages could be ablated by LPA1 and LPA3 receptor knockdown, indicating that the two receptors strongly contribute to the malignant phenotype that cells acquire during tumor progression in PDAC. The same work also suggested that increased cell motility of the long-term cisplatin treated cells is, at least in part, regulated by extracellular LPA produced by ATX [40]. On a similar note, PANC-R9 cells (a highly invasive cell line established from PANC-1 pancreatic cancer cell line) were found to have higher expression levels of LPA1 receptor compared to the parental cell line, while the expression level of LPA3 receptor was decreased. Interestingly, the invasive capacity was amplified by LPA only in the PANC-R9 cells, an event that was accompanied with increased expression level of ATX [41]. Recently, it has been revealed that PDAC cells uptake FAs and particularly LPC derived from PSCs, which they then use either for membrane synthesis, or as precursors to produce signaling lipids, mainly LPA, through ATX-mediated LPC hydrolysis. Of note, ATX is overexpressed in human PDAC and its inhibition, either genetically or pharmacologically, suppressed tumor growth in an orthotopic PDAC mouse model [42]. LPA has also been involved in PDAC cell chemotaxis and metastasis in vivo [43]. Key player of this regulation is the Neural Wiskott-Aldrich Syndrome Protein (N-WASP), which drives LPA1 receptor recycling and prevents its degradation. Overall, these findings imply that the two LPA receptors, as well as LPA per se, may have a direct impact in tumor aggressiveness and resistance to drugs and their targeting could simultaneously thwart multiple hallmarks of cancer, while sparing normal cells.
The connection between the expression of LPA receptors and drug resistance has also been investigated in a CRC setting. DLD1 cells that underwent long term treatment with fluorouracil (5-FU) exhibit a robust increase in the expression levels of LPA1 receptor. Conversely, cisplatin treatment led to a markedly elevated expression of LPA6 receptor [44]. Using the same CRC cell line model, Shida et al. showed that LPA through LPA1 receptor stimulates cell migration, proliferation, adhesion, and the secretion of both VEGF and -in a dose dependent manner- IL-8, boosting their metastatic potential. The mechanism of LPA-induced IL-8 secretion, included degradation of IκBα and consequent activation of NF-κB [45].
Similarly, in a KrasG12D–driven lung adenocarcinoma mouse model, genetic deletion of ATX or LPA1 receptor suppressed lung tumorigenesis, confirming a pro-tumorigenic role of LPA signaling also in this context [46]. Notably, ATX was recently found to modulate also the TME because it can have a chemorepulsive action on tumor-infiltrating lymphocytes and circulating CD8+ T cells [47].
Taken altogether, it is becoming evident that KRAS-driven tumors heavily rely on the ATX-LPA signaling axis to acquire and maintain their aggressive and often therapy resistant phenotype. The mechanisms through which this is achieved seem to be cell line dependent and dictates further research. The heterogeneity in the expression pattern of LPA receptors across the different cancer types acts once more as a reminder that a “one size fits all” approach cannot be applied when planning a treatment strategy. It remains to be investigated why certain tumors rely on different LPA receptors, or their inhibition, to promote functions that will allow their propagation.

4. The Role of Polyunsaturated Fatty Acids (PUFAs) and Cholesterol

In view of the dependency of KRAS-driven tumors on exogenous lipids, when designing approaches to target lipid signaling it is important to consider the impact of the different extracellularly derived lipids on KRAS mutant cancer cells. During the last decades, a strong link between diet and cancer has been established, with high fat diet promoting tumor initiation, progression and, eventually, poorer survival [48]. Intriguingly, high calorie diet-induced obesity which leads to hypertrophic tumor-associated adipocytes in mouse and human PDAC setting, seems to increase the incidence and accelerate tumorigenesis by increasing inflammation [49][50][51]. However, in KrasG12D-driven lung cancer, a high calorie diet dampens tumor progression if given before tumor onset, by causing defective unfolded protein response (UPR) and unresolved endoplasmic reticulum (ER) stress, but increases tumor burden when administered after tumor onset [52]. Whether the timing of high calorie diet is important for tumor development and the exact underpinning mechanisms remain to be investigated, however, ER chaperones are unveiled as an attractive way to selectively target KRAS-driven lung tumors.
Although the role of specific FAs in different human cancer types is not fully unraveled yet, it is becoming clear that a high uptake of ω-3 PUFAs, such as docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA), can reduce the risk of initiation and progression of some cancers including colorectal, prostate, and breast cancer [53][54][55][56][57]. In fact, Collet et al. initially proposed a direct reduction of RAS localization to the plasma membrane by DHA [58]. However, it was later shown that incorporation of DHA and EPA into the plasma membrane phospholipids suppresses phosphatidic acid-dependent oncogenic KRAS-driven effector interactions (i.e., ERK signaling), thus suppressing hyperproliferation of cancer cells in vitro and in vivo [59]. Therefore, DHA rises as a promising compound that could be used to treat RAS-driven cancers.
Accumulating data obtained from both in vitro and in vivo studies point towards a direct cytotoxic effect of certain PUFAs on cancer cells [60][61][62]. This is because certain phosphoglycerolipids (mainly phosphatidylethanolamines) containing polyunsaturated acyl chains are subjected to either oxidation by ROS or selective oxygenation by lipoxygenases (LOX), a process named lipid peroxidation, which leads to ferroptosis (an iron-dependent, non-apoptotic cell death induced by the accumulation of lipid hydroperoxides) [63][64][65]. Therefore, KRAS mutant tumors are equipped with potent antioxidant systems to be able to overcome the cytotoxic effects of PUFAs and generally survive in the presence of high ROS levels [66]. In this context, the glutathione peroxidase 4 (GPX4) is an important ferroptosis regulator as it converts toxic lipid hydroperoxides to non-toxic lipid alcohols at the expense of glutathione (GSH) [67]. This dependency exposes a cancer cell vulnerability, in which inhibition of GPX4 either directly or indirectly with erastin (a compound which inhibits the import of cystine, leading to GSH depletion and inactivation of GPX4) results in excessive lipid peroxidation causing cancer cell death [68]. Indeed, with the view of the ongoing effort to develop drugs that can efficiently target KRAS mutated cancer cells, there are many recent reports that successfully leveraged the ferroptosis machinery to promote cell death in KRAS-driven tumors [69][70][71].

This entry is adapted from the peer-reviewed paper 10.3390/ijms24021793

References

  1. Medes, G.; Thomas, A.; Weinhouse, S. Metabolism of neoplastic tissue. IV. A study of lipid synthesis in neoplastic tissue slices in vitro. Cancer Res. 1953, 13, 27–29.
  2. Padanad, M.S.; Konstantinidou, G.; Venkateswaran, N.; Melegari, M.; Rindhe, S.; Mitsche, M.; Yang, C.; Batten, K.; Huffman, K.E.; Liu, J.; et al. Fatty Acid Oxidation Mediated by Acyl-CoA Synthetase Long Chain 3 Is Required for Mutant KRAS Lung Tumorigenesis. Cell Rep. 2016, 16, 1614–1628.
  3. Kamphorst, J.J.; Cross, J.R.; Fan, J.; de Stanchina, E.; Mathew, R.; White, E.P.; Thompson, C.B.; Rabinowitz, J.D. Hypoxic and Ras-transformed cells support growth by scavenging unsaturated fatty acids from lysophospholipids. Proc. Natl. Acad. Sci. USA 2013, 110, 8882–8887.
  4. Wen, Y.-A.; Xing, X.; Harris, J.W.; Zaytseva, Y.Y.; Mitov, M.I.; Napier, D.L.; Weiss, H.L.; Mark Evers, B.; Gao, T. Adipocytes activate mitochondrial fatty acid oxidation and autophagy to promote tumor growth in colon cancer. Cell Death Dis. 2017, 8, e2593.
  5. Gong, J.; Lin, Y.; Zhang, H.; Liu, C.; Cheng, Z.; Yang, X.; Zhang, J.; Xiao, Y.; Sang, N.; Qian, X.; et al. Reprogramming of lipid metabolism in cancer-associated fibroblasts potentiates migration of colorectal cancer cells. Cell Death Dis. 2020, 11, 267.
  6. Okumura, T.; Ohuchida, K.; Sada, M.; Abe, T.; Endo, S.; Koikawa, K.; Iwamoto, C.; Miura, D.; Mizuuchi, Y.; Moriyama, T.; et al. Extra-pancreatic invasion induces lipolytic and fibrotic changes in the adipose microenvironment, with released fatty acids enhancing the invasiveness of pancreatic cancer cells. Oncotarget 2017, 8, 18280–18295.
  7. Qiao, S.; Koh, S.-B.; Vivekanandan, V.; Salunke, D.; Patra, K.C.; Zaganjor, E.; Ross, K.; Mizukami, Y.; Jeanfavre, S.; Chen, A.; et al. REDD1 loss reprograms lipid metabolism to drive progression of RAS mutant tumors. Genes Dev. 2020, 34, 751–766.
  8. Carrer, A.; Trefely, S.; Zhao, S.; Campbell, S.L.; Norgard, R.J.; Schultz, K.C.; Sidoli, S.; Parris, J.L.D.; Affronti, H.C.; Sivanand, S.; et al. Acetyl-CoA Metabolism Supports Multistep Pancreatic Tumorigenesis. Cancer Discov. 2019, 9, 416–435.
  9. Petrova, E.; Scholz, A.; Paul, J.; Sturz, A.; Haike, K.; Siegel, F.; Mumberg, D.; Liu, N. Acetyl-CoA carboxylase inhibitors attenuate WNT and Hedgehog signaling and suppress pancreatic tumor growth. Oncotarget 2017, 8, 48660–48670.
  10. Svensson, R.U.; Parker, S.J.; Eichner, L.J.; Kolar, M.J.; Wallace, M.; Brun, S.N.; Lombardo, P.S.; Van Nostrand, J.L.; Hutchins, A.; Vera, L.; et al. Inhibition of acetyl-CoA carboxylase suppresses fatty acid synthesis and tumor growth of non-small-cell lung cancer in preclinical models. Nat. Med. 2016, 22, 1108–1119.
  11. Tadros, S.; Shukla, S.K.; King, R.J.; Gunda, V.; Vernucci, E.; Abrego, J.; Chaika, N.V.; Yu, F.; Lazenby, A.J.; Berim, L.; et al. De Novo Lipid Synthesis Facilitates Gemcitabine Resistance through Endoplasmic Reticulum Stress in Pancreatic Cancer. Cancer Res. 2017, 77, 5503–5517.
  12. Ventura, R.; Mordec, K.; Waszczuk, J.; Wang, Z.; Lai, J.; Fridlib, M.; Buckley, D.; Kemble, G.; Heuer, T.S. Inhibition of de novo Palmitate Synthesis by Fatty Acid Synthase Induces Apoptosis in Tumor Cells by Remodeling Cell Membranes, Inhibiting Signaling Pathways, and Reprogramming Gene Expression. EBioMedicine 2015, 2, 808–824.
  13. Gouw, A.M.; Eberlin, L.S.; Margulis, K.; Sullivan, D.K.; Toal, G.G.; Tong, L.; Zare, R.N.; Felsher, D.W. Oncogene KRAS activates fatty acid synthase, resulting in specific ERK and lipid signatures associated with lung adenocarcinoma. Proc. Natl. Acad. Sci. USA 2017, 114, 4300–4305.
  14. Singh, A.; Ruiz, C.; Bhalla, K.; Haley, J.A.; Li, Q.K.; Acquaah-Mensah, G.; Montal, E.; Sudini, K.R.; Skoulidis, F.; Wistuba, I.I.; et al. De novo lipogenesis represents a therapeutic target in mutant Kras non-small cell lung cancer. FASEB J. 2018, 32, 7018–7027.
  15. Bian, Y.; Yu, Y.; Wang, S.; Li, L. Up-regulation of fatty acid synthase induced by EGFR/ERK activation promotes tumor growth in pancreatic cancer. Biochem. Biophys. Res. Commun. 2015, 463, 612–617.
  16. Röhrig, F.; Schulze, A. The multifaceted roles of fatty acid synthesis in cancer. Nat. Rev. Cancer 2016, 16, 732.
  17. Vriens, K.; Christen, S.; Parik, S.; Broekaert, D.; Yoshinaga, K.; Talebi, A.; Dehairs, J.; Escalona-Noguero, C.; Schmieder, R.; Cornfield, T.; et al. Evidence for an alternative fatty acid desaturation pathway increasing cancer plasticity. Nature 2019, 566, 403–406.
  18. Contat, C.; Ancey, P.-B.; Zangger, N.; Sabatino, S.; Pascual, J.; Escrig, S.; Jensen, L.; Goepfert, C.; Lanz, B.; Lepore, M.; et al. Combined deletion of Glut1 and Glut3 impairs lung adenocarcinoma growth. eLife 2020, 9, e53618.
  19. Scafoglio, C.R.; Villegas, B.; Abdelhady, G.; Bailey, S.T.; Liu, J.; Shirali, A.S.; Wallace, W.D.; Magyar, C.E.; Grogan, T.R.; Elashoff, D.; et al. Sodium-glucose transporter 2 is a diagnostic and therapeutic target for early-stage lung adenocarcinoma. Sci. Transl. Med. 2018, 10, eaat5933.
  20. Shimano, H.; Yahagi, N.; Amemiya-Kudo, M.; Hasty, A.H.; Osuga, J.; Tamura, Y.; Shionoiri, F.; Iizuka, Y.; Ohashi, K.; Harada, K.; et al. Sterol regulatory element-binding protein-1 as a key transcription factor for nutritional induction of lipogenic enzyme genes. J. Biol. Chem. 1999, 274, 35832–35839.
  21. Towle, H.C. Metabolic regulation of gene transcription in mammals. J. Biol. Chem. 1995, 270, 23235–23238.
  22. Ricoult, S.J.; Yecies, J.L.; Ben-Sahra, I.; Manning, B.D. Oncogenic PI3K and K-Ras stimulate de novo lipid synthesis through mTORC1 and SREBP. Oncogene 2016, 35, 1250–1260.
  23. Sun, Y.; He, W.; Luo, M.; Zhou, Y.; Chang, G.; Ren, W.; Wu, K.; Li, X.; Shen, J.; Zhao, X.; et al. SREBP1 regulates tumorigenesis and prognosis of pancreatic cancer through targeting lipid metabolism. Tumour Biol. 2015, 36, 4133–4141.
  24. Ruiz, C.F.; Montal, E.D.; Haley, J.A.; Bott, A.J.; Haley, J.D. SREBP1 regulates mitochondrial metabolism in oncogenic KRAS expressing NSCLC. FASEB J. 2020, 34, 10574–10589.
  25. Wen, Y.-A.; Xiong, X.; Zaytseva, Y.Y.; Napier, D.L.; Vallee, E.; Li, A.T.; Wang, C.; Weiss, H.L.; Evers, B.M.; Gao, T. Downregulation of SREBP inhibits tumor growth and initiation by altering cellular metabolism in colon cancer. Cell Death Dis. 2018, 9, 265.
  26. Viale, A.; Pettazzoni, P.; Lyssiotis, C.A.; Ying, H.; Sanchez, N.; Marchesini, M.; Carugo, A.; Green, T.; Seth, S.; Giuliani, V.; et al. Oncogene ablation-resistant pancreatic cancer cells depend on mitochondrial function. Nature 2014, 514, 628–632.
  27. Mathew, R.; Karantza-Wadsworth, V.; White, E. Role of autophagy in cancer. Nat. Rev. Cancer 2007, 7, 961–967.
  28. Levy, J.M.M.; Towers, C.G.; Thorburn, A. Targeting autophagy in cancer. Nat. Rev. Cancer 2017, 17, 528–542.
  29. Guo, J.Y.; Chen, H.Y.; Mathew, R.; Fan, J.; Strohecker, A.M.; Karsli-Uzunbas, G.; Kamphorst, J.J.; Chen, G.; Lemons, J.M.; Karantza, V.; et al. Activated Ras requires autophagy to maintain oxidative metabolism and tumorigenesis. Genes Dev. 2011, 25, 460–470.
  30. Yang, A.; Rajeshkumar, N.V.; Wang, X.; Yabuuchi, S.; Alexander, B.M.; Chu, G.C.; Von Hoff, D.D.; Maitra, A.; Kimmelman, A.C. Autophagy Is Critical for Pancreatic Tumor Growth and Progression in Tumors with p53 Alterations. Cancer Discov. 2014, 4, 905.
  31. Karsli-Uzunbas, G.; Guo, J.Y.; Price, S.; Teng, X.; Laddha, S.V.; Khor, S.; Kalaany, N.Y.; Jacks, T.; Chan, C.S.; Rabinowitz, J.D.; et al. Autophagy is required for glucose homeostasis and lung tumor maintenance. Cancer Discov. 2014, 4, 914–927.
  32. Sousa, C.M.; Biancur, D.E.; Wang, X.; Halbrook, C.J.; Sherman, M.H.; Zhang, L.; Kremer, D.; Hwang, R.F.; Witkiewicz, A.K.; Ying, H.; et al. Pancreatic stellate cells support tumour metabolism through autophagic alanine secretion. Nature 2016, 536, 479–483.
  33. Dai, E.; Han, L.; Liu, J.; Xie, Y.; Kroemer, G.; Klionsky, D.J.; Zeh, H.J.; Kang, R.; Wang, J.; Tang, D. Autophagy-dependent ferroptosis drives tumor-associated macrophage polarization via release and uptake of oncogenic KRAS protein. Autophagy 2020, 16, 2069–2083.
  34. Wolpin, B.M.; Rubinson, D.A.; Wang, X.; Chan, J.A.; Cleary, J.M.; Enzinger, P.C.; Fuchs, C.S.; McCleary, N.J.; Meyerhardt, J.A.; Ng, K.; et al. Phase II and pharmacodynamic study of autophagy inhibition using hydroxychloroquine in patients with metastatic pancreatic adenocarcinoma. Oncologist 2014, 19, 637–638.
  35. Tokumura, A. A family of phospholipid autacoids: Occurrence, metabolism and bioactions. Prog. Lipid Res. 1995, 34, 151–184.
  36. Fukushima, N.; Ishii, I.; Contos, J.J.; Weiner, J.A.; Chun, J. Lysophospholipid receptors. Annu. Rev. Pharmacol. Toxicol. 2001, 41, 507–534.
  37. Ishii, I.; Fukushima, N.; Ye, X.; Chun, J. Lysophospholipid receptors: Signaling and biology. Annu Rev. Biochem 2004, 73, 321–354.
  38. Umezu-Goto, M.; Kishi, Y.; Taira, A.; Hama, K.; Dohmae, N.; Takio, K.; Yamori, T.; Mills, G.B.; Inoue, K.; Aoki, J.; et al. Autotaxin has lysophospholipase D activity leading to tumor cell growth and motility by lysophosphatidic acid production. J. Cell Biol. 2002, 158, 227–233.
  39. Fourcade, O.; Simon, M.F.; Viode, C.; Rugani, N.; Leballe, F.; Ragab, A.; Fournie, B.; Sarda, L.; Chap, H. Secretory phospholipase A2 generates the novel lipid mediator lysophosphatidic acid in membrane microvesicles shed from activated cells. Cell 1995, 80, 919–927.
  40. Fukushima, K.; Takahashi, K.; Yamasaki, E.; Onishi, Y.; Fukushima, N.; Honoki, K.; Tsujiuchi, T. Lysophosphatidic acid signaling via LPA1 and LPA3 regulates cellular functions during tumor progression in pancreatic cancer cells. Exp. Cell Res. 2017, 352, 139–145.
  41. Fukushima, K.; Otagaki, S.; Takahashi, K.; Minami, K.; Ishimoto, K.; Fukushima, N.; Honoki, K.; Tsujiuchi, T. Promotion of cell-invasive activity through the induction of LPA receptor-1 in pancreatic cancer cells. J. Recept. Signal. Transduct. 2018, 38, 367–371.
  42. Auciello, F.R.; Bulusu, V.; Oon, C.; Tait-Mulder, J.; Berry, M.; Bhattacharyya, S.; Tumanov, S.; Allen-Petersen, B.L.; Link, J.; Kendsersky, N.D.; et al. A Stromal Lysolipid-Autotaxin Signaling Axis Promotes Pancreatic Tumor Progression. Cancer Discov. 2019, 9, 617–627.
  43. Juin, A.; Spence, H.J.; Martin, K.J.; McGhee, E.; Neilson, M.; Cutiongco, M.F.A.; Gadegaard, N.; Mackay, G.; Fort, L.; Lilla, S.; et al. N-WASP Control of LPAR1 Trafficking Establishes Response to Self-Generated LPA Gradients to Promote Pancreatic Cancer Cell Metastasis. Dev. Cell 2019, 51, 431–445.e7.
  44. Takahashi, K.; Fukushima, K.; Otagaki, S.; Ishimoto, K.; Minami, K.; Fukushima, N.; Honoki, K.; Tsujiuchi, T. Effects of LPA1 and LPA6 on the regulation of colony formation activity in colon cancer cells treated with anticancer drugs. J. Recept. Signal Transduct. 2018, 38, 71–75.
  45. Shida, D.; Kitayama, J.; Yamaguchi, H.; Okaji, Y.; Tsuno, N.H.; Watanabe, T.; Takuwa, Y.; Nagawa, H. Lysophosphatidic Acid (LPA) Enhances the Metastatic Potential of Human Colon Carcinoma DLD1 Cells through LPA1. Cancer Res. 2003, 63, 1706.
  46. Magkrioti, C.; Oikonomou, N.; Kaffe, E.; Mouratis, M.A.; Xylourgidis, N.; Barbayianni, I.; Megadoukas, P.; Harokopos, V.; Valavanis, C.; Chun, J.; et al. The Autotaxin-Lysophosphatidic Acid Axis Promotes Lung Carcinogenesis. Cancer Res. 2018, 78, 3634–3644.
  47. Matas-Rico, E.; Frijlink, E.; van der Haar Avila, I.; Menegakis, A.; van Zon, M.; Morris, A.J.; Koster, J.; Salgado-Polo, F.; de Kivit, S.; Lanca, T.; et al. Autotaxin impedes anti-tumor immunity by suppressing chemotaxis and tumor infiltration of CD8(+) T cells. Cell Rep. 2021, 37, 110013.
  48. Willett, W.C. Diet and cancer: One view at the start of the millennium. Cancer Epidemiol. Biomark. Prev. 2001, 10, 3–8.
  49. Incio, J.; Liu, H.; Suboj, P.; Chin, S.M.; Chen, I.X.; Pinter, M.; Ng, M.R.; Nia, H.T.; Grahovac, J.; Kao, S.; et al. Obesity-Induced Inflammation and Desmoplasia Promote Pancreatic Cancer Progression and Resistance to Chemotherapy. Cancer Discov. 2016, 6, 852–869.
  50. Berrington de Gonzalez, A.; Sweetland, S.; Spencer, E. A meta-analysis of obesity and the risk of pancreatic cancer. Br. J. Cancer 2003, 89, 519–523.
  51. Calle, E.E.; Rodriguez, C.; Walker-Thurmond, K.; Thun, M.J. Overweight, obesity, and mortality from cancer in a prospectively studied cohort of U.S. adults. N. Engl. J. Med. 2003, 348, 1625–1638.
  52. Ramadori, G.; Konstantinidou, G.; Venkateswaran, N.; Biscotti, T.; Morlock, L.; Galie, M.; Williams, N.S.; Luchetti, M.; Santinelli, A.; Scaglioni, P.P.; et al. Diet-Induced Unresolved ER Stress Hinders KRAS-Driven Lung Tumorigenesis. Cell Metab. 2015, 21, 117–125.
  53. Lipworth, L. Epidemiology of breast cancer. Eur. J. Cancer Prev. 1995, 4, 7–30.
  54. Potter, J.D. Risk factors for colon neoplasia—Epidemiology and biology. Eur. J. Cancer 1995, 31, 1033–1038.
  55. Key, T. Risk factors for prostate cancer. Cancer Surv. 1995, 23, 63–77.
  56. Moro, K.; Nagahashi, M.; Ramanathan, R.; Takabe, K.; Wakai, T. Resolvins and omega three polyunsaturated fatty acids: Clinical implications in inflammatory diseases and cancer. World J. Clin. Cases 2016, 4, 155–164.
  57. Cockbain, A.J.; Toogood, G.J.; Hull, M.A. Omega-3 polyunsaturated fatty acids for the treatment and prevention of colorectal cancer. Gut 2012, 61, 135.
  58. Collett, E.D.; Davidson, L.A.; Fan, Y.-Y.; Lupton, J.R.; Chapkin, R.S. n-6 and n-3 polyunsaturated fatty acids differentially modulate oncogenic Ras activation in colonocytes. Am. J. Physiol. Cell Physiol. 2001, 280, C1066–C1075.
  59. Fuentes, N.R.; Mlih, M.; Barhoumi, R.; Fan, Y.Y.; Hardin, P.; Steele, T.J.; Behmer, S.; Prior, I.A.; Karpac, J.; Chapkin, R.S. Long-Chain n-3 Fatty Acids Attenuate Oncogenic KRas-Driven Proliferation by Altering Plasma Membrane Nanoscale Proteolipid Composition. Cancer Res. 2018, 78, 3899–3912.
  60. Klurfeld, D.M.; Bull, A.W. Fatty acids and colon cancer in experimental models. Am. J. Clin. Nutr. 1997, 66, 1530S–1538S.
  61. Chapkin, R.S.; Seo, J.; McMurray, D.N.; Lupton, J.R. Mechanisms by which docosahexaenoic acid and related fatty acids reduce colon cancer risk and inflammatory disorders of the intestine. Chem. Phys. Lipids 2008, 153, 14–23.
  62. Trombetta, A.; Maggiora, M.; Martinasso, G.; Cotogni, P.; Canuto, R.A.; Muzio, G. Arachidonic and docosahexaenoic acids reduce the growth of A549 human lung-tumor cells increasing lipid peroxidation and PPARs. Chem. Biol. Interact. 2007, 165, 239–250.
  63. Stockwell, B.R.; Friedmann Angeli, J.P.; Bayir, H.; Bush, A.I.; Conrad, M.; Dixon, S.J.; Fulda, S.; Gascon, S.; Hatzios, S.K.; Kagan, V.E.; et al. Ferroptosis: A Regulated Cell Death Nexus Linking Metabolism, Redox Biology, and Disease. Cell 2017, 171, 273–285.
  64. Yang, W.S.; Kim, K.J.; Gaschler, M.M.; Patel, M.; Shchepinov, M.S.; Stockwell, B.R. Peroxidation of polyunsaturated fatty acids by lipoxygenases drives ferroptosis. Proc. Natl. Acad. Sci. USA 2016, 113, E4966–E4975.
  65. Dixon, S.J.; Lemberg, K.M.; Lamprecht, M.R.; Skouta, R.; Zaitsev, E.M.; Gleason, C.E.; Patel, D.N.; Bauer, A.J.; Cantley, A.M.; Yang, W.S.; et al. Ferroptosis: An iron-dependent form of nonapoptotic cell death. Cell 2012, 149, 1060–1072.
  66. DeNicola, G.M.; Karreth, F.A.; Humpton, T.J.; Gopinathan, A.; Wei, C.; Frese, K.; Mangal, D.; Yu, K.H.; Yeo, C.J.; Calhoun, E.S.; et al. Oncogene-induced Nrf2 transcription promotes ROS detoxification and tumorigenesis. Nature 2011, 475, 106–109.
  67. Ursini, F.; Maiorino, M.; Valente, M.; Ferri, L.; Gregolin, C. Purification from pig liver of a protein which protects liposomes and biomembranes from peroxidative degradation and exhibits glutathione peroxidase activity on phosphatidylcholine hydroperoxides. Biochim. Biophys. Acta 1982, 710, 197–211.
  68. Yang, W.S.; Stockwell, B.R. Ferroptosis: Death by Lipid Peroxidation. Trends Cell Biol. 2016, 26, 165–176.
  69. Yang, J.; Mo, J.; Dai, J.; Ye, C.; Cen, W.; Zheng, X.; Jiang, L.; Ye, L. Cetuximab promotes RSL3-induced ferroptosis by suppressing the Nrf2/HO-1 signalling pathway in KRAS mutant colorectal cancer. Cell Death Dis. 2021, 12, 1079.
  70. Bartolacci, C.; Andreani, C.; Vale, G.; Berto, S.; Melegari, M.; Crouch, A.C.; Baluya, D.L.; Kemble, G.; Hodges, K.; Starrett, J.; et al. Targeting de novo lipogenesis and the Lands cycle induces ferroptosis in KRAS-mutant lung cancer. Nat. Commun. 2022, 13, 4327.
  71. Hu, K.; Li, K.; Lv, J.; Feng, J.; Chen, J.; Wu, H.; Cheng, F.; Jiang, W.; Wang, J.; Pei, H.; et al. Suppression of the SLC7A11/glutathione axis causes synthetic lethality in KRAS-mutant lung adenocarcinoma. J. Clin. Investig. 2020, 130, 1752–1766.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
This entry is offline, you can click here to edit this entry!
Video Production Service
Feedback