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Chemoresistance constitute nowadays the major contributor to therapy failure in most cancers. There are main factors that mitigate cell response to therapy, such as target organ, inherent sensitivity to the administered compound, its metabolism, drug efflux and influx or alterations on specific cellular targets, among others. We now know that intrinsic properties of cancer cells, including metabolic features, substantially contribute to chemoresistance. In fact, during the last years, numerous reports indicate that cancer cells resistant to chemotherapy demonstrate significant alterations in mitochondrial metabolism, membrane polarization and mass. Metabolic activity and expression of several mitochondrial proteins are modulated under treatment to cope with stress, making these organelles central players in the development of resistance to therapies. Here, we review the role of mitochondria in chemoresistant cells in terms of metabolic rewiring and function of key mitochondria-related proteins.
Mitochondria, chemoresistance, oxidative phosphorylation, fatty acid oxidation, metabolism, uncoupling proteins, ABC transporters
Initial studies on cancer metabolism in early 20th century overlooked mitochondria in tumors, as they were deemed dysfunctional. Further reports established their critical role in cancer metabolism and, thus, reflected on their key function in disease survival and progression. Although traditionally known for their function in cellular bioenergetics coordinating ATP production from cellular respiration, mitochondria play a role in multitude of central cellular activities in physiological and pathophysiological conditions. Indeed, through their multiple regulatory activities, they boost the tumorigenic and metastatic potential in cancer cells[3-5].
On the one hand, they produce precursors for the synthesis of macromolecules such as lipids, proteins, DNA and RNA, which are key to sustain the high proliferative rates of cancer cells. Additionally, mitochondria are central for redox balance: although they are the main producers of reactive oxygen species (ROS) as respiration by-products, mitochondria modulate redox equivalents and gather the majority of enzymatic and non-enzymatic antioxidants. The control of redox balance and excessive ROS accumulation is essential for cancer cells, known to present high ROS levels to increase pro-proliferative signaling and activity, and further promoting tumorigenesis through accumulation of new mutations. Apart from ROS, mitochondria also dispose of other cellular waste products, such as ammonia through the urea cycle.
Mitochondria provide a common site for multiple metabolic reactions in order to meet energy and biomolecule demands, thus controlling and integrating the diverse metabolic pathways taking place in these organelles. As such, mitochondria enable cancer cells to modulate their metabolism under various stress conditions providing metabolic flexibility[8-11].
Together with cell death regulation, the above-mentioned functions make mitochondria crucial stress sensors, allowing flexibility to adapt to adverse environments such as nutrient deprivation, oxidative stress and DNA damage; all of them present during cancer progression and especially during treatment.
As mentioned above, mitochondria integrate and balance the different metabolic pathways within these organelles, with direct influence on other routes taking place in alternative sites such as the cytoplasm, peroxisomes, etc. This metabolic plasticity is a key factor for adaptation to stressful situations, and development of resistance to diverse anticancer drugs.
Targeting metabolic pathways of cancer has emerged as a promising therapeutic approach. Indeed, myriad metabolic inhibitors have shown encouraging results as anticancer therapeutic agents. However, the emergence of resistant populations, due to metabolic rewiring or intrinsic metabolic heterogeneity found in tumors, highlight some weaknesses in this approach.
Biguanides are a class of well-known metabolic drugs thanks to metformin, which is widely prescribed as anti-diabetic drug. The interest on the antitumoral effects of metformin emerged upon the publication of a retrospective report suggesting that type 2 diabetes patients taking oral metformin showed reduced cancer incidence. Metformin has shown promising results against various cancers, such as breast, endometrial and prostate cancer[13,14]. Nowadays, it is well accepted that systemic effects on glucose blood levels and direct inhibition of the mitochondrial electron transport chain (ETC) contribute to the antitumoral effects of this drug. At the cellular level, metformin treatment induces energetic stress subsequent to reduced ATP production upon ETC blockade, inhibiting proliferation and inducing cell death. Importantly, metformin seems to be especially toxic for cancer stem cells (CSCs), the main tumorigenic and chemoresistant subpopulation found in tumors, due to their dependence on mitochondrial activity[16,17]. However, further studies showed that prolonged exposure to metformin gave rise to resistance in vivo and in vitro in several cancer models, due to the rearrangement of metabolic routes. Although resistance to metformin in pancreatic cancer cells was mainly linked to the ability of cells to increase compensatory glycolysis, other studies suggest that mitochondria are able to adapt to the dysfunctional ETC by upregulating alternative mitochondrial pathways. In fact, initial studies on hepatocytes found that metformin treatment induced lipid metabolism, guiding subsequent studies in cancer cells. For example, studies conducted on lung cancer cells suggested that metformin treatment altered cellular metabolic phenotype, via AMP-activated protein kinase (AMPK) activation, to reduce dependence on oxidative phosphorylation (OXPHOS) and keep the TCA (tricarboxylic acid) cycle activity for lipid synthesis. Under hypoxic conditions, ETC inhibition led to an increased influx of glutamine (Gln) into the TCA cycle, highlighting another mechanism used by cancer cells to adapt to suppressed ETC activity. In fact, the susceptibility to metformin of cancer cells, particularly the CSC subset, may be dependent on Gln metabolism: only the combination of metformin with a glutaminase C inhibitor had a substantial CSC suppressing effect.
Phenformin, another biguanide targeting complex I of mitochondria, was found to be more potent at inhibiting cell growth in non-small cell lung cancer. However, the emergence of a phenformin-resistant population was observed. It has been suggested that the development of resistance is dependent on functional LKB1-AMPK signaling, which promotes a shift in their metabolism to overcome inhibitory effects of phenformin. This drug has been withdrawn from the market due to its extreme side effects, although research on phenformin has been reactivated lately due to the mild effects observed upon metformin treatment.
Cancer cells have been found to be dependent on Gln, with a significant upregulation in its uptake[24,25]. For that reason, the effects of Gln deprivation or inhibition of the enzymes related to its metabolism has been widely studied in vitro. Glutaminase (GA) is a mitochondrial enzyme that converts Gln to glutamate, which is further converted to α-ketoglutarate to fuel the TCA cycle for energy or biomolecule synthesis. GA expression is upregulated in cancers to boost Gln metabolism, and thus has been proposed as an attractive therapeutic target[26-30]. GA inhibitors were found to slow tumor growth, increase cell death and sensitize chemotherapy-resistant cells[31-33]. Along with drugs, interference RNA is a strategy used to inhibit activity of glutaminase[34,31]. However, Gln-dependent cancer cells were able to adapt to GA knockdown by increasing pyruvate carboxylase activity, allowing glucose-derived pyruvate to be shunted into the TCA cycle through oxalacetate.
Gln-addicted cells may also be dependent on Gln synthetase (GS) to cover their Gln needs: this enzyme catalyzes the synthesis of Gln from glutamate and ammonia. De novo synthesis of Gln plays an important role in cancer survival and adaptation to altered cellular needs and, consequently, many cancer cells upregulate the expression of this enzyme[36-38]. Diverse studies underlined the importance of GS in the development of resistance to Gln deprivation. In fact, GS knockdown inhibited the growth of sarcoma cell lines in Gln-deprived media, but had no effect in media supplemented with the aminoacid. Importantly, GS overexpression may confer resistance to Gln deprivation in a non-autonomous fashion: luminal breast cancer cells overexpressed GS that, not only allowed them to grow without Gln, but also supported the growth of basal cancer cells in co-culture. On the other hand, these studies suggest that cancer cells may develop resistance to anti-GA therapy by increasing de novo Gln synthesis via GS to meet their needs, and inhibition of GA and GS together would be necessary to overcome resistance.
Lipid metabolism is frequently altered in cancer cells to meet energy demands, synthesize cellular membranes and mediate cell signaling[40-42]. Indeed, cancer cells largely rely on de novo lipid synthesis by overexpression of fatty acid synthase (FASN) making FASN inhibition an attractive therapeutic approach with remarkable anticancer activity[44-46]. However, studies in colorectal cancer suggest development of resilience to FASN inhibitors. Although some evidence points towards altered signaling of metabolism-related kinases (AKT and AMPK) for development of resistance to FASN inhibitors, metabolic plasticity conferred by mitochondrial lipid metabolism constitutes the strongest candidate[40,42]. Furthermore, cancer cells are also efficient in taking up fatty acids from the microenvironment, another reason FASN inhibition alone may only target a small subset of cells and combination therapies would be necessary.
BRAF is a common oncogene in many cancers, conferring constitutive activation of proliferation pathways. For that reason, BRAF inhibitors have been extensively studied, showing promising results against various cancer types, especially melanoma. However, the emergence of resistance to BRAF inhibitors is a major challenge associated with these drugs. Among the various proposed mechanisms for drug resistance, mitochondrial metabolism seems to play a key role. On the one hand, resistant melanoma cells demonstrated an increment of mitochondrial respiration conferred by increased expression of ETC genes and densification of mitochondrial cristae. In addition, they showed a high dependency on mitochondrial metabolic activity, maintained on withdrawal of the drug, making them sensitive to mitochondria-targeting compounds in combined therapy. On the other hand, BRAF inhibition with vemurafenib led to an increase in Gln uptake consequence of Gln synthesis inhibition. For that reason, resistant cells were found to be dependent on mitochondrial Gln, constituting an important vulnerability of these cells.
The tyrosine kinase inhibitor imatinib is used mainly to treat chronic myeloid leukemia. This anticancer agent has also been linked to the development of resistant populations. Contrary to BRAF inhibition-resistant cells, the development of resistance to imatinib in various cancers has been linked to increase in glycolysis rather than mitochondrial metabolism. Nevertheless, chemoresistance originates in the mitochondria: mitochondrial malfunction in imatinib-resistant cells promote the accumulation of TCA cycle intermediates and NADH, as well as increased ROS generation, as compared to sensitive cells. Interestingly, ROS accumulation was found to be the primary cause of resistance to this drug.
The antimetabolite 5-fluoro-uracil (5FU) has been extensively used as anticancer treatment for a long time. A major issue with this drug, however, is the development of resistance leading to limited clinical use. Among the studies tackling the role of mitochondria on resistance to 5FU, most of them carried out in colon cancer cells, we can identify two opposing mechanisms: resistance through either Warburg-related features or OXPHOS-dependency. On the one hand, increased glucose uptake and metabolism, coupled to mitochondrial dysfunction, has been traditionally considered a hallmark of the Warburg effect. Related to this, it was found that the glucose transporter GLUT1 inhibitor resensitized resistant cancer cells to 5FU. Additionally, other studies linked the development of resistance to downregulation of mitochondrial ATP synthase activity, which altered cellular bioenergetics and increased glucose dependency. On the other hand, resistance to 5FU has been related to increased mitochondrial mass and activity, amplified expression of ETC enzymes and higher rates of oxygen consumption[57,58]. Interestingly, due to their OXPHOS-dependency, resistant cells were found to be sensitive to Complex I inhibition by metformin. Strikingly, OXPHOS involvement in resistance to 5FU was linked to the development of stemness-related features, directly connecting CSCs to mitochondrial metabolism as suggested previously[59,60]. In line with this, tumor-associated mesenchymal stem cells conferred chemoresistance to 5FU by promoting stemness in gastric cancer cells via mitochondrial fatty acid oxidation (FAO) activation. In fact, treatment with the FAO inhibitor etomoxir was able to partially mitigate resistance to 5FU.
The development of resistance to the alkylating agent cisplastin has been associated with a metabolic shift from glycolysis to OXPHOS in adenocarcinoma and hepatoma cells. The rewiring of cellular metabolism in those tumors was controlled by SIRT1, a master regulator of cellular functions with important implications on metabolism. Interestingly, cisplastin resistance in ovarian cancer cells has been also linked to altered metabolism and increased dependency on OXPHOS-generated energy[63,64]. In fact, treatment with the Bcl-2 proteins inhibitor ABT-737 was able to reverse cisplastin resistance, possibly by targeting mitochondrial metabolism. Additionally, the FAO inhibitor perhexiline also sensitized cisplastin-resistant ovarian cancer cells.
Resistance to the antimicrotubule agent paclitaxel has also been associated to mitochondrial metabolism, although in this case independently of glucose. In fact, chemoresistant cells showed a distinct metabolic profile from their sensitive counterparts and their proliferation was inhibited on treatment with FAO inhibitors like mercaptoacetate and etomoxir.
The cell cycle inhibitor Cytarabine is a commonly administered chemotherapeutic drug for leukemias and lymphomas, with reduced clinical implications due to development of resistance[67,68]. Among the possible mechanisms, Cytarabine treatment favored the expansion of a chemoresistant leukemic stem cell subset with high FAO/OXPHOS activity. Accordingly, the FAO inhibitor etomoxir repressed oxygen consumption in acute myeloid leukemia cells and sensitized cells to cytarabine.
Besides metabolism, mitochondria contribute to drug resistance through specific mitochondrial proteins allowing for survival and proliferation under treatment. In fact, the key role of mitochondria in drug resistance has been traditionally attributed to some of the proteins discussed below.
Uncoupling proteins (UCPs) have been found to be overexpressed in different cancers and affect mitochondrial potential and ROS levels[70-72]. UCPs are inner mitochondrial membrane proteins that provide an alternative path for proton entry into mitochondrial matrix, thereby leading to a reduced electric gradient across the membranes. As the name suggests, they uncouple ATP formation from the ETC and promote the free flow of protons across the membrane, thus leading to a decrease in membrane potential and ROS formation.
UCPs have been found to have major implications in cancer and are associated with metabolic reprogramming and therapy resistance[74,75]. As mentioned above, these proteins reduce the coupling between two necessary pathways, namely: energy and biomolecule generation pathways. In fact, it has been proposed they modulate cellular metabolism to deal with stress. In cancer cells, UCP2 expression was found to promote glycolysis and fatty acid metabolism, thus playing a major role in metabolic plasticity. On the other hand, UCP2-expressing cells demonstrated better tolerance towards oxidative stress, which is key in chemoresistance. For instance, UCP2 expression was found to reduce superoxide production by gemcitabine to produce a protective effect on pancreatic cancer cells, which was reverted by UCP2 inhibition with genipin. Similar results were found in human leukemia, where drug-resistant cells overexpressed UCP2 and genipin sensitized these cells to epirubicin and doxorubicin by increasing intracellular ROS. Further, due to decrease in ROS formation, UCPs also promote the antiapoptotic activity of cells by inhibition of cytochrome c release and increasing antiapoptotic protein expression[83-85].
Several drug transporters are expressed on mitochondrial membrane and their expression is modulated by mitochondrial activity, further contributing to developing resistance by excessive drug efflux[86,87]. ATP-binding cassette (ABC) is a super family of transporters known to be responsible for drug efflux and, therefore, drug resistance. In fact, drug-resistant cancer cells, particularly CSCs, are found to overexpress ABCs[89,90]. Under oxidative stress, cells elevate the expression of various ABC transporters in order to facilitate the entry of glutathione to mitigate it. For that reason, it has been suggested that ABC expression is higher in cancer cells which are mitochondria-dependent (including CSCs): they balance their oxidative state controlling ETC-produced ROS, while mitochondrial activity would provide the substrate (ATP) required for ABC activity[91,92].
It is well known that many DNA-targeting chemotherapeutic agents used in cancer induce drug resistance through upregulation of multidrug resistance (MRD) associated genes and multidrug resistance associated proteins (MRP), members of the ABC family of transporters. This was initially described for etoposide-resistance in ovarian cancer cells, in which MRPs allowed efflux of the drug, thereby disallowing its activity. A similar mechanism was found in breast cancer cells where MRP expression promoted efflux of the drug and reduced its cellular activity. Interestingly, some families of ABCs, including MRPs, can be expressed on mitochondrial membranes, mediating drug resistance by either altering drug transport in order to protect mtDNA (mitochondrial DNA) or inhibiting mitochondrial outer membrane permeability, thus blocking apoptosis. In fact, doxorubicin resistance in melanoma cells was associated with expression of ABCB8, a member of ABC transporter family expressed on the inner mitochondrial membrane: ABCB8 overexpression provided doxorubicin resistance protecting mtDNA from damage, while its knockdown resensitized cells to doxorubicin. On the other hand, resistance to the topoisomerase inhibitor mitoxantrone could be induced by overexpression of ABCG2 on the mitochondrial membrane, increasing the efflux of the drug. The study of mitochondrial drug transporters on therapy resistance is still elusive and needs to be explored further.
The role of mitochondria and mitochondrial proteins in apoptosis has been extensively studied and reviewed[96,97]. Anti-apoptotic proteins are overexpressed in cancer, making them an important contributor in the development of resistance[98-101].
Cancer cells are enriched in antiapoptotic proteins that inhibit the activity of proapoptotic counterparts. In fact, the prosurvival Bcl-2 proteins bind proapoptotic proteins to maintain the mitochondrial membrane potential and inhibit the release of cytochrome c from mitochondria, thus evading apoptosis[102,103]. Bcl-2 is upregulated in chemoresistant cells from small cell lung cancer, prostate cancer, ovarian and breast cancer[63,103-106]. Similarly, Bcl-xL and Bcl-w, also members of Bcl-2 pro-survival family, are overexpressed in some chemoresistant cancer cells[63,107]. Importantly, the upregulation of Bcl-2 expression enabled the development of cross-resistance to chemotherapeutic drugs.
Interestingly, alternative mechanisms have been described to favor an antiapoptotic balance in chemoresistant cancer cells. In fact, cells with depleted mtDNA accumulated unprocessed caspases (necessary for apoptosis), which has been suggested as one of the resistance mechanisms. Further, loss of signaling metabolites in cancer cells upregulated pro-survival proteins, promoting chemoresistance.
Pro-survival Bcl-2 proteins are negatively regulated by BH3-only (BCL-2 homology domain only) proteins. This has led to the development of small inhibitory molecules called “BH3 mimetics”[110,111]. These small molecules successfully sensitized chemoresistant cancer cells to treatment. For example, the Bcl-2 antagonist ABT-263 induced mitochondria-dependent apoptosis, enhancing activity of chemotherapeutic agents to inhibit small cell lung carcinoma and lymphoid malignancies in vitro and in vivo. Similarly, the Bcl-2 inhibitor ABT-737 antagonized Bcl-2 prosurvival proteins to induce the regression of solid tumors. ABT-737 was able to induce significant cytotoxicity in ovarian cancer cells by inhibiting mitochondria respiration and thus sensitized therapy-resistant ovarian cancer cells to chemotherapy. Interestingly, the FAO inhibitors etomoxir and ranolazine activated Bak, a proapoptotic member of the Bcl-2 family, enhancing ABT-737 pro-apoptotic activity in leukemia. Obatoclax, another BH3 mimetic, was found to be effective in targeting chemoresistant cells, as well as increasing efficiency of chemotherapeutic drugs.
Along with altered mitochondrial bioenergetics, cancer cells may increase their mitochondrial mass to rescue them from energy scarcity, and promote survival, growth and propagation[116-120]. The main regulators of mitochondrial biogenesis are the PGC-1 [PPAR (peroxisome proliferator-activated receptor)-γ coactivator] family of transcription factors, expressed in energetic cells[121,122]. PGC-1 family members, particularly PGC-1α, have been found to be overexpressed in several cancer types, as well as CSC subsets[18,123-126], contributing to tumorigenesis and metastasis.
Importantly, PGC-1α upregulation and enhanced mitochondrial biogenesis are important contributors of chemoresistance development[127-131]. In fact, cancer cells alter PGC-1α expression in response to therapy-induced stress, leading to the development of resistance in various ways.
On the one hand, DNA-targeting anticancer agents promote mitochondrial biogenesis[132,133], a process controlled by AMPK-dependent PGC-1α upregulation in HeLa cells. Indeed, cells resistant to these agents showed high mitochondrial mass that provides protection against DNA damage[63,135].
PGC-1α induction may also confer chemoresistance by promoting a metabolic shift to overcome ATP requirements, as described for cells treated with BRAF inhibitors (Haq et al. 2013). This is also the case for 5FU resistance, which enhanced PGC-1α expression and consequently altered cellular metabolism to mitigate the energetic stress induced by treatment[58,102,131].
Finally, it is important to highlight the regulatory role of PGC-1α and mitochondrial mass for CSCs[18,135,137], the main subpopulation responsible for chemoresistance. For instance, PGC-1α silencing sensitized chemoresistant stem-like cells to chemotherapy.
These reports suggest that PGC-1α targeting could help to overcome therapeutic resistance. Although no specific inhibitors of PGC-1α are available to date, indirect inhibition through upstream modulators may be possible. In fact, MAPK inhibitor-resistant melanoma cells were sensitized to therapy when treated with mTORC inhibitors, which decreased PGC-1α expression and reversed metabolic changes.
Contrary to the traditional view, mitochondria are now considered the “Achilles heel” of cancer due to the heavy dependency of cancer cells (especially CSCs) on both mitochondrial metabolism and ability to cope with stress. In fact, these mitochondrial properties allow cancer cells to develop resistance to various therapies [Figure 1], suggesting that multimodal therapies targeting mitochondria would constitute promising therapeutic strategies[33,138,139].
Figure 1. Overview of the mitochondrial determinants of resistance. Some therapeutic drugs promote a shift in metabolism. Drugs like biguanides that target the complex I of mitochondria promote a shift to glycolysis to meet ATP needs. Similarly, glutaminase inhibitors promote cells to switch to oxidative phosphorylation and BRAF protein inhibitors push cells to metabolize glutamine. Mitochondrial FAO contributes to chemoresistance to 5-fluoro-uracil (5FU), Cisplatin and Cytarabine. Increased intracellular ROS induced by enhanced expression of uncoupling proteins (UCPs), electron transport chain (ETC) activity or drug action are also responsible for chemoresistance. In addition to metabolic shifts, therapeutic drugs also elevate the expression of mitochondrial ABC transporters proteins that allow drug efflux and helps to minimize the inhibitory activity of the drug. Bcl-2 family of prosurvival proteins block apoptosis induced by chemotherapeutic drugs. Mitochondrial biogenesis is also induced during therapy-induced stress by elevation of PGC1α
However, much research is still needed to unveil the role of mitochondria in cancer. In fact, tumor metabolism may not only change with disease progression and course of treatment: due to intrinsic heterogeneity, mitochondrial function may be variable across tumors and cell subpopulations within a tumor. In fact, it is well known that the activity of mitochondria is largely dependent on various factors, including physical location, nutrient and oxygen availability and the microenvironment. Indeed, cancer-associated cells also exhibit different metabolic phenotypes and directly affect cancer cells metabolism, adding an extra layer of variability. Therefore, carefully designed therapies targeting diverse populations and considering the tumor microenvironment will be crucial in order to develop successful treatment strategies.
We would like to thank Beatriz Parejo-Alonso for critical reading of the manuscript, and Laura Sancho Andrés for language editing.Authors’ contributions
Wrote the main draft and designed the figure: Bokil A
Developed the study concept, obtained funding and wrote the final version of the manuscript: Sancho P
Contributed to the conception and design of the manuscript, wrote sections, revised, read and approved the submitted version: Both authorsAvailability of data and materials
Not applicable.Financial support and sponsorship
This work was supported by the Miguel Servet Program (CP16/00121) and a FIS (Fondo Investigaciones Sanitarias) grant (PI17/00082), both from the Instituto de Salud Carlos III and cofinanced by European funds (FSE: “el FSE invierte en tu futuro” and FEDER: “una manera de hacer Europa,” respectively).Conflicts of interest
All authors declared that there are no conflicts of interest.Ethical approval and consent to participate
Not applicable.Consent for publication
© The Author(s) 2019.
1. Warburg O, Wind F, Negelein E. The metabolism of tumors in the body. J Gen Physiol 1927;8:519-30.DOIPubMedPMC
2. Pedersen PL. Tumor mitochondria and the bioenergetics of cancer cells. Prog Exp Tumor Res 1978;22:190-274.DOIPubMed
3. Pavlova NN, Thompson CB. The emerging hallmarks of cancer metabolism. Cell Metab 2016;23:27-47.DOIPubMedPMC
4. Weinberg SE, Chandel NS. Targeting mitochondria metabolism for cancer therapy. Nat Chem Biol 2015;11:9.DOIPubMedPMC
5. Wallace DC. Mitochondria and cancer. Nat Rev Cancer 2012;12:685-98.DOIPubMedPMC
6. Vyas S, Zaganjor E, Haigis MC. Mitochondria and cancer. Cell 2016;166:555-66.DOIPubMedPMC
7. Spinelli JB, Haigis MC. The multifaceted contributions of mitochondria to cellular metabolism. Nat Cell Biol 2018;20:745-54.DOIPubMedPMC
8. Cluntun AA, Lukey MJ, Cerione RA, Locasale JW. Glutamine metabolism in cancer: understanding the Heterogeneity. Trends Cancer 2017;3:169-80.DOIPubMedPMC
9. Lukey MJ, Katt WP, Cerione RA. Targeting amino acid metabolism for cancer therapy. Drug Discov Today 2017;22:796-804.DOIPubMedPMC
10. DeBerardinis RJ, Chandel NS. Fundamentals of cancer metabolism. Sci Adv 2016;2:e1600200.DOIPubMedPMC
11. Gentric G, Mieulet V, Mechta-Grigoriou F. Heterogeneity in cancer metabolism: new concepts in an old field. Antioxid Redox Signal 2017;26:462-85.DOIPubMedPMC
12. Evans JMM, Donnelly LA, Emslie-Smith AM, Alessi DR, Morris AD. Metformin and reduced risk of cancer in diabetic patients. BMJ 2005;330:1304-5.DOIPubMedPMC
13. Bayraktar S, Hernadez-Aya LF, Lei X, Meric-Bernstam F, Litton JK, et al. Effect of metformin on survival outcomes in diabetic patients with triple receptor-negative breast cancer. Cancer 2012;118:1202-11.DOIPubMedPMC
14. Bodmer M, Becker C, Meier C, Jick SS, Meier CR. Use of metformin and the risk of ovarian cancer: a case-control analysis. Gynecol Oncol 2011;123:200-4.DOIPubMed
15. Wheaton WW, Weinberg SE, Hamanaka RB, Soberanes S, Sullivan LB, et al. Metformin inhibits mitochondrial complex I of cancer cells to reduce tumorigenesis. Elife 2014;3:e02242.DOIPubMedPMC
16. Jagust P, de Luxán-Delgado B, Parejo-Alonso B, Sancho P. Metabolism-based therapeutic strategies targeting cancer stem cells. Front Pharmacol 2019;10:203.DOIPubMedPMC
17. Heeschen C, Sancho P. More challenges ahead-metabolic heterogeneity of pancreatic cancer stem cells. Mol Cell Oncol 2016;3:e1105353.DOIPubMedPMC
18. Sancho P, Burgos-Ramos E, Tavera A, Bou Kheir T, Jagust P, et al. MYC/PGC-1alpha balance determines the metabolic phenotype and plasticity of pancreatic cancer stem cells. Cell Metab 2015;22:590-605.DOIPubMed
19. Zhou G, Myers R, Li Y, Chen Y, Shen X, et al. Role of AMP-activated protein kinase in mechanism of metformin action. J Clin Invest 2001;108:1167-74.DOIPubMedPMC
20. Griss T, Vincent EE, Egnatchik R, Chen J, Ma EH, et al. Metformin antagonizes cancer cell proliferation by suppressing mitochondrial-dependent biosynthesis. PLoS Biol 2015;13:e1002309.DOIPubMedPMC
21. Wang Y, Bai C, Ruan Y, Liu M, Chu Q, et al. Coordinative metabolism of glutamine carbon and nitrogen in proliferating cancer cells under hypoxia. Nat Commun 2019;10:201.DOIPubMedPMC
22. Kim JH, Lee KJ, Seo Y, Kwon JH, Yoon JP, et al. Effects of metformin on colorectal cancer stem cells depend on alterations in glutamine metabolism. Sci Rep 2018;8:409.DOIPubMedPMC
23. Shackelford DB, Abt E, Gerken L, Vasquez DS, Seki A, et al. LKB1 inactivation dictates therapeutic response of non-small cell lung cancer to the metabolism drug phenformin. Cancer Cell 2013;23:143-58.DOIPubMedPMC
24. Márquez J, Alonso FJ, Matés JM, Segura JA, Martín-Rufián M, et al. Glutamine Addiction In Gliomas. Neurochem Res 2017;42:1735-46.DOIPubMed
25. Mohamed A, Deng X, Khuri FR, Owonikoko TK. Altered glutamine metabolism and therapeutic opportunities for lung cancer. Clin Lung Cancer 2014;15:7-15.DOIPubMedPMC
26. Gao P, Tchernyshyov I, Chang TC, Lee YS, Kita K, et al. c-Myc suppression of miR-23a/b enhances mitochondrial glutaminase expression and glutamine metabolism. Nature 2009;458:762.DOIPubMedPMC
27. Gross MI, Demo SD, Dennison JB, Chen L, Chernov-Rogan T, et al. Antitumor activity of the glutaminase inhibitor CB-839 in triple-negative breast cancer. Mol Cancer Ther 2014;13:890-901.DOIPubMed
28. Qie S, Chu C, Li W, Wang C, Sang N. ErbB2 activation upregulates glutaminase 1 expression which promotes breast cancer cell proliferation. J Cell Biochem 2014;115:498-509.DOIPubMedPMC
29. Erickson JW, Cerione RA. Glutaminase: a hot spot for regulation of cancer cell metabolism? Oncotarget 2010;1:734-40.DOIPubMedPMC
30. Son J, Lyssiotis CA, Ying H, Wang X, Hua S, et al. Glutamine supports pancreatic cancer growth through a KRAS-regulated metabolic pathway. Nature 2013;496:101.DOIPubMedPMC
31. Seltzer MJ, Bennett BD, Joshi AD, Gao P, Thomas AG, et al. Inhibition of glutaminase preferentially slows growth of glioma cells with mutant IDH1. Cancer Res 2010;70:8981-7.DOIPubMedPMC
32. Tanaka K, Sasayama T, Irino Y, Takata K, Nagashima H, et al. Compensatory glutamine metabolism promotes glioblastoma resistance to mTOR inhibitor treatment. J Clin Invest 2015;125:1591-602.DOIPubMedPMC
33. Masamha CP, LaFontaine P. Molecular targeting of glutaminase sensitizes ovarian cancer cells to chemotherapy. J Cell Biochem 2018;119:6136-45.DOIPubMed
34. Wang J-B, Erickson JW, Fuji R, Ramachandran S, Gao P, et al. Targeting mitochondrial glutaminase activity inhibits oncogenic transformation. Cancer Cell 2010;18:207-19.DOIPubMedPMC
35. Cheng T, Sudderth J, Yang C, Mullen AR, Jin ES, et al. Pyruvate carboxylase is required for glutamine-independent growth of tumor cells. Proc Natl Acad Sci U S A 2011;108:8674-9.DOIPubMedPMC
36. Christa L, Simon MT, Flinois JP, Gebhardt R, Brechot C, et al. Overexpression of glutamine synthetase in human primary liver cancer. Gastroenterology 1994;106:1312-20.DOIPubMed
37. Issaq SH, Mendoza A, Fox SD, Helman LJ. Glutamine synthetase is necessary for sarcoma adaptation to glutamine deprivation and tumor growth. Oncogenesis 2019;8:20.DOIPubMedPMC
38. Yang L, Achreja A, Yeung TL, Mangala LS, Jiang D, et al. Targeting stromal glutamine synthetase in tumors disrupts tumor microenvironment-regulated cancer cell growth. Cell Metab 2016;24:685-700.DOIPubMed
39. Kung HN, Marks JR, Chi JT. Glutamine synthetase is a genetic determinant of cell type-specific glutamine independence in breast epithelia. PLoS Genet 2011;7:e1002229.DOIPubMedPMC
40. Kuo CY, Ann DK. When fats commit crimes: fatty acid metabolism, cancer stemness and therapeutic resistance. Cancer Commun (Lond) 2018;38:47.DOIPubMedPMC
41. Mancini R, Noto A, Pisanu ME, Vitis CD, Maugeri-Saccà M, et al. Metabolic features of cancer stem cells: the emerging role of lipid metabolism. Oncogene 2018;37:2367.DOIPubMed
42. Yi M, Li J, Chen S, Cai J, Ban Y, et al. Emerging role of lipid metabolism alterations in Cancer stem cells. J Exp Clin Cancer Res 2018;37:118.DOIPubMedPMC
43. Menendez JA, Lupu R. Fatty acid synthase and the lipogenic phenotype in cancer pathogenesis. Nat Rev Cancer 2007;7:763-77.DOIPubMed
44. Schcolnik-Cabrera A, Chavez-Blanco A, Dominguez-Gomez G, Taja-Chayeb L, Morales-Barcenas R, et al. Orlistat as a FASN inhibitor and multitargeted agent for cancer therapy. Expert Opin Investig Drugs 2018;27:475-89.DOIPubMed
45. Flavin R, Peluso S, Nguyen PL, Loda M. Fatty acid synthase as a potential therapeutic target in cancer. Future Oncol 2010;6:551-62.DOIPubMedPMC
46. Papaevangelou E, Almeida GS, Box C, deSouza NM, Chung YL. The effect of FASN inhibition on the growth and metabolism of a cisplatin-resistant ovarian carcinoma model. Int J Cancer 2018;143:992-1002.DOIPubMedPMC
47. Zaytseva YY, Rychahou PG, Le AT, Scott TL, Flight RM, et al. Preclinical evaluation of novel fatty acid synthase inhibitors in primary colorectal cancer cells and a patient-derived xenograft model of colorectal cancer. Oncotarget 2018;9:24787-800.DOIPubMedPMC
48. Nieman KM, Kenny HA, Penicka CV, Ladanyi A, Buell-Gutbrod R, et al. Adipocytes promote ovarian cancer metastasis and provide energy for rapid tumor growth. Nat Med 2011;17:1498-503.DOIPubMedPMC
49. Alcalá AM, Flaherty KT. BRAF inhibitors for the treatment of metastatic melanoma: clinical trials and mechanisms of resistance. Clin Cancer Res 2012;18:33-9.DOIPubMed
50. Baenke F, Chaneton B, Smith M, van den Broek N, Hogan K, et al. Resistance to BRAF inhibitors induces glutamine dependency in melanoma cells. Mol Oncol 2016;10:73-84.DOIPubMedPMC
51. Corazao-Rozas P, Guerreschi P, Jendoubi M, André F, Jonneaux A, et al. Mitochondrial oxidative stress is the Achille’s heel of melanoma cells resistant to Braf-mutant inhibitor. Oncotarget 2013;4:1986-98.DOIPubMedPMC
52. Hernandez-Davies JE, Tran TQ, Reid MA, Rosales KR, Lowman XH, et al. Vemurafenib resistance reprograms melanoma cells towards glutamine dependence. J Transl Med 2015;13:210.DOIPubMedPMC
53. Serkova N, Boros LG. Detection of resistance to Imatinib by metabolic profiling. Am J Pharmacogenomic 2005;5:293-302.DOIPubMed
54. Kluza J, Jendoubi M, Ballot C, Dammak A, Jonneaux A, et al. Exploiting mitochondrial dysfunction for effective elimination of Imatinib-resistant leukemic cells. PLoS One 2011;6:e21924.DOIPubMedPMC
55. Liu W, Fang Y, Wang XT, Liu J, Dan X, et al. Overcoming 5-Fu resistance of colon cells through inhibition of Glut1 by the specific inhibitor WZB117. Asian Pac J Cancer Prev 2014;15:7037-41.DOIPubMed
56. Shin YK, Yoo BC, Chang HJ, Jeon E, Hong SH, et al. Down-regulation of mitochondrial F1F0-ATP synthase in human colon cancer cells with induced 5-fluorouracil resistance. Cancer Res 2005;65:3162-70.DOIPubMed
57. Denise C, Paoli P, Calvani M, Taddei ML, Giannoni E, et al. 5-Fluorouracil resistant colon cancer cells are addicted to OXPHOS to survive and enhance stem-like traits. Oncotarget 2015;6:41706-21.DOIPubMedPMC
58. Vellinga TT, Borovski T, Boer VCJ de, Fatrai S, van Schelven S, et al. SIRT1/PGC1α-dependent increase in oxidative phosphorylation supports chemotherapy resistance of colon cancer. Clin Cancer Res 2015;21:2870-9.DOIPubMed
59. Sancho P, Barneda D, Heeschen C. Hallmarks of cancer stem cell metabolism. Br J Cancer 2016;114:1305-12.DOIPubMedPMC
60. Jagust P, de Luxán-Delgado B, Parejo-Alonso B, Sancho P. Metabolism-based therapeutic strategies targeting cancer stem cells. Front Pharmacol 2019;10:203.DOIPubMedPMC
61. He W, Liang B, Wang C, Li S, Zhao Y, et al. MSC-regulated lncRNA MACC1-AS1 promotes stemness and chemoresistance through fatty acid oxidation in gastric cancer. Oncogene 2019;38:4637.DOIPubMed
62. Liang XJ, Finkel T, Shen DW, Yin JJ, Aszalos A, et al. SIRT1 contributes in part to cisplatin resistance in cancer cells by altering mitochondrial metabolism. Mol Cancer Res 2008;6:1499-506.DOIPubMedPMC
63. Xu Y, Gao W, Zhang Y, Wu S, Liu Y, et al. ABT737 reverses cisplatin resistance by targeting glucose metabolism of human ovarian cancer cells. Int J Oncol 2018;53:1055-68.DOIPubMedPMC
64. Matassa DS, Amoroso MR, Lu H, Avolio R, Arzeni D, et al. Oxidative metabolism drives inflammation-induced platinum resistance in human ovarian cancer. Cell Death Differ 2016;23:1542.DOIPubMedPMC
65. Zhu J, Wu G, Song L, Cao L, Tan Z, et al. NKX2-8 deletion-induced reprogramming of fatty acid metabolism confers chemoresistance in epithelial ovarian cancer. EBioMedicine 2019;43:238-52.DOIPubMedPMC
66. Li J, Zhao S, Zhou X, Zhang T, Zhao L, et al. Inhibition of lipolysis by Mercaptoacetate and Etomoxir specifically sensitize drug-resistant lung adenocarcinoma cell to Paclitaxel. PLoS One 2013;8:e74623.DOIPubMedPMC
67. Freiburghaus C, Emruli VK, Johansson A, Olsson R, Ek F, et al. Resistance to cytarabine in mantle cell lymphoma is mediated by down-regulation of deoxycytidine kinase at the protein level. Blood 2016;128:1769.
68. Lokody I. Drug resistance: overcoming resistance in acute myeloid leukaemia treatment. Nat Rev Cancer 2014;14:452-3.DOIPubMed
69. Farge T, Saland E, Toni Fd, Aroua N, Hosseini M, et al. Chemotherapy-resistant human acute myeloid leukemia cells are not enriched for leukemic stem cells but require oxidative metabolism. Cancer Discov 2017;7:716-35.DOIPubMedPMC
70. Carretero MV, Torres L, Latasa U, García-Trevijano ER, Prieto J, et al. Transformed but not normal hepatocytes express UCP2. FEBS Lett 1998;439:55-8.DOIPubMed
71. Giatromanolaki A, Balaska K, Kalamida D, Kakouratos C, Sivridis E, et al. Thermogenic protein UCP1 and UCP3 expression in non-small cell lung cancer: relation with glycolysis and anaerobic metabolism. Cancer Biol Med 2017;14:396-404.DOIPubMedPMC
72. Horimoto M, Resnick MB, Konkin TA, Routhier J, Wands JR, et al. Expression of uncoupling protein-2 in human colon cancer. Clin Cancer Res 2004;10:6203-7.DOIPubMed
73. Klingenberg M. Uncoupling protein--a useful energy dissipator. J Bioenerg Biomembr 1999;31:419-30.DOIPubMed
74. Baffy G. Uncoupling protein-2 and cancer. Mitochondrion 2010;10:243-52.DOIPubMed
75. Samudio I, Fiegl M, Andreeff M. Mitochondrial uncoupling and the Warburg effect: molecular basis for the reprogramming of cancer cell metabolism. Cancer Res 2009;69:2163-6.DOIPubMedPMC
76. Baffy G. Mitochondrial uncoupling in cancer cells: liabilities and opportunities. Biochim Biophys Acta Bioenerg 2017;1858:655-64.DOIPubMed
77. Derdak Z, Mark NM, Beldi G, Robson SC, Wands JR, et al. The mitochondrial uncoupling protein-2 promotes chemoresistance in cancer cells. Cancer Res 2008;68:2813-9.DOIPubMedPMC
78. Nedergaard J, Cannon B. The ‘novel’ ‘uncoupling’ proteins UCP2 and UCP3: what do they really do? Pros and cons for suggested functions. Exp Physiol 2003;88:65-84.DOIPubMed
79. Pecqueur C, Alves-Guerra C, Ricquier D, Bouillaud F. UCP2, a metabolic sensor coupling glucose oxidation to mitochondrial metabolism? IUBMB Life 2009;61:762-7.DOIPubMed
80. Koziel A, Sobieraj I, Jarmuszkiewicz W. Increased activity of mitochondrial uncoupling protein 2 improves stress resistance in cultured endothelial cells exposed in vitro to high glucose levels. Am J Physiol Heart Circ Physiol 2015;309:H147-56.DOIPubMed
81. Dalla Pozza E, Fiorini C, Dando I, Menegazzi M, Sgarbossa A, et al. Role of mitochondrial uncoupling protein 2 in cancer cell resistance to gemcitabine. BBA-Mol Cell Res 2012;1823:1856-63.DOIPubMed
82. Mailloux RJ, Adjeitey CNK, Harper ME. Genipin-induced inhibition of uncoupling protein-2 sensitizes drug-resistant cancer cells to cytotoxic agents. PLoS One 2010;5:e13289.DOIPubMedPMC
83. Collins P, Jones C, Choudhury S, Damelin L, Hodgson H. Increased expression of uncoupling protein 2 in HepG2 cells attenuates oxidative damage and apoptosis. Liver Int 2005;25:880-7.DOIPubMed
84. Liu WY, He W, Li H. Exhaustive training increases uncoupling protein 2 expression and decreases Bcl-2/Bax ratio in rat skeletal muscle. Oxid Med Cell Longev 2013;2013:23365696.DOIPubMedPMC
85. Deng S, Yang Y, Han Y, Li X, Wang X, et al. UCP2 inhibits ROS-mediated apoptosis in A549 under hypoxic conditions. PLoS One 2012;7:e30714.DOIPubMedPMC
86. Cox J, Weinman S. Mechanisms of doxorubicin resistance in hepatocellular carcinoma. Hepat Oncol 2016;3:57-9.DOIPubMedPMC
87. Elliott AM, Al-Hajj MA. ABCB8 mediates doxorubicin resistance in melanoma cells by protecting the mitochondrial genome. Mol Cancer Res 2009;7:79-87.DOIPubMed
88. Szakács G, Paterson JK, Ludwig JA, Booth-Genthe C, Gottesman MM. Targeting multidrug resistance in cancer. Nat Rev Drug Discov 2006;5:219.DOIPubMed
89. Chuthapisith S, Eremin J, El-Sheemey M, Eremin O. Breast cancer chemoresistance: emerging importance of cancer stem cells. Surg Oncol 2010;19:27-32.DOIPubMed
90. Eyre R, Harvey I, Stemke-Hale K, Lennard TWJ, Tyson-Capper A, et al. Reversing paclitaxel resistance in ovarian cancer cells via inhibition of the ABCB1 expressing side population. Tumor Biol 2014;35:9879-92.DOIPubMed
91. Begicevic RR, Falasca M. ABC transporters in cancer stem cells: beyond chemoresistance. Int J Mol Sci 2017;18:2362.DOIPubMedPMC
92. Adamska A, Falasca M. ATP-binding cassette transporters in progression and clinical outcome of pancreatic cancer: What is the way forward? World J Gastroenterol 2018;24:3222-38.DOIPubMedPMC
93. Kubota N, Nishio K, Takeda Y, Ohmori T, Funayama Y, et al. Characterization of an etoposide-resistant human ovarian cancer cell line. Cancer Chemother Pharmacol 1994;34:183-90.DOIPubMed
94. Alpsoy A, Yasa S, Gunduz U. Etoposide resistance in MCF-7 breast cancer cell line is marked by multiple mechanisms. Biomed Pharmacother 2014;68:351-5.DOIPubMed
95. Solazzo M, Fantappiè O, D’Amico M, Sassoli C, Tani A, et al. Mitochondrial expression and functional activity of breast cancer resistance protein in different multiple drug-resistant cell lines. Cancer Res 2009;69:7235-42.DOIPubMed
96. Green DR, Reed JC. Mitochondria and apoptosis. Science 1998;281:1309-12.DOIPubMed
97. Wang X, Green DR, Reed JC. The expanding role of mitochondria in apoptosis. Genes Dev 1998;281:1309-12.
98. Liu R, Page C, Beidler DR, Wicha MS, Núñez G. Overexpression of Bcl-xL promotes chemotherapy resistance of mammary tumors in a syngeneic mouse model. Am J Pathol 1999;155:1861-7.DOIPubMedPMC
99. Tu Y, Xu FH, Liu J, Vescio R, Berenson J, et al. Upregulated expression of BCL-2 in multiple myeloma cells induced by exposure to doxorubicin, etoposide, and hydrogen peroxide. Blood 1996;88:1805-12.PubMed
100. Kutuk O, Letai A. Alteration of the mitochondrial apoptotic pathway is key to acquired paclitaxel resistance and can be reversed by ABT-737. Cancer Res 2008;68:7985-94.DOIPubMedPMC
101. Kang MH, Reynolds CP. Bcl-2 inhibitors: targeting mitochondrial apoptotic pathways in cancer therapy. Clin Cancer Res 2009;15:1126-32.DOIPubMedPMC
102. Adams JM, Cory S. The Bcl-2 apoptotic switch in cancer development and therapy. Oncogene 2007;26:1324-37.DOIPubMedPMC
103. Sartorius UA, Krammer PH. Upregulation of bcl-2 is involved in the mediation of chemotherapy resistance in human small cell lung cancer cell lines. Int J Cancer 2002;97:584-92.DOIPubMed
104. Real PJ, Sierra A, Juan Ad, Segovia JC, Lopez-Vega JM, et al. Resistance to chemotherapy via Stat3-dependent overexpression of Bcl-2 in metastatic breast cancer cells. Oncogene 2002;21:7611.DOIPubMed
105. Bauer JJ, Sesterhenn IA, Mostofi FK, McLeod DG, Srivastava S, et al. Elevated levels of apoptosis regulator proteins P53 and BCL-2 are independent prognostic biomarkers in surgically treated clinically localized prostate cancer. J Urology 1996;156:1511-6.DOIPubMed
106. Cho HJ, Kim JK, Kim KD, Yoon HK, Cho MY, et al. Upregulation of Bcl-2 is associated with cisplatin-resistance via inhibition of Bax translocation in human bladder cancer cells. Cancer Lett 2006;237:56-66.DOIPubMed
107. Amundson SA, Myers TG, Scudiero D, Kitada S, Reed JC, et al. An informatics approach identifying markers of chemosensitivity in human cancer cell lines. Cancer Res 2000;60:6101-10.PubMed
108. Biswas G, Guha M, Avadhani NG. Mitochondria-to-nucleus stress signaling in mammalian cells: nature of nuclear gene targets, transcription regulation, and induced resistance to apoptosis. Gene 2005;354:132-9.DOIPubMedPMC
109. Colié S, van Veldhoven PP, Kedjouar B, Bedia C, Albinet V, et al. Disruption of Sphingosine 1-Phosphate Lyase confers resistance to chemotherapy and promotes oncogenesis through Bcl-2/Bcl-xL upregulation. Cancer Res 2009;69:9346-53.DOIPubMed
110. Renault TT, Elkholi R, Bharti A, Chipuk JE. B cell lymphoma-2 (BCL-2) homology domain 3 (BH3) mimetics demonstrate differential activities dependent upon the functional repertoire of pro- and anti-apoptotic BCL-2 family proteins. J Biol Chem 2014;289:26481-91.DOIPubMedPMC
111. Vogler M, Dinsdale D, Dyer MJS, Cohen GM. Bcl-2 inhibitors: small molecules with a big impact on cancer therapy. Cell Death Differ 2009;16:360.DOIPubMed
112. Tse C, Shoemaker AR, Adickes J, Anderson MG, Chen J, et al. ABT-263: a potent and orally bioavailable Bcl-2 family inhibitor. Cancer Res 2008;68:3421-8.DOIPubMed
113. Oltersdorf T, Elmore SW, Shoemaker AR, Armstrong RC, Augeri DJ, et al. An inhibitor of Bcl-2 family proteins induces regression of solid tumours. Nature 2005;435:677-81.DOIPubMed
114. Samudio I, Harmancey R, Fiegl M, Kantarjian H, Konopleva M, et al. Pharmacologic inhibition of fatty acid oxidation sensitizes human leukemia cells to apoptosis induction. J Clin Invest 2010;120:142-56.DOIPubMedPMC
115. Steele TM, Talbott GC, Sam A, Tepper CG, Ghosh PM, et al. Obatoclax, a BH3 mimetic, enhances cisplatin-induced apoptosis and decreases the clonogenicity of muscle invasive bladder cancer cells via mechanisms that involve the inhibition of pro-survival molecules as well as cell cycle regulators. Int J Mol Sci 2019;20:1285.DOIPubMedPMC
116. Luca A de, Fiorillo M, Peiris-Pagès M, Ozsvari B, Smith DL, et al. Mitochondrial biogenesis is required for the anchorage-independent survival and propagation of stem-like cancer cells. Oncotarget 2015;6:14777-95.DOIPubMedPMC
117. Salem AF, Whitaker-Menezes D, Howell A, Sotgia F, Lisanti MP. Mitochondrial biogenesis in epithelial cancer cells promotes breast cancer tumor growth and confers autophagy resistance. Cell Cycle 2012;11:4174-80.DOIPubMedPMC
118. Carracedo A, Weiss D, Leliaert AK, Bhasin M, Boer VCJ de, et al. A metabolic prosurvival role for PML in breast cancer. J Clin Invest 2012;122:3088-100.DOIPubMedPMC
119. Martinez-Outschoorn UE, Pavlides S, Sotgia F, Lisanti MP. Mitochondrial biogenesis drives tumor cell proliferation. Am J Pathol 2011;178:1949-52.DOIPubMedPMC
120. Haq R, Shoag J, Andreu-Perez P, Yokoyama S, Edelman H, et al. Oncogenic BRAF regulates oxidative metabolism via PGC1α and MITF. Cancer Cell 2013;23:302-15.DOIPubMedPMC
121. Puigserver P, Wu Z, Park CW, Graves R, Wright M, et al. A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis. Cell 1998;92:829-39.DOIPubMed
122. Lin J, Handschin C, Spiegelman BM. Metabolic control through the PGC-1 family of transcription coactivators. Cell Metab 2005;1:361-70.DOIPubMed
123. Tan Z, Luo X, Xiao L, Tang M, Bode AM, et al. The role of PGC1α in cancer metabolism and its therapeutic implications. Mol Cancer Ther 2016;15:774-82.DOIPubMed
124. Shin SW, Yun SH, Park ES, Jeong JS, Kwak JY, et al. Overexpression of PGC-1α enhances cell proliferation and tumorigenesis of HEK293 cells through the upregulation of Sp1 and Acyl-CoA binding protein. Int J Oncol 2015;46:1328-42.DOIPubMed
125. Bhalla K, Hwang BJ, Dewi RE, Ou L, Twaddel W, et al. PGC1α promotes tumor growth by inducing gene expression programs supporting lipogenesis. Cancer Res 2011;71:6888-98.DOIPubMedPMC
126. Srivastava S, Barrett JN, Moraes CT. PGC-1alpha/beta upregulation is associated with improved oxidative phosphorylation in cells harboring nonsense mtDNA mutations. Hum Mol Genet 2007;16:993-1005.DOIPubMedPMC
127. Kim B, Jung JW, Jung J, Han Y, Suh DH, et al. PGC1α induced by reactive oxygen species contributes to chemoresistance of ovarian cancer cells. Oncotarget 2017;8:60299-311.DOIPubMedPMC
128. Gopal YNV, Rizos H, Chen G, Deng W, Frederick DT, et al. Inhibition of mTORC1/2 overcomes resistance to MAPK pathway inhibitors mediated by PGC1alpha and oxidative phosphorylation in melanoma. Cancer Res 2014;74:7037-47.DOIPubMedPMC
129. Gabrielson M, Björklund M, Carlson J, Shoshan M. Expression of mitochondrial regulators PGC1α and TFAM as putative markers of subtype and chemoresistance in epithelial ovarian carcinoma. PLoS One 2014;9:e107109.DOIPubMedPMC
130. Luo C, Widlund HR, Puigserver P. PGC-1 coactivators: shepherding the mitochondrial biogenesis of tumors. Trends Cancer 2016;2:619-31.DOIPubMedPMC
131. Yun CW, Han YS, Lee SH. PGC-1α controls mitochondrial biogenesis in drug-resistant colorectal cancer cells by regulating endoplasmic reticulum stress. Int J Mol Sci 2019;20:1707.DOIPubMedPMC
132. Douarre C, Sourbier C, Dalla Rosa I, Brata Das B, Redon CE, et al. Mitochondrial Topoisomerase I is critical for mitochondrial integrity and cellular energy metabolism. PLoS One 2012;7:e41094.DOIPubMedPMC
133. Kluza J, Marchetti P, Gallego MA, Lancel S, Fournier C, et al. Mitochondrial proliferation during apoptosis induced by anticancer agents: effects of doxorubicin and mitoxantrone on cancer and cardiac cells. Oncogene 2004;23:7018.DOIPubMed
134. Fu X, Wan S, Lyu YL, Liu LF, Qi H. Etoposide Induces ATM-dependent mitochondrial biogenesis through AMPK activation. PLoS One 2008;3:e2009.DOIPubMedPMC
135. Farnie G, Sotgia F, Lisanti MP. High mitochondrial mass identifies a sub-population of stem-like cancer cells that are chemo-resistant. Oncotarget 2015;6:30472-86.DOIPubMedPMC
136. Haq R, Shoag J, Andreu-Perez P, Yokoyama S, Edelman H, et al. Oncogenic BRAF regulates oxidative metabolism via PGC1α and MITF. Cancer Cell 2013;23:302-15.DOIPubMedPMC
137. Lamb R, Bonuccelli G, Ozsvári B, Peiris-Pagès M, Fiorillo M, et al. Mitochondrial mass, a new metabolic biomarker for stem-like cancer cells: understanding WNT/FGF-driven anabolic signaling. Oncotarget 2015;6:30453-71.DOIPubMedPMC
138. Rose PG, Mossbruger K, Fusco N, Smrekar M, Eaton S, et al. Gemcitabine reverses cisplatin resistance: demonstration of activity in platinum- and multidrug-resistant ovarian and peritoneal carcinoma. Gynecol Oncol 2003;88:17-21.DOIPubMed
139. Rudin CM, Yang Z, Schumaker LM, VanderWeele DJ, Newkirk K, et al. Inhibition of glutathione synthesis reverses Bcl-2-mediated cisplatin resistance. Cancer Res 2003;63:312-8.PubMed
Lauren K. Meyer et al., Cancer Drug Resistance, 2019
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