Oxidative Medicine and Cellular Longevity

Oxidative Medicine and Cellular Longevity / 2020 / Article

Research Article | Open Access

Volume 2020 |Article ID 4971525 | https://doi.org/10.1155/2020/4971525

Leslie Amaral, Filipa Mendes, Manuela Côrte-Real, Maria João Sousa, Susana Rodrigues Chaves, "Lactate Induces Cisplatin Resistance in S. cerevisiae through a Rad4p-Dependent Process", Oxidative Medicine and Cellular Longevity, vol. 2020, Article ID 4971525, 8 pages, 2020. https://doi.org/10.1155/2020/4971525

Lactate Induces Cisplatin Resistance in S. cerevisiae through a Rad4p-Dependent Process

Academic Editor: Kum Kum Khanna
Received20 Jan 2020
Revised09 Sep 2020
Accepted30 Sep 2020
Published24 Oct 2020

Abstract

Cisplatin is a widely used antineoplastic agent that has DNA as the main target, though cellular resistance hampers its therapeutic efficacy. An emerging hallmark of cancer cells is their altered metabolism, characterized by increased glycolysis even under aerobic conditions, with increased lactate production (known as the Warburg effect). Although this altered metabolism often results in increased resistance to chemotherapy, it also provides an opportunity for targeted therapeutic intervention. It has been suggested that cisplatin cytotoxicity can be affected by tumor metabolism, though with varying effects. We therefore sought to better characterize how lactate affects cisplatin sensitivity in the simplified Saccharomyces cerevisiae model. We show that lactate renders yeast cells resistant to cisplatin, independently of growth rate or respiration ability. We further show that histone acetylation is not affected, but histone phosphorylation is decreased in lactate-containing media. Finally, we show that Rad4p, essential for nucleotide excision repair, is required for the observed phenotype and thus likely underlies the mechanism responsible for lactate-mediated resistance to cisplatin. Overall, understanding how lactate modulates cisplatin sensitivity will aid in the development of new strategies to overcome drug resistance.

1. Introduction

Cisplatin is a platinum-based drug used in the treatment of various solid neoplasms, such as head and neck, lung, colorectal, and ovarian cancers [1]. This chemotherapeutic drug is administered either alone or combined with other drugs [2] and, depending on the dose, can induce irreparable DNA damage leading to cell cycle arrest or cell death, usually by apoptosis [3]. Despite its effectiveness, cellular resistance still occurs, either intrinsic or acquired [4]. Resistance is a multifactorial problem, and the most studied mechanisms range from defects in mismatch repair, aneuploidy, increased cisplatin detoxification, and failure to undergo apoptosis [5, 6]. More recently, increased attention has been given to the metabolism of tumor cells and how it affects chemoresistance. Unlike normal cells, cancer cells predominantly present a glycolytic profile even in the presence of oxygen, associated with high uptake of glucose and lactic acid production [7, 8], known as the Warburg effect or “aerobic glycolysis” [9, 10]. It is postulated that this type of metabolism can lead to a faster production of ATP and accumulation of bioprecursors to fit cellular demand [11] but may also influence the response of cancer cells to therapy [12]. Indeed, a high lactate concentration in tumors is associated with bad prognosis [13]. However, the connection between metabolism and response to therapy is not always clear-cut. Indeed, tumors are inherently heterogeneous and can include both oxidative and glycolytic cells, depending, among other factors, on vascularization and oxygen availability (reviewed in [14, 15]). A “Reverse Warburg Effect” has also been proposed, whereby cells in the tumor microenvironment are induced to provide metabolites to fuel cancer cells. For instance, lactate released by glycolytic cells can be recaptured by oxidative cancer cells to use as fuel, but also function as a signaling molecule affecting chemosensitivity (reviewed in [16, 17]) [18]. Nonetheless, results are often contradictory. In particular, it has been reported that treatment with cisplatin can lead to inhibition of glycolysis [19], and that inhibition of glucose metabolism can increase sensitivity to the drug, often through increased ROS levels [20, 21]. Cisplatin treatment also decreases mitochondrial respiration in in vitro cultures of HT-29, HCT116, HepG2, and MDA-MB-231, followed by a decrease in glycolysis [22]. However, while some cisplatin-resistant cell lines display higher rates of glycolysis [2326], others have increased oxidative metabolism [27, 28]. This indicates that the relation between metabolism and cisplatin sensitivity depends on the type of tumor or cell line under study, as well as experimental conditions. Since we have previously shown that cisplatin induces an active, but mitochondrial-independent cell death process in Saccharomyces cerevisiae [29], we took advantage of this simpler model to clarify the role of lactate in cisplatin-induced cell death.

2. Materials and Methods

2.1. Growth Conditions and Treatments

The Saccharomyces cerevisiae strains used in the experimental procedures and respective genotypes are listed in Table 1. Cells were grown overnight, at 30°C and 200 rpm, in synthetic complete (SC) medium (5.0 g/L ammonium sulphate ((NH4)2SO4), 1.4 g/L dropout mix, 1.7 g/L yeast nitrogen base w/o amino acids and w/o ammonium sulphate ((NH4)2SO4), and 2% glucose (w/v)). After overnight growth, cells were collected at OD (A640) 0.5-0.7 and transferred to new SC medium containing 2% of the indicated carbon source (glucose or lactate). The pH of all media was adjusted to 5.0-5.5. Cisplatin (CDDP; Sigma) was added afterwards to a final concentration of 600 μg/mL or 450 μg/mL. Cisplatin was stored in aliquots in the dark and resuspended in DMSO immediately prior to use. However, although the phenotype in relation to control was consistent, some variability in cytotoxicity between independent experiments and different cisplatin lots was unavoidable. Methyl methanesulfonate (MMS; Fluka) was used as a positive control at a final concentration of 0.1%. α-mating factor was used at a final concentration of 1 mg/mL in glucose-containing media, to decrease the specific growth rate by arresting cells at the G1 cell cycle phase.


StrainGenotypeReference

BY4741MATa his3Δ1; leu2Δ0; met15Δ0; ura3Δ0[42]
rho0BY4741 rho0A. Rego
bar1ΔBY4741 bar1Δ::KanMX4Euroscarf (Germany)
rad4ΔBY4741 rad4Δ::URA3This study

2.2. Viability Assays

For semiquantitative viability assays, 5 μL of cell suspensions and tenfold serial dilutions (10-1 to 10-4) were spotted on YPD plates (1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) glucose, and 2% (w/v) agar). After 2 days of incubation at 30°C, all plates were photographed using Chemidoc XRS (BioRad) and analyzed by Quantity One (BioRad).

For quantitative viability assays, 40 μL of cell suspensions were spotted on YPD plates. After 2 days of incubation at 30°C, colony-forming units (c.f.u.) were counted and normalized to the OD (A640) values. The percentage of cell viability was then calculated in relation to T0 (considered as 100% of cell viability). Results are expressed as . Each experiment was performed at least 3 times. GraphPad Prism 5 was used for statistical analysis.

2.3. Western Blot

Total cellular extracts were separated in a 12.5% polyacrylamide gel (SDS-PAGE). Following electrophoresis, proteins were transferred onto a nitrocellulose membrane at 60 mA for 90 minutes, using a semidry system (TE77X, Hoefer, USA). Membranes were blocked for 1 h in 5% nonfat dry milk and diluted in PBS-T (PBS containing 0.05% (v/v) Tween-20), under low agitation. Subsequently, for phosphorylated H2A, H3 acetylation, and Pgk1p detection, membranes were incubated overnight at 4°C with the primary polyclonal antibodies anti-γH2AX (1 : 5000, Abcam), anti-acetyl-Histone H3 (Lys9) (1 : 5000, Millipore), or anti-PGK1 (1 : 5000, Molecular Probes), respectively. The next day, membranes were washed minutes with PBS-T, followed by a 1 h room temperature incubation with the secondary antibodies anti-rabbit (for γH2AX and acetyl-Histone H3) or anti-mouse (for Pgk1p) from Jackson Laboratories. Chemiluminescence detection was performed using the Immobilon ECL detection system (Millipore-Merck) and a Chemi-DOC XRS system (BioRad) or X-ray ortho CP-G films in an X-ray film processor (Curix 60, Agfa Healthcare).

3. Results and Discussion

3.1. Lactate Protects S. cerevisiae Cells from Cisplatin-Induced Cell Death

To uncover the influence of lactate on cisplatin sensitivity, we assessed the viability of BY4741 cells exposed to cisplatin in medium containing glucose or lactate for up to 180 min. As a control, we also assessed viability of cells exposed to methyl methanesulfonate (MMS), a DNA alkylating agent [30]. As seen in Figure 1, lactate increased viability of cells exposed to cisplatin, but did not seemingly affect viability of cells exposed to MMS. This indicates that lactate specifically protects cells from cisplatin exposure, which may be contingent on the type of DNA damage or the pathways involved in its repair.

Indeed, MMS is a DNA-alkylating agent that causes alterations in guanine and adenine residues, resulting in base mispairing, and is predominantly repaired by the base excision repair (BER) pathway and DNA alkyltransferase [31]. In contrast, intrastrand crosslinks caused by cisplatin are mainly repaired by the nucleotide excision repair (NER) pathway [32].

Another contributing factor that may influence resistance to cisplatin could be the slower growth rate of yeast cells. Indeed, cells grow slower in lactate-containing media, which could result in additional time to repair DNA damage induced by cisplatin and underlie the observed increased resistance. To address this hypothesis, we assessed whether decreasing the growth rate in glucose-containing media affected cisplatin resistance. For this purpose, BY4741 bar1Δ cells were grown overnight in glucose-containing medium, pretreated with α-factor and exposed to cisplatin for 180 min (the BAR1 deletion was used to decrease α-factor degradation). As seen in Figure 2, α-factor decreased the specific growth rate over the 3 h of cisplatin exposure to a level comparable or even lower to that of cells grown in lactate-containing media, consistent with a cell cycle arrest during this period. However, similar viability was observed in cells exposed to cisplatin in the presence or absence of α-factor (with a tendency for lower viability in the presence of α-factor that was not significant). This indicates that it is not the decreased growth rate but the presence of lactate that increases cisplatin resistance.

Apart from the decreased growth rate, cells grown in lactate media also display a different metabolic status, i.e., cells switch to a respiratory metabolism. As referred above, several studies report a connection between cisplatin and oxidative metabolism, though the mechanism and effects are complex. On one hand, studies show that exposure to cisplatin can itself lead to increased oxidative phosphorylation, suggesting that inhibition of tumor glycolytic metabolism could underlie decreased proliferation in response to the drug [19, 33]. On the other hand, although cisplatin-resistant and cisplatin-sensitive cells tend to display different energy metabolism, opposing effects are often found (reviewed in [12]). For instance, cisplatin-resistant ovarian and cervical cancer cells had higher rates of glycolysis whereas cisplatin-resistant lung cancer cells displayed increased oxidative phosphorylation in comparison with their cisplatin-sensitive counterparts [2428]. To assess whether respiration affected cisplatin resistance in yeast cells, we determined whether the protective effect of lactate was still observed in the respiratory-deficient rho0 strain, which lacks mitochondrial DNA.

As seen in Figure 3, rho0 cells were still more resistant to cisplatin in lactate-containing media, as observed in wild-type BY4741 cells. This indicates that it is not respiration per se that increases cisplatin resistance when shifting from fermentative to nonfermentative carbon sources during cisplatin exposure. These results therefore suggest that it is the lactate signaling role, in the modulation of processes other from respiration, that protects cells from cisplatin-induced death. Indeed, lactate is increasingly regarded as not just the end result of the Warburg effect but as an important molecule in driving carcinogenesis [710] (reviewed in [34]). While some of its effects are specific to multicellular organisms (such as increased angiogenesis), others occur at the cellular level, such as the effects of lactate on chromatin and transcription.

3.2. Cisplatin-Induced DNA Damage Response Is Decreased in Lactate-Containing Medium

It has previously been described that lactate is a weak histone deacetylase (HDAC) inhibitor in HCT116 cells [35], resulting in increased levels of histone acetylation, associated with more relaxed chromatin. Histone acetylation has also been implicated in the regulation of the DNA damage response (DDR) in mammalian cells, and lactate has been shown to increase chromatin accessibility and DNA repair in cervical cancer cells [36]. We therefore sought to determine whether lactate could alter the levels of histone acetylation in yeast cells. For this purpose, we assessed the levels of H3 acetylation in extracts of cells exposed to cisplatin in glucose or lactate-containing media, by Western blot (Figure 4).

We could not detect any increase in H3 acetylation levels under our experimental conditions, suggesting that the observed cisplatin resistance in the presence of lactate is not related with increased histone acetylation. Though lactate did not seem to act by epigenetic regulation of chromatin in this case, it could still affect DDR through other mechanisms. One particularly well-known and early event that occurs after DNA damage is the phosphorylation of histone H2A or H2AX, in higher eukaryotes. In yeast, it is well established that H2A is phosphorylated in response to DNA damage, as we have previously shown in response to cisplatin [37]. We therefore assessed the levels of H2A phosphorylation in extracts of cells exposed to cisplatin in glucose or lactate-containing media, by Western blot (Figure 5).

We observed that the levels of phospho-H2A were much lower in the latter, suggestive of lower levels of damaged DNA or inhibition of the DNA damage response. However, decreased DDR should not result in increased viability; quite on the contrary, it usually results in increased sensitivity to DNA damage [38, 39]. It is, therefore, more likely that exposure to cisplatin in the presence of lactate results in decreased initial DNA damage or increased DNA repair in yeast cells.

3.3. Nucleotide Excision Repair Is Required for Lactate-Induced Cisplatin Resistance

Taken together, the previous results indicate that the presence of lactate results in decreased initial cisplatin DNA damage or increased damage repair, which is difficult to distinguish biochemically. We therefore sought to determine whether DNA repair was required for the observed phenotype. As mentioned above, the DNA damage induced by platinum drugs is mainly repaired by the NER pathway [40], and it is well established that Rad4p is indispensable for NER activity [41]. We therefore deleted the RAD4 gene in BY4741 cells and assessed whether lactate still rescued rad4∆ cells from cisplatin-induced cell death (Figure 6).

Because rad4∆ cells are highly sensitive to cisplatin, a lower concentration of cisplatin was also used. For both concentrations, it was possible to observe that lactate did not greatly increase the resistance of rad4∆ cells to cisplatin (Figure 6). Indeed, the percentage of surviving wild-type cells was significantly greater in lactate media than in glucose media after 180 min of cisplatin exposure (panels b and c). In contrast, only a minor increase was observed in the rad4∆ strain (measured at the 120 min time point since, after 180 min, there were virtually no viable cells exposed to either concentration of cisplatin). Taken together, though a contribution from another mechanism cannot be excluded, these results indicate that the NER pathway plays an important role in the protection from cisplatin-induced cell death imparted by lactate.

4. Conclusions

In this work, we show that lactate specifically increases resistance to cisplatin, independently of the cellular growth rate and respiratory capacity. This process was associated with decreased DNA damage, probably through increased NER, as Rad4p was required for the observed resistance. It is well established that repair-deficient tumor cells are particularly sensitive to cisplatin, which has been attributed to a decreased capacity to repair the DNA lesions. Our work suggests that NER deficiency may further contribute to chemotherapy efficacy by attenuating acquired resistance of tumor cells in a lactate-containing microenvironment, which warrants further investigation.

Data Availability

The data used to support the findings of this study are available from the corresponding author upon request.

Conflicts of Interest

The authors declare that they have no conflicts of interest.

Authors’ Contributions

Leslie Amaral and Filipa Mendes contributed equally to this work.

Acknowledgments

This work was supported by the "Contrato-Programa" (UID/BIA/04050/2020) funded by national funds through FCT I.P., as well as a Post-doc fellowship to S. Chaves (SFRH/BPD/89980/2012).

References

  1. M. Galanski, “Recent developments in the field of anticancer platinum complexes,” Recent Patents on Anti-Cancer Drug Discovery, vol. 1, no. 2, pp. 285–295, 2006. View at: Publisher Site | Google Scholar
  2. A. R. Timerbaev, C. G. Hartinger, S. S. Aleksenko, and B. K. Keppler, “Interactions of antitumor metallodrugs with serum proteins: advances in characterization using modern analytical methodology,” Chemical Reviews, vol. 106, no. 6, pp. 2224–2248, 2006. View at: Publisher Site | Google Scholar
  3. J. Campisi, “Aging, cellular senescence, and cancer,” Annual Review of Physiology, vol. 75, no. 1, pp. 685–705, 2013. View at: Publisher Site | Google Scholar
  4. D. W. Shen, L. M. Pouliot, M. D. Hall, and M. M. Gottesman, “Cisplatin resistance: a cellular self-defense mechanism resulting from multiple epigenetic and genetic changes,” Pharmacological Reviews, vol. 64, no. 3, pp. 706–721, 2012. View at: Publisher Site | Google Scholar
  5. L. H. Einhorn, “Curing metastatic testicular cancer,” Proceedings of the National Academy of Sciences of the United States of America, vol. 99, no. 7, pp. 4592–4595, 2002. View at: Publisher Site | Google Scholar
  6. R. P. Perez, “Cellular and molecular determinants of cisplatin resistance,” European Journal of Cancer, vol. 34, no. 10, pp. 1535–1542, 1998. View at: Publisher Site | Google Scholar
  7. O. Warburg, “On the origin of cancer cells,” vol. 123, no. 3, pp. 412–421, 1956. View at: Google Scholar
  8. R. A. Gatenby and R. J. Gillies, “Why do cancers have high aerobic glycolysis?” Nature Reviews. Cancer, vol. 4, no. 11, pp. 891–899, 2004. View at: Publisher Site | Google Scholar
  9. D. Hanahan and R. A. Weinberg, “Hallmarks of cancer: the next generation,” Cell, vol. 144, no. 5, pp. 646–674, 2011. View at: Publisher Site | Google Scholar
  10. R. Cairns, I. Harris, and T. Mak, “Regulation of cancer cell metabolism,” Nature Reviews. Cancer, vol. 11, no. 2, pp. 85–95, 2011. View at: Publisher Site | Google Scholar
  11. M. G. Vander Heiden, L. C. Cantley, and C. B. Thompson, “Understanding the Warburg effect: the metabolic requirements of cell proliferation,” Science, vol. 324, no. 5930, pp. 1029–1033, 2009. View at: Publisher Site | Google Scholar
  12. E. A. Zaal and C. R. Berkers, “The influence of metabolism on drug response in cancer,” Frontiers in Oncology, vol. 8, pp. 1–15, 2018. View at: Publisher Site | Google Scholar
  13. P. J. Vlachostergios, K. G. Oikonomou, E. Gibilaro, and G. Apergis, “Elevated lactic acid is a negative prognostic factor in metastatic lung cancer,” Cancer Biomarkers, vol. 15, no. 6, pp. 725–734, 2015. View at: Publisher Site | Google Scholar
  14. F. D. S. E. Melo, L. Vermeulen, E. Fessler, and J. P. Medema, “Cancer heterogeneity—a multifaceted view,” EMBO Reports, vol. 14, no. 8, pp. 686–695, 2013. View at: Publisher Site | Google Scholar
  15. X. X. Sun and Q. Yu, “Intra-tumor heterogeneity of cancer cells and its implications for cancer treatment,” Acta Pharmacologica Sinica, vol. 36, no. 10, pp. 1219–1227, 2015. View at: Publisher Site | Google Scholar
  16. L. Wilde, M. Roche, M. Domingo-Vidal et al., “Metabolic coupling and the reverse Warburg effect in cancer: implications for novel biomarker and anticancer agent development,” Seminars in Oncology, vol. 44, no. 3, pp. 198–203, 2017. View at: Publisher Site | Google Scholar
  17. S. Romero-Garcia, M. M. B. Moreno-Altamirano, H. Prado-Garcia, and F. J. Sánchez-García, “Lactate contribution to the tumor microenvironment: mechanisms, effects on immune cells and therapeutic relevance,” Frontiers in Immunology, vol. 7, 2016. View at: Publisher Site | Google Scholar
  18. P. Sonveaux, F. Végran, T. Schroeder et al., “Targeting lactate-fueled respiration selectively kills hypoxic tumor cells in mice,” The Journal of Clinical Investigation, vol. 118, no. 12, pp. 3930–3942, 2008. View at: Publisher Site | Google Scholar
  19. M. V. Shirmanova, I. N. Druzhkova, M. M. Lukina et al., “Chemotherapy with cisplatin: insights into intracellular pH and metabolic landscape of cancer cells in vitro and in vivo,” Scientific Reports, vol. 7, no. 1, pp. 1–13, 2017. View at: Google Scholar
  20. P. Loar, H. Wahl, M. Kshirsagar, G. Gossner, K. Griffith, and J. R. Liu, “Inhibition of glycolysis enhances cisplatin-induced apoptosis in ovarian cancer cells,” American Journal of Obstetrics and Gynecology, vol. 202, no. 4, pp. 371–379, 2010. View at: Google Scholar
  21. W. H. Gotlieb, J. Saumet, M.-C. Beauchamp et al., “In vitro metformin anti-neoplastic activity in epithelial ovarian cancer,” Gynecologic Oncology, vol. 110, no. 2, pp. 246–250, 2008. View at: Publisher Site | Google Scholar
  22. H. Alborzinia, S. Can, P. Holenya et al., “Real-time monitoring of cisplatin-induced cell death,” PLoS One, vol. 6, no. 5, article e19714, 2011. View at: Publisher Site | Google Scholar
  23. R. Rattan, R. P. Graham, J. L. Maguire, S. Giri, and V. Shridhar, “Metformin suppresses ovarian cancer growth and metastasis with enhancement of cisplatin cytotoxicity in vivo,” Neoplasia, vol. 13, no. 5, pp. 483–IN28, 2011. View at: Publisher Site | Google Scholar
  24. C. Xintaropoulou, C. Ward, A. Wise et al., “Expression of glycolytic enzymes in ovarian cancers and evaluation of the glycolytic pathway as a strategy for ovarian cancer treatment,” BMC Cancer, vol. 18, no. 1, pp. 1–15, 2018. View at: Google Scholar
  25. R. Rashmi, X. Huang, J. M. Floberg et al., “Radioresistant cervical cancers are sensitive to inhibition of glycolysis and redox metabolism,” Cancer Research, vol. 78, no. 6, pp. 1392–1403, 2018. View at: Publisher Site | Google Scholar
  26. G. R. Leisching, B. Loos, M. H. Botha, and A.-M. Engelbrecht, “The role of mTOR during cisplatin treatment in an in vitro and ex vivo model of cervical cancer,” Toxicology, vol. 335, pp. 72–78, 2015. View at: Publisher Site | Google Scholar
  27. M. Wangpaichitr, C. Wu, Y. Y. Li et al., “Exploiting ROS and metabolic differences to kill cisplatin resistant lung cancer,” Oncotarget, vol. 8, no. 30, pp. 49275–49292, 2017. View at: Publisher Site | Google Scholar
  28. L. G. F. Medhi Wangpaichitr, G. Theodoropoulos, C. Wu, M. You, N. S. Macus, and T. Kuo, “The relationship of thioredoxin-1 and cisplatin resistance: its impact on ROS and oxidative metabolism in lung cancer cells,” Molecular Cancer Therapeutics, vol. 11, no. 3, pp. 604–615, 2013. View at: Google Scholar
  29. D. Cunha, R. Cunha, M. Côrte-Real, and S. R. Chaves, “Cisplatin-induced cell death in Saccharomyces cerevisiae is programmed and rescued by proteasome inhibition,” DNA Repair, vol. 12, no. 6, pp. 444–449, 2013. View at: Publisher Site | Google Scholar
  30. G. Evensen and E. Seeberg, “Adaptation to alkylation resistance involves the induction of a DNA glycosylase,” Nature, vol. 296, no. 5859, pp. 773–775, 1982. View at: Publisher Site | Google Scholar
  31. T. Lindahl and R. D. Wood, “Quality control by DNA repair,” Science, vol. 286, no. 5446, pp. 1897–1905, 1999. View at: Publisher Site | Google Scholar
  32. L. P. Martin, T. C. Hamilton, and R. J. Schilder, “Platinum resistance: the role of DNA repair pathways,” Clinical Cancer Research, vol. 14, no. 5, pp. 1291–1295, 2008. View at: Publisher Site | Google Scholar
  33. A. Cruz-Bermúdez, R. Laza-Briviesca, R. J. Vicente-Blanco et al., “Cisplatin resistance involves a metabolic reprogramming through ROS and PGC-1α in NSCLC which can be overcome by OXPHOS inhibition,” Free Radical Biology and Medicine, vol. 135, pp. 167–181, 2019. View at: Publisher Site | Google Scholar
  34. I. San-Millán and G. A. Brooks, “Reexamining cancer metabolism: lactate production for carcinogenesis could be the purpose and explanation of the Warburg effect,” Carcinogenesis, vol. 38, no. 2, pp. 119–133, 2016. View at: Publisher Site | Google Scholar
  35. T. Latham, L. Mackay, D. Sproul et al., “Lactate, a product of glycolytic metabolism, inhibits histone deacetylase activity and promotes changes in gene expression,” Nucleic Acids Research, vol. 40, no. 11, pp. 4794–4803, 2012. View at: Publisher Site | Google Scholar
  36. W. Wagner, W. M. Ciszewski, and K. D. Kania, “L- and D-lactate enhance DNA repair and modulate the resistance of cervical carcinoma cells to anticancer drugs via histone deacetylase inhibition and hydroxycarboxylic acid receptor 1 activation,” Cell Communication and Signaling: CCS, vol. 13, no. 1, pp. 1–16, 2015. View at: Google Scholar
  37. A. R. Costa, N. Machado, A. Rego, M. J. Sousa, M. Côrte-Real, and S. R. Chaves, “Proteasome inhibition prevents cell death induced by the chemotherapeutic agent cisplatin downstream of DNA damage,” DNA Repair, vol. 73, pp. 28–33, 2019. View at: Publisher Site | Google Scholar
  38. V. G. Gorgoulis, L.-V. F. Vassiliou, P. Karakaidos et al., “Activation of the DNA damage checkpoint and genomic instability in human precancerous lesions,” Nature, vol. 434, no. 7035, pp. 907–913, 2005. View at: Publisher Site | Google Scholar
  39. S. Sun, M. D. Osterman, and M. Li, “Tissue specificity of DNA damage response and tumorigenesis,” Cancer Biology & Medicine, vol. 16, no. 3, pp. 396–414, 2019. View at: Publisher Site | Google Scholar
  40. J. T. Reardon, A. Vaisman, S. G. Chaney, and A. Sancar, “Efficient nucleotide excision repair of cisplatin, oxaliplatin, and Bis-aceto-ammine-dichloro-cyclohexylamine-platinum(IV) (JM216) platinum intrastrand DNA diadducts,” Cancer Research, vol. 59, no. 16, pp. 3968–3971, 1999. View at: Google Scholar
  41. S. Boiteux and S. Jinks-Robertson, “DNA repair mechanisms and the bypass of DNA damage in Saccharomyces cerevisiae,” Genetics, vol. 193, no. 4, pp. 1025–1064, 2013. View at: Publisher Site | Google Scholar
  42. C. B. Brachmann, A. Davies, G. J. Cost, and E. Caputo, “Designer deletion strains derived fromSaccharomyces cerevisiae S288C: a useful set of strains and plasmids for PCR-mediated gene disruption and other applications,” Yeast, vol. 14, no. 2, pp. 115–132, 1998. View at: Publisher Site | Google Scholar

Copyright © 2020 Leslie Amaral et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


More related articles

 PDF Download Citation Citation
 Download other formatsMore
 Order printed copiesOrder
Views210
Downloads269
Citations

Related articles

Article of the Year Award: Outstanding research contributions of 2020, as selected by our Chief Editors. Read the winning articles.