Posts Tagged ‘LOW GRADE’

Accumulation of 2-hydroxyglutarate is not a biomarker for malignant progression of IDH-mutated low grade gliomas 

Larry H Bernstein, MD, FCAP, Curator


UPDATED 9/26/2021

Blockade of Glutathione Metabolism in IDH1-Mutated Glioma

Xiaoying TangXiao FuYang LiuDi YuSabrina J. Cai and Chunzhang Yang

source: https://mct.aacrjournals.org/content/19/1/221


Mutations in genes encoding isocitrate dehydrogenases (IDH) 1 and 2 are common cancer-related genetic abnormalities. Malignancies with mutated IDHs exhibit similar pathogenesis, metabolic pattern, and resistance signature. However, an effective therapy against IDH1-mutated solid tumor remains unavailable. In this study, we showed that acquisition of IDH1 mutation results in the disruption of NADP+/NADPH balance and an increased demand for glutathione (GSH) metabolism. Moreover, the nuclear factor erythroid 2–related factor 2 (Nrf2) plays a key protective role in IDH1-mutated cells by prompting GSH synthesis and reactive oxygen species scavenging. Pharmacologic inhibition of the Nrf2/GSH pathway via brusatol administration exhibited a potent tumor suppressive effect on IDH1-mutated cancer in vitro and in vivo. Our findings highlight a possible therapeutic strategy that could be valuable for IDH1-mutated cancer treatment.


Isocitrate dehydrogenases (IDH) are a family of enzymes that mediate the oxidative decarboxylation of isocitrate to α-ketoglutarate. These enzymes depend on NAD+/NADP+, as they reduce NAD+/NADP+ for NADH/NADPH production (1). While IDH1 and IDH2 isozymes are homodimers that use NADP+ as a cofactor, IDH3 is a heterotetramer that uses NAD+ as a cofactor (2). Genetic abnormalities in IDH1/2 are common in multiple types of human tumors. For example, mutations in IDH1/2 have been found in over 80% of World Health Organization grade II/III gliomas, including astrocytoma and oligodendroglioma (3). These mutations are found in 73% of secondary glioblastomas, which are derived from lower grade gliomas, but are less frequent in primary glioblastoma multiforme (4). Furthermore, the IDH1/2 mutations are commonly identified in acute myeloid leukemia (AML), central chondrosarcoma, central/periosteal chondromas, and cholangiocarcinoma (5, 6). However, despite the widespread prevalence of these mutations, effective therapies for IDH-mutated solid tumors remain unavailable.

The majority of cancer-associated IDH mutations are amino acid substitutions of an arginine residue in its catalytic center. For IDH1, the 132 arginine (R) residue is frequently altered to histidine (H) or cysteine (C). The R132H (73.67%) and R132C (13.35%) variant comprise over 87% of all IDH1 mutations in human. A seminal study by Dang and colleagues (7) revealed that these amino acid substitutions in IDH1 lead to a neomorphic activity of the enzyme, which is the NADPH-dependent consumption of α-ketoglutarate for 2-hydroxyglutarate (2-HG) production. The accumulation of 2-HG has been reported to associate with glioma oncogenesis by inhibiting α-ketoglutarate dioxygenases (8–10). Several pioneer studies also suggest that IDH mutants are associated with depletion of NADPH and glutathione (GSH), accompanied with elevated reactive oxygen species (ROS) levels (11, 12). The neomorphic catalytic function, 2-HG accumulation and occurrence of oxidative stress, suggest a distinctive oncogenesis mechanism, which could be exploited as a therapeutic vulnerability in IDH1-mutated malignancies.

In this study, we investigated the association between mutant IDH1 enzyme and ROS levels. Furthermore, we analyzed the role of nuclear factor erythroid 2–related factor 2 (Nrf2) in the regulation of GSH metabolism that maintains cellular redox homeostasis and survival. Moreover, we investigated the efficacy of Nrf2/GSH metabolism blockade as a therapeutic approach in IDH1-mutated malignancies.


Neomorphic activity in cancer-associated mutant IDH1 triggers oxidative stress

To better understand the effect of IDH1 mutants on redox homeostasis, we established a doxycycline-induced IDH1-mutant U251 cell line (Supplementary Fig. S1A). We noticed that upon the expression of mutant IDH1 enzymes, the overall quantity of NADP decreased, in both of its oxidized (NADP+) and reduced (NADPH) forms (Fig. 1A). Moreover, the NADP+/NADPH ratio significantly increased (Fig. 1B), suggesting that the neomorphic enzyme activity of mutant IDH1 exhausted the cellular pool of NADPH. The balance between NADP+ and NADPH is a critical factor to maintain cellular redox homeostasis, as NADPH is a general cofactor in reductive biosynthetic reactions, and to provide electrons for metabolic pathways, such as the reduction of GSSG back to GSH and ROS neutralization (23). Furthermore, by quantification of H2O2, we showed that mutant IDH1 enzymes led to severe oxidative stress in both U251 cells and BTIC TS603 (Fig. 1C; Supplementary Fig. S2A). Such an increase in ROS levels depended on the presence of mutant enzymes. The treatment with AGI-5198, a specific inhibitor of mutant IDH1 (13), reduced ROS accumulation in cells expressing the IDH1 R132H variant (Fig. 1D). Furthermore, the elevated ROS levels led to oxidative stress to macromolecules and subcellular organelles, evidenced by increased lipid peroxidation (Fig. 1E and F) and mitochondrial ROS level (Fig. 1G and H). The ROS scavenger catalase and MnTBAP, but not mannitol, abrogated the accumulation of ROS in IDH1-mutated cells, indicating the majority form of oxidative stress is derived from hydrogen peroxide and superoxide anion (Supplementary Fig. S1B).

Figure 1.

Cancer-associated IDH1 mutants trigger oxidative stress. A, NADP+ level was measured in U251 cells with doxycycline (Dox)-induced expression of IDH1-mutant enzymes (R132C and R132H). **, P < 0.01. B, Measurement of NADP+/NADPH ratio in U251 cells with expression of IDH1-mutant enzymes. **, P < 0.01. C, ROS-Glo measurement in U251 cells with expression of IDH1-mutant enzymes. **, P < 0.01. D, ROS-Glo measurement in IDH1-R132H U251 cells with AGI-5198 treatment (1 μmol/L, 24 hours; **, P < 0.01). E, Lipid peroxidation staining measures membrane oxidative damage in U251 cells with expression of IDH1-mutant enzymes. Scale bar, 10 μm. F, Quantification of lipid peroxidation in E. **, P < 0.01. G, MitoSOX staining measures mitochondrial ROS in IDH1-mutated U251 cells. Scale bar, 10 μm. H, Quantification of MitoSOX signal in G. **, P < 0.01.

GSH metabolism supports redox balance and survival in IDH1-mutated cells

Considering the remarkable ROS accumulation in IDH1-mutated cells, we speculate that antioxidant pathways, such as GSH-dependent ROS scavenging systems, may be triggered to maintain redox homeostasis. To test this hypothesis, we first measured the protein levels of GSH synthesis enzymes by Western blotting. We found that upon introduction of pathogenic mutant IDH1 enzymes, the levels of key enzymes in GSH biosynthesis, such as glutamate-cysteine ligase (GCLC, catalytic subunit; GCLM, modifier subunit), and cystine/glutamate transporter (SLC7A11, xCT transporter), increased (Fig. 2A). We also noticed that the levels of Nrf2 protein (NFE2L2), the major transcriptional factor responsible for ROS sensing, increased in IDH1-mutated cells. The enhancement of Nrf2 and GSH-dependent ROS scavenging pathways in IDH1-mutated U251 cells, as well as IDH1-mutated BTIC TS603, was also confirmed by qPCR (Fig. 2B; Supplementary Fig. S2B). To further understand the role of GSH in IDH1-mutated cells, we quantified GSH/GSSG levels in U251 cells expressing R132C/H IDH1 mutants. We recorded a substantial decrease in the GSH/GSSG ratio compared with that in cells expressing wild-type IDH1, suggesting that there is an elevated demand from GSH-dependent ROS scavenging (Fig. 2C). We also measured GSH/GSSG levels in BTICs, the result consistently showed that IDH1-mutated BTIC TS603 has lower GSH/GSSG ratio compared with IDH1 wild-type BTIC GSC827 and GSC 923 (Supplementary Fig. S2C). Moreover, the addition of an exogenous antioxidant enzyme, catalase, partially restored the GSH/GSSG ratio, indicating that the GSH/GSSG imbalance could be a result of GSH oxidation by hydrogen peroxide decomposition pathways (e.g., GSH peroxidases). Importantly, GSH biosynthesis exhibited a critical protective role in cells with mutant IDH1 enzyme. A loss-of-function experiment showed that 72 hours after genetic silencing of GCLC/GCLM resulted in remarkable apoptotic changes. The annexin V/PI apoptosis assay showed that apoptotic cell population increased remarkably upon siRNA treatment (R132H, siCont = 0.45% vs. siGCLM.2 = 31.2%; Fig. 2D and E). Moreover, Western blot analysis confirmed that cleaved caspase-3 was increased in U251 IDH1 R132H cells after genetic silencing of GCLC and GCLM (Supplementary Fig. S1C). ROS scavenger catalase restored caspase-3 cleavage in the presence of GCLC and GCLM RNA interference (Supplementary Fig. S1D). The increase of apoptosis was not observed when mutant IDH1 enzyme is absent, suggesting that the blockade of GSH metabolism is selectively toxic to IDH1-mutated cells. Furthermore, we recorded a great increase in cytoplasmic ROS levels after 48 hours of siRNA treatment, suggesting that apoptosis could be caused by ROS-derived cellular damage (Fig. 2F).

Figure 2.

GSH de novo synthesis support cellular physiology in IDH1-mutated cells. A, Western blot measures the expression of GSH synthesis enzymes in U251 cells with IDH1-mutant expression. β-Actin was used as internal control. B, qRT-PCR analysis measures mRNA level of GSH synthesis enzymes. *, P < 0.05; **, P < 0.01. C, GSH and GSSG level was measured in IDH1-mutated U251 cells. Catalase (Cata) was used as exogenous ROS scavenger (500 U/mL, 24 hours; **, P < 0.01). D, Annexin V/PI apoptotic analysis in IDH1-mutated U251 cells with genetic silencing of GCLC and GCLME, Quantification of apoptotic cells in DF, ROS-Glo assay in IDH1-mutated U251 cells with genetic silencing of GCLC and GCLM. Dox, doxycycline.

Activation of Nrf2 antioxidant pathway in IDH1-mutated cells

The transcription factor Nrf2 is a basic leucine zipper (bZIP) protein that regulates the cellular responses to oxidative stress by activating the expression of antioxidant genes (24). Under physiologic conditions, Nrf2 is tightly controlled by interaction with Kelch-like ECH-associated protein 1 (Keap1), which is a protein adaptor for E3 ubiquitin ligases, and proteasomal degradation. When cells are challenged by oxidative stress, Keap1–Nrf2 interaction is disrupted and the dissociated Nrf2 translocate into the nucleus for transcriptional activation (25). In IDH1-mutated cells, the increased ROS levels may trigger Nrf2 stabilization and gene transcription, which could be relevant to prompt GSH synthesis. To test this hypothesis, we first evaluated Nrf2-associated gene transcription via ARE-luciferase reporter assay. We found that mutant IDH1 expression was associated with enhanced Nrf2-dependent transcriptional activation (Fig. 3A). Furthermore, through ChIP assay, we demonstrated that the affinity of Nrf2 to antioxidant gene promoters was strongly enhanced after mutant IDH1 introduction, indicating that Nrf2 transactivation plays a central role in ROS homeostasis in IDH1-mutated cells (Fig. 3B). Consistent with these findings, genetic silencing of Nrf2 resulted in downregulation of antioxidant genes, such as GCLC, GCLM, HMOX1, NQO1, and SLC7A11 in IDH1-mutated cells (Fig. 3C; Supplementary Fig. S1E). The induction of Nrf2 transcription activity was accompanied by its prolonged protein stability. Immunoprecipitation assay showed that Nrf2 ubiquitination is compromised in the presence of mutant IDH1 (Fig. 3D). Furthermore, the protein stability of Nrf2 was elevated when mutant IDH1 enzymes were expressed (Fig. 3E and F). The protein half-lives of Nrf2 were prolonged from 22.4 to 49.2 minutes (R132C) or 62.3 minutes (R132H).

Figure 3.

Nrf2 regulates GSH metabolism in IDH1-mutated cells. A, ARE-Luciferase reporter assay was performed in U251 cells with IDH1-mutant expression. *, P < 0.05. B, ChIP PCR assay showed antioxidant genes promoter affinity of Nrf2 in IDH1-mutated U251 cells. **, P < 0.01. C, Real-time PCR assay showed mRNA level of antioxidant genes after 48 hours genetic silencing of Nrf2 in U251 cells with IDH1-mutant expression. **, P < 0.01. D, Immunoprecipitation (IP) assay measures Nrf2 ubiquitination in U251 cells with IDH1-mutant expression. E, Cycloheximide (CHX) pulse chase assay measures Nrf2 protein stability in IDH1-mutated U251 cells. F, Quantification of Nrf2 half-lives from results in E. Dox, doxycycline; IB, immunoblot; n.s., not significant.

Suppressing Nrf2/GSH axis results in oxidative damage in IDH1-mutated cells

Considering the central role of Nrf2 in the physiology of IDH1-mutated cells, blockade of Nrf2/GSH axis may be an effective therapeutic approach for tumors with IDH1 R132 variants. To investigate this, we tested a Nrf2 inhibitor, brusatol, in IDH1-mutated cells. Brusatol has been shown to strongly reduce Nrf2 transcriptional activity and enhance chemosensitivity in transformed cells (21, 26). Here, we confirmed that brusatol promoted Nrf2 degradation in IDH1-mutated cells, as evidenced by increased Nrf2 protein ubiquitination (Fig. 4A). Moreover, cycloheximide pulse chase assay confirmed that Nrf2 protein stability is compromised upon brusatol treatment (Fig. 4B). The protein half-lives decreased for both IDH1 R132C (67.1 vs. 11.1 minutes) and R132H (41.6 vs. 10.34 minutes) variants (Fig. 4C). Furthermore, Western blot analysis showed that Nrf2 protein levels drastically decreased after brusatol treatment (Fig. 4D). Accordingly, ChIP-PCR assay showed that the Nrf2 affinity for DNA sharply decreased in the presence of brusatol (Fig. 4E).

Figure 4.

Suppressing Nrf2/GSH axis results in oxidative damage in IDH1-mutated cells. A, Immunoprecipitation (IP) assay measures Nrf2 ubiquitination in U251 cells with IDH1-mutant expression after brusatol (Bru) treatment (40 nmol/L, 12 hours). B, Cycloheximide (CHX) pulse chase assay measures Nrf2 protein stability in IDH1-mutated U251 cells after brusatol treatment. C, Quantification of Nrf2 half-lives from results in BD, Western blot measures the expression of GSH synthesis enzymes with brusatol treatment (40 nmol/L, 24 hours) in IDH1-mutated U251 cells. E, ChIP PCR assay showed antioxidant genes promoter affinity of Nrf2 with brusatol (40 nmol/L, 24 hours) in IDH1-mutated U251 cells. *, P < 0.05; **, P < 0.01. F, Annexin V/PI apoptosis assay showed apoptotic changes in IDH1-mutated U251 cells with brusatol treatment (40 nmol/L, 72 hours). Exogenous antioxidant catalase (Cata) was used as exogenous ROS scavenger. G, Quantification of apoptotic cells in F. ***, P < 0.001. H, ARE-Luciferase reporter assay showed Nrf2-associated gene transcription with brusatol (40 nmol/L, 24 hours) in IDH1-mutated U251 cells. *, P < 0.05; **, P < 0.01. I, GSH/GSSG measurement in IDH1-mutated U251 cells with brusatol treatment (40 nmol/L, 24 hours). **, P < 0.01. J, ROS-Glo measurement in IDH1-mutated U251 cells with brusatol treatment (40 nmol/L, 24 hours; **, P < 0.01). IB, immunoblot.

Importantly, the suppression of Nrf2 activity resulted into exacerbated oxidative damage and cell death in IDH1-mutated cells. Annexin V/PI flow cytometry assay showed that brusatol increased apoptotic rates by 1.9-fold and 2.7-fold in IDH1 R132C and R132H U251 cells, respectively (Fig. 4F and G). Consistently, brusatol also resulted in cell apoptosis in IDH1-mutated BTIC TS603, but the trend was much less in IDH1 wild-type BTICs (Supplementary Fig. S2D and S2E). We noticed that brusatol treatment resulted in reduced ARE-luciferase activity, suggesting that Nrf2 activity is suppressed in these cells (Fig. 4H). Quantification of GSH revealed that brusatol further reduced GSH availability in IDH1-mutated cells. Brusatol decreased the GSH/GSSG ratio by 75.8% and 75.9% in IDH1 R132C and R132H cells, respectively (Fig. 4I). Accordingly, the cytoplasmic levels of ROS were significantly elevated by brusatol treatment (Fig. 4J

Targeting Nrf2/GSH axis suppresses IDH1-mutated xenografts

The aforementioned in vitro experiments strongly indicate that the blockade of GSH metabolism could be a valuable approach to suppress malignancies with mutant IDH1 enzymes. To better test this hypothesis, we established a xenograft mice model based on a patient-derived IDH1-mutated cell line TS603 (Fig. 5A). Patient-derived TS603 glioma cells with intrinsic mutant IDH1 enzyme were injected into NSG immunocompromised mice to establish xenograft tumor. When the tumor mass approaches 50 mm3, mice were treated with either brusatol and/or the exogenous antioxidant NAC. Tumor growth curve showed that brusatol significantly reduced the expansion of tumor mass (Fig. 5B and C). Notably, NAC abolished the suppressive effect of brusatol, suggesting ROS played a critical role in brusatol effects on tumor growth. No significant loss of body weight was observed during the treatment (Supplementary Fig. S1F). Histologic analysis revealed that brusatol treatment reduced the expression of antioxidant genes such as Nrf2, SLC7A11, GCLC, and GCLM (Fig. 5D). NAC slightly restored antioxidant gene expression in the xenografts. On the other hand, brusatol led to reduced expression of Ki67, but elevated levels of DNA damage markers, γH2A.X and TUNEL (Fig. 5E). Similarly, NAC treatment minimized cytotoxicity in IDH1-mutated xenografts.

Figure 5.

Targeting Nrf2/GSH axis suppresses IDH1-mutated xenografts. A, Schematic illustration for the xenograft and treatment schedule. B, Tumor growth curve of TS603 xenografts. n = 10 for each group. **, P < 0.01. C, Gross anatomy of TS603 xenografts. D, IHC assay showed the expression of Ki67, γH2AX, and terminal deoxynucleotidyl transferase–mediated dUTP nick end labeling (TUNEL) assay in tumor sections. Scale bar, 50 μm. E, IHC assay showed the expression of Nrf2, SLC7A11, GCLC, and GCLM in tumor sections. Scale bar, 50 μm. Bru, brusatol.


IDH1-mutated malignancies and therapeutic approaches

Mutations of IDH1/2 genes are widespread genetic abnormalities detected in several types of human malignancies, including lower grade glioma, leukemia, chondroma, chondrosarcoma, and cholangiocarcinoma. Cancer-associated IDH mutations cause amino acid substitution of an arginine residue in the IDH enzyme catalytic center. Biochemical studies showed that IDH1 R132 mutants have elevated affinity for both NADPH (Km = 0.44 μmol/L) and α-ketoglutarate (Km = 965 μmol/L), indicating that the mutant enzyme prefers NADPH and α-ketoglutarate, whereas wild-type IDH enzyme prefers NADP+ and isocitrate for its catalytic function (7, 27).

Several pioneered studies showed that direct targeting mutant IDH1 enzyme is an effective treatment for IDH1-mutated hematopoietic malignancies, such as relapsed or refractory AML (28). Regarding IDH1-mutated solid tumors, Rohle and colleagues (13) showed that the inhibition of mutant IDH1 delayed IDH1-mutated xenograft expansion in vivo. However, preliminary data from several early phase clinical trials showed modest impact on objective response rate and delay of progression of IDH1-mutated solid tumors. More effective therapies, such as developing refined inhibitors of mutant IDH1, or targeting IDH-related pathways, have been urged to improve disease outcome of IDH1-mutated malignancies. Besides direct targeting the mutant enzyme, several lines of evidence show that metabolic reprogramming in IDH1-mutated cells could be targeted to synergize with conventional chemo/radiotherapies. We and other colleagues demonstrated that NAD+ depletion, as well as 2-HG–mediated deficiency in homologous DNA recombination establish vulnerability to PARP inhibitors in IDH1-mutated cells (29–32). The glutaminase inhibitor, CB-839, has also been proposed to be useful for the treatment of IDH1-mutated cancers (33, 34). In this study, we extended the investigation of effective therapy for IDH1-mutated cancer and discovered that Nrf2/GSH metabolism could be another therapeutic vulnerability in malignancies that harbor IDH1 mutation.

IDH-mutated cells develop dependency on GSH ROS scavenging

The production of ROS is involved in several aspects of cancer biology, such as genomic instability, loss of growth control, cellular motility, and tumor invasiveness (35, 36). On the other hand, excessive ROS is harmful to biological molecules, resulting in oxidative damage to DNA, lipid, and proteins (37). Maintaining appropriate ROS levels is key to cancer cells during oncogenesis and therapeutic resistance. GSH is an endogenous antioxidant tripeptide that participates in the elimination of reactive molecules, such as free radicals, peroxides, lipid peroxides, and metals. The thiol group in reduced GSH is responsible for its reducing activity, which alleviates oxidative stress through direct reduction of disulfide bonds in the cysteine residues of cytoplasmic proteins and eliminates ROS through the GSH-ascorbate cycle (38). For IDH1-mutated malignancies, several pioneered studies suggested the correlation with GSH depletion and ROS accumulation (39). Although failed control of intracellular ROS has been indicated with tumorigenesis process (40), there is still lack of a direct evidence showing IDH1-mutant–derived ROS promote tumor development.

In this study, we found that IDH1-mutated malignancies exhibit a tendency to suffer oxidative stress, as the introduction of mutant IDH1 is closely associated with elevated ROS levels in cytoplasm and mitochondria (Fig. 1C–H). Similarly, recent research suggests that tumoral GSH levels negatively correlate with 2-HG, suggesting that IDH-mutated cells have elevated demands for GSH (41). Our findings showed that, in IDH1-mutated cells, the key regulatory enzymes of GSH biosynthesis were upregulated to meet the increased demands of endogenous antioxidant systems (Fig. 2A–C). Loss-of-function experiments demonstrated that blocking GSH synthesis led to remarkably elevated ROS levels, oxidative stress, and apoptotic changes (Fig. 2D and E). Overall, our findings suggest that upregulation of GSH-based ROS scavenging pathways play a central role to maintain cellular homeostasis in IDH1-mutated cancers.

Nrf2-regulated GSH metabolism

The multifunctional transcriptional factor, Nrf2, governs the cellular response to oxidative stress by triggering antioxidant gene transcription. It regulates a variety of genes for electrophile and oxidant metabolism, as well as genes that support cellular survival under stress conditions (42). In this study, we recorded enhanced Nrf2 transcriptional activity that was associated with the presence of mutant IDH1 enzyme (Fig. 3A and B), suggesting that Nrf2-driven antioxidant response is a compensatory response to IDH1-associated oxidative stress. As a further validation, it was shown that several known Nrf2 transcription targets, such as GCLC, GCLM, HMOX1, NQO1, and SLC7A11, were upregulated in IDH1-mutated cells (Fig. 3C). Importantly, these genes play central role in cysteine uptake and de novo GSH biosynthesis, indicating that the protective effect of Nrf2 is a result of increased intracellular GSH pool. Protein stability tests showed that Nrf2 was less ubiquitinated and degraded in IDH1-mutated cells, which would lead to the activation of antioxidant expression (Fig. 3D–F). Moreover, the activation of Nrf2/antioxidant pathway not only relieves the metabolic stress for IDH1-mutated cells but may also support cellular viability and promote growth advantage during oncogenesis. Our findings highlighted the role of Nrf2-dependent GSH metabolism in IDH1-mutated cells, indicating a selective vulnerability of IDH1-mutated malignancies.

Targeting Nrf2/GSH metabolism as a new strategy for IDH1-mutated malignancies

Considering the critical role of Nrf2/GSH metabolism in cancer biology and therapeutic resistance, several attempts have been made to achieve specific targeting of this pathway using small-molecular compounds. For example, by high-throughput screening assay, Singh and colleagues (43) reported that the small-molecule compound ML-385 exhibits inhibitory effect on Nrf2 transcriptional activity. However, the effective dosage of ML-385 is too high for further preclinical studies in animals. In another example, Ren and colleagues (21) reported that brusatol, a quassinoid compound, exhibits potent inhibitory effect on Nrf2 transcriptional activity. In our study, we confirmed that brusatol is able to block Nrf2 activity in IDH1-mutated cells, which strongly suppressed antioxidant pathways, such as de novo GSH biosynthesis (Fig. 4). Interestingly, brusatol has been used as a sensitizer for conventional chemotherapy (21, 26, 44). In contrast, our study showed that at a similar dosage, brusatol monotherapy is sufficient to cause apoptotic changes in IDH1-mutated cells (45). We speculate that the brusatol potent efficacy is due to the induction of ROS levels, which leads to cell death in concert with the inhibition of endogenous antioxidants expression. The xenograft experiments confirmed this hypothesis. The introduction of an exogenous ROS scavenger, NAC, compromised the tumor-suppressing effect of brusatol, suggesting that ROS play a key role in brusatol-induced cytotoxicity (Fig. 5).

Overall, our study showed that neomorphic activity of mutant IDH1 enzyme results in mitochondrial and cytoplasmic ROS accumulation by disrupting NADP+/NADPH balance. The major regulator of antioxidant responses, Nrf2, controls the GSH de novo synthesis and plays a central role in the cellular physiology of IDH1-mutated cells. Blockade of Nrf2/antioxidant pathway exhibited selective cytotoxicity in cells with IDH1 mutation (Fig. 6). Our findings highlight the importance of GSH metabolism in IDH1-mutated cells and indicate a novel therapeutic approach for malignancies with IDH1 mutation.

Figure 6.

Targeting Nrf2/GSH axis for IDH1-mutated cancers. Cancer-associated IDH1 mutants cause metabolic deficiency and accumulation of oxidative stress. Nrf2 plays a key protective role in IDH1-mutated cells by transcribing GSH synthesis enzymes. The enhanced de novo GSH synthesis neutralizes excessive ROS, and therefore avoids oxidative damages to macromolecules. Targeting Nrf2/GSH could be a novel therapeutic strategy in these types of human malignancies.

Accumulation of 2-hydroxyglutarate is not a biomarker for malignant progression in IDH-mutated low-grade gliomas

Neuro Oncol. 2013 Jun; 15(6): 682–690.
Online 2013 Feb 14. doi: 10.1093/neuonc/not006


To determine whether accumulation of 2-hydroxyglutarate in IDH-mutated low-grade gliomas (LGG; WHO grade II) correlates with their malignant transformation and to evaluate changes in metabolite levels during malignant progression.


Samples from 54 patients were screened for IDH mutations: 17 patients with LGG without malignant transformation, 18 patients with both LGG and their consecutive secondary glioblastomas (sGBM; n = 36), 2 additional patients with sGBM, 10 patients with primary glioblastomas (pGBM), and 7 patients without gliomas. The cellular tricarboxylic acid cycle metabolites, citrate, isocitrate, 2-hydroxyglutarate, α-ketoglutarate, fumarate, and succinate were profiled by liquid chromatography–tandem mass spectrometry. Ratios of 2-hydroxyglutarate/isocitrate were used to evaluate differences in 2-hydroxyglutarate accumulation in tumors from LGG and sGBM groups, compared with pGBM and nonglioma groups.


IDH1 mutations were detected in 27 (77.1%) of 37 patients with LGG. In addition, in patients with LGG with malignant progression (n = 18), 17 patients were IDH1 mutated with a stable mutation status during their malignant progression. None of the patients with pGBM or nonglioma tumors had an IDH mutation. Increased 2-hydroxyglutarate/isocitrate ratios were seen in patients with IDH1-mutated LGG and sGBM, in comparison with those with IDH1-nonmutated LGG, pGBM, and nonglioma groups. However, no differences in intratumoral 2-hydroxyglutarate/isocitrate ratios were found between patients with LGG with and without malignant transformation. Furthermore, in patients with paired samples of LGG and their consecutive sGBM, the 2-hydroxyglutarate/isocitrate ratios did not differ between both tumor stages.


Although intratumoral 2-hydroxyglutarate accumulation provides a marker for the presence of IDHmutations, the metabolite is not a useful biomarker for identifying malignant transformation or evaluating malignant progression.

Keywords: α-ketoglutarate, IDH1 mutations, liquid chromatography–tandem mass spectrometry, low-grade gliomas, secondary glioblastomas, 2-hydroxyglutarate

Low-grade gliomas (LGG) occur in the cerebral hemispheres and represent 10%–15% of all astrocytic brain tumors.1 Despite long-term survival in many patients, 50%–75% of patients with LGG eventually die of either progression of a low-grade tumor or transformation to a malignant glioma.2 The time to progression can vary from a few months to several years,35 and the median survival among patients with LGG ranges from 5 to 10 years.6,7 Among several risk factors, only age, histology, tumor location, and Karnofsky performance index have generally been accepted as prognostic factors for patients with LGG.8,9 As a prognostic molecular marker, only 1p19q codeletion was identified as such in pure oligodendrogliomas. However, this association was not seen in either astrocytomas or oligoastrocytomas.10

Somatic mutations in human cytosolic isocitrate dehydrogenases 1 (IDH1) were first described in 2008 in ∼12% of glioblastomas11 and later in acute myeloid leukemia, in which the reported mutations were missense and specific for a single R132 residue.11,12 Some gliomas lacking cytosolic IDH1 mutations were later observed to have mutations in IDH2, the mitochondrial homolog of IDH1.12IDH mutations are the most commonly mutated genes in many types of gliomas, with incidences of up to 75% in grade II and grade III gliomas.13,14 Further frequent mutations in patients with LGG were recently identified, including inactivating alterations in alpha thalassemia/mental retardation syndrome X-linked (ATRX), inactivating mutations in 2 suppressor genes, homolog of Drosophila capicua (CIC) and far-upstream binding protein 1 (FUBP1), in about 70% of grade II gliomas and 57% of sGBM.1517 The association between ATRX mutations with IDHmutations and the association between CIC/FUBP1 mutations and IDH mutations and 1p/19q loss are especially common among the grade II-III gliomas and remarkably homogeneous in terms of genetic alterations and clinical characteristics.16

It was thought that IDH mutations might be a prognostic factor in LGG, predicting a prolonged survival from the beginning of the disease.1823 However, this assumption, as shown in our and other earlier studies, had to be corrected because survival among patients who have LGG with IDH mutations is only improved after transformation to secondary high-grade gliomas.18,19,24 Furthermore, it had already been demonstrated that an IDH mutation is not a biomarker for further malignant transformation in LGG.18 IDH1 and IDH2 catalyze the oxidative decarboxylation of isocitrate to α-ketoglutarate (α-KG) and reduce NADP to NADPH.25 The mutations inactivate the standard enzymatic activity of IDH112 and confer novel activity on IDH1 for conversion of α-KG and NADPH to 2-hydroxyglutarate (2HG) and NADP+, supporting the evidence thatIDH1 and 2 are proto-oncogenes. This gain of function causes an accumulation of 2HG in glioma and acute myeloid leukemia samples.26,27 The 2HG levels in cancers with IDH mutations are found to be consistently elevated by 10–100-fold, compared with levels in samples lacking mutations of IDH1 or IDH2.26,28Nevertheless, how exactly the production or accumulation of 2HG by mutant IDH might drive cancer development is not well understood.

In the present study, we postulate that intratumoral 2HG could be a useful biomarker that predicts the malignant transformation of WHO grade II LGG. We therefore screened for IDH mutations in patients with LGG and measured the accumulation of 2HG in 2 populations of patients, patients with LGG with and without malignant transformation, with use of liquid chromatography–tandem mass spectrometry (LC-MS/MS). Furthermore, we compared the concentrations of 2HG in LGG and their consecutive secondary glioblastomas (sGBM) to evaluate changes in metabolite levels during the malignant progression.

Go to:Methods and Materials


According to aforementioned criteria, a total of 72 tumor samples from 54 patients were analyzed (Table 1). The samples were from 17 patients with LGG without malignant transformation, 18 patients with both LGG and consecutive sGBM (n = 36), 2 additional patients with sGBM, 10 patients with pGBM, and 7 patients with nonglioma tumors. The nonglioma samples comprised 3 meningiomas, 2 metastases of breast cancers, 1 cavernoma, and 1 reactive gliosis from a patient with epilepsy.

Table 1.

Patient characteristics

DNA Isolation and IDH Mutation Detection

Tumor tissue samples were taken intraoperatively and were snap frozen at −80°C. To ensure a tumor cell content of at least 80% for nucleic acid extraction, control slides stained with hematoxylin and eosin were examined by the local neuropathologist. IDH1 and IDH2 mutations were assessed using direct DNA sequencing, as reported previously.18

Progression and Survival

Progression-free survival (PFS) was defined as the time from first diagnosis of an LGG to tumor progression or end of follow-up. Time to malignant transformation was defined as the time from the day of first surgery for an LGG to the day of surgery for malignant progression to a secondary high-grade glioma. Overall survival (OS) was the time from the day of first surgery to death or end of follow-up. All patient data were updated on June 15, 2012.

LC-MS/MS Analysis of Tricarboxylic Acid Cycle (TCA) Metabolites

Instrumentation included an AB Sciex QTRAP 5500 triple quadruple mass spectrometer coupled to a high-performance liquid chromatography (HPLC) system from Shimadzu containing a binary pump system, an autosampler, and a column oven. Targeted analyses of citrate, isocitrate, α-ketoglutarate (α-KG), succinate, fumarate (Sigma-Aldrich), and 2-hydroxyglutarate (2HG; SiChem GmbH) were performed in multiple reaction monitoring (MRM) scan mode with use of negative electrospray ionization (-ESI). Expected mass/charge ratios (m/z), assumed as [M-H+], were m/z 190.9, m/z 191.0, m/z 145.0, m/z 116.9, m/z 114.8, and m/z 147.0 for citrate, isocitrate, α-KG, succinate, fumarate, and 2HG, respectively. For quantification, ratios of analytes and respective stable isotope-labeled internal standards (IS) (Table 2) were used. For quantification of isocitrate and 2HG, stable isotope-labeled succinate was used as IS because of unavailability of labeled analogs. MRM transitions are summarized in Table 2.

Table 2.

MRM transitions and respective fragmentation parameters


Patient Characteristics

According to the malignant progression of the LGG, patients were divided into two groups: group 1 patients with LGG (LGG1) without malignant transformation and group 2 patients with LGG (LGG2) with histologically confirmed malignant progression.

Among 35 patients with LGG who were included in this study, 19 (54%) were women, and 16 (46%) were men. Furthermore, histologically, 6 patients had oligoastrocytomas, and the remaining 29 patients had diffuse astrocytomas.

The median age of all patients with LGG was 37.4 years at the time of the first diagnosis. Patients in LGG2 had a median age of 37.1 years, which did not differ significantly from that of patients in LGG1, who had a median age of 41.4 years.

The median time to malignant transformation among patients in the LGG2 group was 3.35 years (range, 2.5–5.4 years). The median OS among all patients with LGG was 13.1 years (11.4 years in LGG1 and 13.1 years in LGG2; P = .97).

IDH1 Mutation and Outcome

An IDH1 mutation was detected in 27 of 35 patients with LGG (77.1%), in 10 of 17 patients in LGG1 (59%), and in 17 of 18 patients in LGG2 (95%). In all cases, IDH1 mutations were found on R132. IDH2mutations were not detected in any of the patients. The IDH1 mutation status was stable during progression from LGG to sGBM in all patients in LGG2. None of the patients with pGBM or nonglioma had an IDHmutation. Patients with LGG with an IDH1 mutation had a median PFS of 3.3 years, which was comparable to that among patients with wild-type LGG (2.8 years; P > .05). Furthermore, the OS among patients with LGG with an IDH1 mutation was not statistically different at 13.0 years compared with that among patients with LGG without an IDH1 mutation, who had an OS of 9.3 years (P = .66).

LC-MS/MS Profiling of TCA Metabolites

TCA metabolites, citrate, isocitrate, α-ketoglutarate, succinate, fumarate, and 2-hydroxyglutarate were measured in glioma samples with and without an IDH1 mutation, in samples identified as primary GBM, and in nonglioma brain tumor specimens (Fig. 1). No differences in citrate, isocitrate, α-KG, succinate, and fumarate concentrations were found when comparing all of the latter groups. Concentrations of 2HG, a side product in IDH1-mutated gliomas, were 20–34-fold higher in IDH1-mutated gliomas (0.64–0.81 µmol/g), compared with non–IDH1-mutated LGG1 (P ≤ .001). No differences were observed between IDH1-mutated gliomas and IDH1-nonmutated LGG2 and sGBM, caused by strongly elevated 2HG levels in either 1 or 2 samples in these groups, respectively. Furthermore, in IDH1-mutated gliomas, 2HG concentrations were a mean of 20 times higher than in pGBM and nongliomas (P ≤ .001) (Fig. 1). No differences were observed between the single groups of IDH1-mutated gliomas LGG1, LGG2, and sGBM in relation to 2HG concentration.

Fig. 1.

Dot-box and whisker plots show concentration ranges for TCA metabolites measured in IDH1-nonmutated (IDH1wt) and IDH1-mutated (IDH1mut) LGG and sGBM and in pGBM and nonglioma tumor specimens; boxes span the 25th and 75th percentiles with median, whiskers

To detect possible differences among the IDH1-mutated LGG1, LGG2, and sGBM, the α-KG/isocitrate and 2HG/isocitrate ratios were used in additional tests. Therefore, the direct precursor-product relation would correct for all differences possibly expected during pre-analytical processing. To prove this, analyte ratios ofIDH1-mutated and nonmutated gliomas were compared. IDH1-mutated gliomas showed a 2HG/isocitrate ratio that was 13 times higher (P ≤ .001) (Fig. 2A), which corresponds to a lower accumulation of 2HG inIDH1-nonmutated gliomas. α-KG/isocitrate ratios were determined to be approximately 10-fold higher inIDH1-mutated gliomas than in IDH1-nonmutated gliomas (P = .005) (Fig. 2B), which also implies lower accumulation of α-KG in IDH1-nonmutated gliomas.

Fig. 2.

2-Hydroxyglutarate to isocitrate ratios (A) and α-ketoglutarate to isocitrate ratios (B) for IDH1-nonmutated (IDH1wt) and IDH1-mutated (IDH1mut) gliomas (LGG and sGBM); boxes span the 25th and 75th percentiles with median, and whiskers represent

2HG/isocitrate and α-KG/isocitrate ratios, respectively, were calculated in all 8 specimen groups (Fig. 3). In addition to the differences in 2HG/isocitrate ratios of IDH1-mutated and nonmutated gliomas (Fig. 2A), the ratios in IDH1-mutated gliomas were 4–9 times higher, compared with those in pGBM (P ≤ .001), and 3–6 times higher, compared with those in non-glioma tumor specimens, which was not statistically significant (Fig. 3A). In detail, ratios of 2HG and isocitrate were established to be 13, 9.4, and 22 times higher in IDH1-mutated LGG1, LGG2, and their consecutive sGBM, respectively, than in IDH1-nonmutated LGG1 (Fig. 3A). No significant differences were observed between IDH1-mutated gliomas and IDH1-nonmutated LGG2 and sGBM. The comparison of 2HG/isocitrate ratios between IDH1-nonmutated gliomas and IDH1-mutated LGG2 and sGBM showed no statistically significant differences. However, a trend toward higher ratios inIDH1-mutated LGG1/2 was seen. Furthermore, no differences could be determined by comparing 2HG/isocitrate ratios measured in the groups of IDH1-mutated LGG1 and LGG2. Although 2HG/isocitrate ratios in IDH1-mutated secondary glioblastomas are 1.7 and 2.3 times higher than in the LGG1 and LGG2 groups, respectively, no statistically significant differences were observed.

Fig. 3.

2-Hydroxyglutarate to isocitrate (A, C) and α-ketoglutarate to isocitrate (B, D) ratios for groups of IDH1-nonmutated (IDH1wt) and mutated gliomas (IDH1mut), pGBM, and nongliomas; dot-box and whisker plots (A, B) span the 25th and 75th percentiles

The absence of a straight trend to higher 2HG/isocitrate ratios during malignant progression is shown by paired analysis of IDH1-mutated LGG2 and their consecutive sGBM (Fig. 3C). Similar findings were observed using the α-KG/isocitrate ratios. Although significant differences were found, with ratios approximately 10 times higher in IDH1-mutated glioblastomas than in IDH1-nonmutated glioblastomas (Fig. 2B), it was not possible to differentiate among the 3 IDH1-mutated glioblastoma groups LGG1, LGG2, and their consecutive sGBM with use of this analyte ratio (Fig. 3B and D).

Go to:Discussion     

On the basis of a comprehensive analysis of cellular TCA metabolites from several cohorts of patients with glioma and nonglioma, our study provides evidence that the level of 2HG accumulation is not suitable as an early biomarker for distinguishing patients with LGG in relation to their course of malignancy. To our knowledge, this is the first report of a paired analysis of 2HG levels in LGG and their consecutive sGBM showing stable 2HG accumulation during malignant progression. This fact assumes that malignant transformation of IDH-mutated LGG appears to be independent of their intracellular 2HG accumulation. Considering these results, we could not stratify patients with LGG into subgroups with distinct survival.

To date, little is known about biomarkers that may predict malignant transformation and, consequently, predict survival in patients with LGG. The investigation of biomarkers in this patient group is relevant because treatment interventions can be tailored to prolong survival, minimize treatment-related adverse effects, and, accordingly, maximize quality of life. In many previous studies, IDH mutations, as the most commonly detected mutations in LGG, were observed to be a significant prognostic factor in patients with glioma, often relating to improved survival among patients with LGG.1823 However, only patients withIDH-mutated LGG with malignant progression have a prolonged OS.18,19,24 In an earlier conducted study, the analysis of 2 groups of patients with LGG with and without malignant transformation failed to provide a significant influence from the IDH mutation, neither on the PFS nor on the OS.18 In agreement with these data, in a recent study, we showed again that PFS and OS among patients with LGG with and without anIDH mutation did not differ significantly, despite the malignant transformation in LGG2. This result is not unexpected because the same patient population was analyzed in both studies.

Accumulation of 2HG in IDH1-mutated gliomas was first described by Dang et al.26 2HG accumulation is an important marker of IDH1/2-mutated gliomas and other neoplasms.26,29,30 Furthermore, it was assumed that 2HG accumulation might be a potential systemic biomarker of gliomas, but it was not detectable in serum samples from patients with glioma.31 Nevertheless, it has become possible in the meantime to detect 2HG production using magnetic resonance spectroscopy in a noninvasive manner to identify patients withIDH1 mutant brain tumors.32

Because 2HG accumulation provides one of the few potential read-outs for mutant IDH enzymatic activity, we suspected that different intratumoral levels of 2HG accumulation in patients with LGG (especially in those with an IDH mutation) may affect their clinical course in relation to malignant transformation. Therefore, intratumoral concentrations of 2HG and other TCA metabolites were quantified by LC-MS/MS, showing concentrations in IDH1-mutated glioma samples that were comparable to the levels described by Dang et al.26 As previously shown,22 no differences in metabolite levels were observed, with the exception of 2HG with increased accumulation in IDH-mutated gliomas, compared with IDH-nonmutated specimens. However, the fold-difference in 2HG levels between IDH1 mutant and IDH1 wild-type LGG in our study was smaller than in some other studies.26,30 As a reason for this issue, we identified 3 samples of IDH1 wild-type LGG and sGBM with strongly elevated 2HG levels. The fact that some IDH1 wild-type tumors might accumulate 2HG was previously described by Wise et al,33 who reported that the increased IDH2-dependent carboxylation of glutamine-derived α-KG in hypoxia is associated with a concomitant increased synthesis of 2HG in cells with wild-type IDH1 and IDH2. Thus, they concluded that, in further support of the increased mitochondrial reductive glutamine metabolism that they observed in hypoxia, the incubation in hypoxia can lead to elevated 2HG levels in cells lacking IDH1/2 mutations.

The ratio of 2HG to isocitrate and the ratio of α-KG to isocitrate were provided. The use of these ratios can function as an internal control because isocitrate is the direct precursor of α-KG and 2HG. However, intracellular accumulation of both metabolites (2HG and of α-KG) did not differ between both IDH-mutated LGG groups with and without malignant progression. In the same way, detection of 2HG concentrations and the 2HG/isocitrate ratios in patients with LGG and their consecutive sGBM were comparable to values during malignant progression. Correspondingly, all other assessed TCA metabolites remained stable during the malignant progression of the LGG to sGBM. Therefore, 2HG accumulation appears, at least for now, merely to represent a highly correlative and stable marker for an emerging class of somatic mutations in theIDH enzymes from the early stage of glioma development and after malignant transformation to high-grade gliomas.

An expected lower level of α-KG in IDH-mutated LGG and sGBM, in comparison with IDH-nonmutated LGG, pGBM, and nonglioma tumors, was not detected in our study. This is in concordance with a similar finding by Dang et al,26 who showed unaffected α-KG levels in whole-tumor cell lysates. However, the latter finding was in contrast to reported results by Zhao et al,34 who showed that forced expression of mutant IDHin cultured cells led to a dose-dependent decrease in α-ketoglutarate levels. A possible explanation for this finding is the mono-allelic heterozygous mutations of the IDH1 gene leading to expression of both wild-type and mutant IDH1 in a single cell and, therefore, to production of 2HG and α-KG at the same time in every mutated cell.11,12 Another possible reason is the reported evidence that the biochemical effects of mutantIDH1 on α-KG-dependent enzymes are not principally attributable to depletion of α-KG but are a competitive antagonism with α-KG.3537 Thus, Xu et al postulated that IDH1 mutations alone do not reduce cellular level of α-KG sufficiently to have a significant tumorigenic consequence, but nonetheless these mutations sensitize α-KG–dependent dioxygenases to the inhibitory effect by the large amounts of intratumoral accumulated 2HG.35

Whether 2HG acts as a mutagen or plays a distinct role in gliomagenesis remains to be determined. Dang et al predicted that patients with LGG may benefit from the therapeutic inhibition of 2HG production, resulting in the slowing or halting of conversion of LGG into a lethal secondary glioblastoma, thus changing the course of the disease.26 However, our data confirm similar values of 2HG accumulation in the different LGG groups (with and without malignant progression) and present comparable ranges of 2HG in the low-grade and high-grade tumor stages. In addition, our study used the ratio of 2HG to isocitrate, which might provide a more sensitive screening tool for IDH-mutated LGG than an increase in the absolute concentration of 2HG.

Finally, more work is needed to provide valuable clues about the precise role that 2HG might play in the initiation and progression of LGG. Moreover, the value of 2HG as a useful biomarker for diagnosis or monitoring of the treatment response of LGG has not yet been realized.

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Author, reporter: Tilda Barliya PhD

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Word Cloud By Danielle Smolyar

Breast cancer is the second most common cancer worldwide after lung cancer, the fifth most  common cause of cancer death, and the leading  cause of cancer death in women. the global burden of  breast cancer exceeds all other cancers and the incidence  rates of breast cancer are increasing (1,2).

The heterogeneity of breast cancers makes them both a fascinating and challenging solid tumor to diagnose and treat. Here is a great review of the molecular pathology of breast cancer progression (3).

The molecular pathology of breast cancer progression” by Alessandro Bombonati  and Dennis C Sgroi.

Breast cancer is the most frequent carcinoma in females and the second most common cause of cancer related mortality in women. Approximately 54 000 and 207 000 new cases of in situ and invasive breast carcinoma, respectively. Overall, breast cancer incidence rates have levelled off since 1990, with a decrease of 3.5%/year from 2001 to 2004.  Most notably, during this same time period, breast cancer mortality rates have declined 24%, with the largest impact among young women and women with estrogen receptor (ER)-positive disease.

The decline in breast cancer mortality has been attributed to the combination of early detection with screening programmes and the advent of more efficacious adjuvant progression have aided in the discovery of novel pathway-specific targeted therapeutics, and the emergence of such effective therapeutics is currently driving the need for molecular-based, ‘patient-tailored’ treatment planning.

Proposed models of human breast cancer progression

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Epidemiological and morp

hological observations led to the formulation of several linear models of breast cancer initiation, transformation and

progression. Figure 1

The ductal and lobular subtypes constitute the majority of all breast cancers worldwide, with the ductal subtype accounting for 40–75% of all diagnosed cases.

The classic model of breast cancer progression of the ductal type proposes thatneoplastic evolution initiates in normal epithelium (normal), progresses to flat epithelial atypia (FEA), advances to atypical ductalhyperplasia (ADH), evolves to ductal carcinoma in situ (DCIS) and culminates as invasive ductal carcinoma (IDC).

The model of lobular neoplasia proposes a multi-step progression from normal epithelium to atypicallobular hyperplasia, lobular carcinoma in situ (LCIS) and invasive lobular carcinoma (ILC).

The cell of origin of breast cancer: the clonal and stem cell hypotheses

The two leading models accounting for breast carcinogenesis are the sporadic clonal evolution model and the cancer stem cell (cSC) model. According to the sporadic clonal evolution hypothesis, any breast epithelial cell can be the target of random mutations. The cells with advantageous genetic and epigenetic alterations are selected over time to contribute to tumour progression. The third alternative cSC model postulates that only stem and progenitor cells (representing a small fraction of the tumor cells within the cancer) can initiate and maintain tumor progression. Figure 2.

Normal breast stem cells (nBSCs) are long-lived, tissue-resident cells capable of self-renewal activity and multi-lineage differentiation that can recapitulate the breast tubulolobular architecture that is composed of luminal and myoepithelial cells.

As normal breast cancer stem cells are long-time tissue residents, it has been proposed that such cells are candidates for accumulating genetic and epigenetic modifications. It has been further proposed that such molecular alterations result in deregulation of normal self-renewal, leading to the development of a cancer stem cell (cSC).

It is believed that the cSC undergoes asymmetrical division, maintaining the stem cell population while at the same time differentiating into committed progenitor(s) cells that give rise to the different breast cancer subtypes.

A second scenario, as it relates to breast cancer development, is one in which the cancer-initiating cells are derived from committed progenitor cells that spawn different breast cancer subtypes. Both scenarios are highly supported.

Molecular analysis of the different stages of breast cancer progression

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Genomic and transcriptomic data in combination with morphological and immunohistochemical data stratify the majority of breast cancers into a “low-grade-like” molecular pathway and a “high-grade-like” molecular pathway. Figure 3. The low-grade-like pathway (left hand side) is characterized by recurrent chromosomal loss of 16q, gains of 1q, a low-grade-like gene expression signature, and the expression of estrogen and progesterone receptors (ER+ and PR+). The progression (vertical arrows) along this pathway (green rectangles) culminates with the formation of low and intermediate grade invasive ductal, (LG IDC and IG IDC) and invasive lobular carcinomas including both the classic (ILC) and the pleomorphic variant (pILC). The tumors arising from the low grade pathway are classified as luminal consisting of a continuum of gene expression frequently associated with the absence (luminal A) or presence of HER2 expression (luminal B). The vast majority of ILCs and pILCs and their precursors cluster together within the luminal subtype. The high grade-like gene expression molecular pathway (right hand side) is characterized by recurrent gain of 11q13 (+11q13), loss of 13q (13q−), expression of a high-grade-like gene expression signature, amplification of 17q12 (17q12AMP), and lack of estrogen and progesterone receptors expression (ER− and PR−). The progression along this pathway (red rectangles) includes intermediate and high grade ductal carcinomas that are stratified as HER2, or basal-like, depending on the expression/amplification of HER2. The molecular apocrine subtype, characterized by the lack of ER expression and presence of AR expression, arises from the high grade pathway. The model also depicts intra-pathway tumor grade progression (horizontal arrows).

Although the genomic and transcriptomic data presented in this review support the divergent model of breast cancer progression, the clinical experience indicates that tumors within each pathway are still fairly heterogeneous with respect to clinical outcome suggesting that even this advanced molecular progression scheme is oversimplified.

The future application of massively parallel sequencing technologies to the preinvasive stages of breast cancer will assist in assessing intratumoral heterogeneity during the transition from preinvasive to invasive breast cancer, and may assist in identifying early tumor initiating genetic events.


Over the past decade the integration of numerous genomic and transcriptomic analyses of the various stages of breast cancer has generated multiple novel insights in the complex process of breast cancer progression.

  • First, human breast cancer appears to progress along two distinct molecular genetic pathways that strongly associate with tumor grade.
  • Second, in the epithelial and non-epithelial components of the tumor microenvironment, the greatest molecular alterations (at the gene expression level) occur prior to local invasion.
  • Third, in the epithelial compartment, no major additional gene expression changes occur between the preinvasive and invasive stages of breast cancer.
  • Fourth, the non-epithelial compartment of the tumor micromilieu undergoes dramatic epigenetic and gene expression alterations occur during the transition form preinvasive to invasive disease. Despite these significant advances, we have only begun to scratch the surface of this multifaceted biological process. With the advent of additional novel high-throughput genetic, epigenetic and proteomic technologies, it is anticipated that the next decade of breast cancer research will gain an equally paralleled appreciation for the complexity breast cancer progression. It is with great hope that knowledge gained from such studies will provide for more effective strategies to not only treat, but also prevent breast cancer.


1. http://www.nature.com/nrclinonc/journal/v7/n12/pdf/nrclinonc.2010.192.pdf

2. Jemal, a. et al. CA Cancer J. Clin. 60, 277–300; 2010

3. Alessandro Bombonati and Dennis C Sgro. The molecular pathology of breast cancer progression. J Pathol 2011; 223: 307–317.



4. Rodney C. Richie and John O. Swanson. Breast Cancer: A Review of the Literature. J Insur Med 2003;35:85–101.

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