Altered cellular metabolism, including an increased dependence on aerobic glycolysis, is a hallmark of cancer. Despite the fact that this observation was first made nearly a century ago, effective therapeutic targeting of glycolysis in cancer has remained elusive. One potentially promising approach involves targeting the glycolytic enzyme lactate dehydrogenase (LDH), which is overexpressed and plays a critical role in several cancers. Here, we used a novel class of LDH inhibitors to demonstrate, for the first time, that Ewing sarcoma cells are exquisitely sensitive to inhibition of LDH. EWS-FLI1, the oncogenic driver of Ewing sarcoma, regulated LDH A (LDHA) expression. Genetic depletion of LDHA inhibited proliferation of Ewing sarcoma cells and induced apoptosis, phenocopying pharmacologic inhibition of LDH. LDH inhibitors affected Ewing sarcoma cell viability both in vitro and in vivo by reducing glycolysis. Intravenous administration of LDH inhibitors resulted in the greatest intratumoral drug accumulation, inducing tumor cell death and reducing tumor growth. The major dose-limiting toxicity observed was hemolysis, indicating that a narrow therapeutic window exists for these compounds. Taken together, these data suggest that targeting glycolysis through inhibition of LDH should be further investigated as a potential therapeutic approach for cancers such as Ewing sarcoma that exhibit oncogene-dependent expression of LDH and increased glycolysis.
LDHA is a pharmacologically tractable EWS-FLI1 transcriptional target that regulates the glycolytic dependence of Ewing sarcoma.
Glycolytic dependence of cancer cells was first described in 1927 by Otto Warburg, who hypothesized that tumor cells preferentially catabolized glucose to lactate even in the presence of oxygen (aerobic glycolysis), whereas normal cells preferentially catabolized glucose to carbon dioxide (oxidative phosphorylation; ref. 1). Targeting this increased dependence on glycolysis in cancer cells presents an opportunity to inhibit their growth while potentially limiting the toxicity delivered to normal cells (2). Since this initial discovery, our understanding of these processes has evolved to reflect the role of genetic alterations in cancer cells, including the activation of genes, such as lactate dehydrogenase A (LDHA), the enzyme responsible for the conversion of pyruvate to lactate, the final enzymatic step in the glycolytic pathway (3). Several studies have shown that LDHA plays a key role in tumor initiation and maintenance, and that inhibition of LDHA reduces cellular growth and metastasis in preclinical cancer models (4–9). Intriguingly, many glycolytic tumors display elevated levels of LDHA, and it appears to be expressed primarily in cancer cells, as opposed to normal tissues (10, 11). LDHA is therefore used as a biomarker for many malignancies (11) and is a promising target for cancer therapeutics.
To date, much of the data on LDH inhibition has been generated through proof-of-principle genetic studies that reveal LDHA knockdown inhibits in vitro and in vivo growth of certain cancer cell lines (4–7, 9). Although novel LDH inhibitors (LDHi) have been described, there are numerous issues with evaluating their translational potential, including suboptimal selectivity, potency, cellular permeability, and pharmacokinetic properties (4, 5, 12–14). Hence, the potential clinical applications of LDHi remain unrealized. Recently, renewed efforts have been made to more efficiently target LDH with agents developed through the National Cancer Institute Experimental Therapeutics (NExT) Program, a consortium that aims to develop drugs for difficult targets (15). NCI-737 and NCI-006 represent two novel LDHi that were developed and validated as part of the NExT Program.
In this study, we sought to evaluate the activity of NCI-737 and NCI-006, and to describe the impact of genetic and pharmacologic inhibition of LDH on cellular metabolism, growth, and survival on in vitro and in vivo preclinical models of Ewing sarcoma. Ewing sarcoma is an aggressive malignancy of the bones and soft tissues that primarily affects adolescents and young adults and is driven by a reciprocal oncogenic translocation between EWSR1 and an ETS family member such as FLI1 or ERG that results in aberrant gene expression (16–18). Ewing sarcoma remains a disease for which new therapies are critically needed, given that outcomes for high-risk patients remain poor and have not improved in decades (19–21). In this study, we show that Ewing sarcoma cells are exquisitely sensitive to inhibition of LDH activity, both genetically and pharmacologically. Further, glycolytic inhibition with either NCI-006 or NCI-737 impairs growth and survival of Ewing sarcoma in vitro and in vivo.
Materials and Methods
NCI-737 and NCI-006, equipotent against LDHA and LDHB, were obtained through the NExT Program (15). Details of compounds and their synthesis can be found in Supplementary Methods. Stock solutions for in vitro use were prepared in DMSO, aliquoted, and stored at -20°C. For in vivo use, powdered compound was dissolved in a volume of 0.1N NaOH, equivalent to 18% of the total solution volume, and added to PBS. Dropwise addition of 1N HCl was performed to achieve a pH of 7.4 to 7.8. Solution was prepared weekly and kept at 4°C.
Cell line screen
The cell line screen was performed by Oncolead using a panel of 94-cell lines and a 72-hour sulforhodamine assay.
Ewing sarcoma cell lines TC32, TC71, EW8, and RDES have been previously described (2). SK-N-MC and CHLA-258 were obtained from Dr. Lee Helman (Children's Hospital of Los Angeles, Los Angeles, CA). 5838 was obtained from the ATCC. Cell lines were authenticated by short tandem repeat DNA fingerprinting and compared with known sequences. TC32 (RRID: CVCL_7151), TC71 (RRID: CVCL_2213), EW8 (RRID: CVCL_V618), and CHLA-258 (RRID: CVCL_A058) were most recently authenticated in 2018 in the lab of Dr. Stephen Channock (National Cancer Institute, Rockville, MD). SK-N-MC (RRID: CVCL_0530) and RDES (RRID: CVCL_2169) were most recently authenticated in 2012 by Genetica Cell Line Testing. 5838 (RRID: CVCL_6255) has not been independently authenticated since purchase from the ATCC.
Mycoplasma testing of these cell lines was most recently performed in January 2019 and confirmed negative results. Experiments were performed on cells that were passaged between 5 and 12 times.
Cells were maintained in RPMI growth medium (Life Technologies) with 10% FBS (Sigma Aldrich), 100 U/mL penicillin and 100 μg/mL streptomycin (Life Technologies), and 2 mmol/L l-glutamine (Life Technologies) at 37°C in an atmosphere of 5% CO2.
Cell proliferation assays
Cellular proliferation was monitored in real-time using the IncuCyte live cell analysis system (Essen BioScience), and cellular viability was determined by MTT assay (Promega) according to the manufacturer's instructions. For both methods, cells were plated at a density of 2,000 cells/well in 96-well plates overnight and treated the following day.
Cell lysates were prepared by plating 1 million cells/10-cm plate overnight and then treating cells for an additional 24 hours. Cells were harvested with 1X RIPA (Santa Cruz Biotechnology) plus phosphatase and protease inhibitor cocktail (ThermoFisher). Protein lysates (30 μg/lane) were quantified by BCA protein assay (ThermoFisher). Blots were prepared as previously described (22) and incubated with primary antibodies, as described in Supplementary Methods.
Cells were plated at 1 million cells/10-cm plate and treated the following day. Cells were harvested using PBS-based, enzyme-free cell dissociation buffer (Life Technologies) at 72 hours and processed according to the Annexin V–FITC apoptosis detection kit (Sigma), before being run through the LSR Fortessa flow cytometer (BD Biosciences).
Lipofectamine RNAi Max (ThermoFisher) was used according to the manufacturer's instructions. Additional details of siRNA sequences and experimental conditions can be found in Supplementary Methods.
Chromatin immunoprecipitation sequencing analysis
FASTQ files from published FLI1 and H3K27ac chromatin immunoprecipitation sequencing (ChIP-seq), as well as associated RNA-seq experiments, were downloaded from NCI GEO (GSE8826 and GSE89026). Reads were aligned to the hg19 reference genome using BWA (version 0.7.10) using an established pipeline (23). Resulting tdf files were visualized using IGV (version 2.3.40).
The YSI 2950D Biochemistry Analyzer (Xylem Inc.) was used to measure glucose and lactate in the media of cells treated with NCI-737 (187 nmol/L) or DMSO (control). Cells were plated in triplicate in 24-well plates as follows: 500,000 cells/well for TC71 and EW8, 600,000 cells/well for RDES, and 250,000 cells/well for TC32. Media were collected after 24 hours of treatment, and the plate with cells was frozen at -80°C and subsequently lysed with 1X RIPA buffer (Santa Cruz Biotechnology) for protein assay.
Intracellular pyruvate concentrations were determined using the Pyruvate Assay Kit (Abcam). Cells were plated at 2 million/10-cm plate and treated with 187 nmol/L NCI-737 for 24 hours. Cells were lysed using 200 μL pyruvate sample buffer. Proteins were precipitated from the lysate with perchloric acid and neutralized with KOH according to the manufacturer's instructions. Pyruvate was measured using the fluorometric assay protocol.
Five thousand cells/well were plated in ViewPlate-96 96-well microplates (PerkinElmer) and treated with up to 187 nmol/L of NCI-737 for 13 hours. NAD+/NADH ratio was determined using the NAD+/NADH-Glo assay (Promega) according to the manufacturer's instructions.
Extracellular flux analysis
Analyses of cellular bioenergetics were performed using the Seahorse XFe96 Extracellular Flux Analyzer (Agilent). Glycolytic stress tests were performed according to the manufacturer's instructions, as previously described (24, 25). Additional experimental details can be found in Supplementary Methods.
LDH activity assay
Cells were plated overnight at 200,000 cells/well in 24-well plates. The following morning, cells were acclimated to XF complete media for 30 minutes prior to treatment with the indicated concentrations of LDHi or DMSO (control) for 1 hour (dose–response experiment) at 37°C in a non-CO2 incubator. For the time-point experiment, cells were incubated in XF complete media with the LDHi for the indicated time points. Cells were lysed using 100 μL LDH assay buffer and the oxidation of NADH measured spectrophotometrically at 340 nm.
Details of sample preparation for liquid chromatography mass spectrometry and nuclear magnetic resonance analyses can be found in Supplementary Methods. Liquid chromatography mass spectrometry analysis has been previously described (26), but further details of this and nuclear magnetic resonance analysis can be found in Supplementary Methods.
Statistical significance was determined by two-tailed Student t test. P < 0.05 was considered significant.
In vivo studies
Animal studies were approved by and performed in accordance with the National Institutes of Health Animal Care and Use Committee guidelines. For all experiments described, female Fox Chase SCID beige mice (CB17.B6-Prkdcscid Lystbg/Crl) were purchased from Charles River Laboratories. Two million cells in a solution of Hank's Balanced Salt Solution were injected orthotopically into the gastrocnemius muscle in the left hind leg of each mouse. Mice were randomized when palpable tumors developed, at which point treatment with agents began. Specific details for each set of experimental conditions may be found in Supplementary Methods.
Tumor volumes were compared between groups using a two-tailed Student t test at serial time points. P < 0.05 was considered significant. Measurements for mice that had already reached endpoint were carried forward until all mice in the group had reached endpoint.
Histopathology and image analysis
Hematoxylin and eosin–stained slides were scanned using the Aperio XT ScanScope whole slide imaging system with a 20X objective lens (Aperio). One whole slide image (SVS file) was prepared from each animal, capturing the entire tumor section on the slide. Details of image analysis can be found in Supplementary Methods.
Hyperpolarized 13C-MRI study was performed as described previously (27). Additional experimental details can be found in Supplementary Methods.
Intratumoral drug measurement
Measurement of tumor concentrations of LDHi was done by mass spectrometric analysis with internal standards (pure NCI-737 and NCI-006) and was performed by Quintara Discovery.
Intratumoral LDH activity measurement
Frozen tumor sections were pulverized in a liquid nitrogen–cooled pestle-mortar apparatus. Pulverized tumors were lysed in 10 volumes of LDH assay buffer and LDH activity measured as described above.
Ewing sarcoma cells are sensitive to genetic and pharmacologic inhibition of LDHA, which decreases cellular proliferation and activates apoptotic pathways
To evaluate the functional activity of NCI-737 and NCI-006, we screened a panel of 94 cancer cell lines using the Oncolead cell panel assay, which represents a diverse array of cancer cell lines. Across this panel, Ewing sarcoma cell lines emerged among the top 10 and top 12 most sensitive cell lines to NCI-006 and NCI-737, respectively, with IC50 values of 100 to 200 nmol/L (median IC50 values for all cell lines in the panel were 1,260 nmol/L for NCI-006 and 845 nmol/L for NCI-737; Fig. 1A). Of note, other pediatric-type sarcoma cell lines (rhabdomyosarcoma and osteosarcoma) in the panel had median IC50 values in line with the general panel (1,037 nmol/L and 712 nmol/L, for NCI-006 and NCI-737, respectively; Supplementary Table S1). To validate these findings, additional in vitro studies of cellular viability were performed on a broader panel of Ewing sarcoma cell lines. By MTT assay, all Ewing sarcoma cell lines displayed nearly identical dose-dependent sensitivity to NCI-737 and NCI-006, with IC50 values ranging from 100 nmol/L (TC71 and TC32) to 1 μmol/L (RDES and EW8) for each compound at 72 hours of treatment (Supplementary Fig. S1A). Given the potential for metabolic inhibitors to affect reagents in this assay, we verified these findings using IncuCyte live cell analysis, which confirmed that both NCI-737 and NCI-006 inhibited cellular proliferation at doses in the 100–500 nmol/L range across multiple cell lines (Fig. 1B; Supplementary Fig. S1B and S1C). Notably, the IC50 values for each of the compounds were very similar for each cell line tested, indicating equal potency of NCI-737 and NCI-006 in the Ewing sarcoma cell line models.
To evaluate the mechanisms behind this loss of viability, we performed Western blot analysis of cells treated with NCI-737, which revealed activation of proapoptotic proteins, cleaved PARP, and cleaved caspase 7 at 24 hours (Fig. 1C). Flow cytometry analysis of Ewing sarcoma cells treated with NCI-737 for 72 hours demonstrated a significant increase in the percentage of cells in both early and late apoptosis (Fig. 1D). Taken together, these findings suggest that the observed decrease in Ewing sarcoma cellular proliferation caused by LDHi is due to apoptotic cell death.
Next, we examined the result of genetic loss of LDHA and LDHB in Ewing sarcoma cell lines by knocking down the enzymes using siRNA to determine whether the results of pharmacologic LDH inhibition would be recapitulated. LDHA knockdown resulted in the expected loss of LDHA protein in Ewing sarcoma cell lines. As with the LDHi, genetic knockdown of LDHA resulted in decreased cellular proliferation and induction of apoptotic markers (Fig. 1E and F; Supplementary Fig. S2A and S2B), consistent with the conclusion that LDHA is important for Ewing sarcoma cell survival. In contrast, knockdown of LDHB with four siRNA sequences in multiple cell lines neither induced apoptosis nor negatively affected proliferation of Ewing sarcoma cells (Supplementary Fig. S2C–S2E).
LDHA is regulated by EWS-FLI1
Ewing sarcoma is characterized by a reciprocal translocation between chromosomes 11 and 22, resulting in the oncogenic transcription factor EWS-FLI1, which acts as the primary driver of the disease (17, 18). Although attempts to target EWS-FLI1 directly have yet to yield an effective therapeutic, examining the downstream effects of EWS-FLI1 has the potential to reveal insights into Ewing sarcoma biology that may be translated in clinically meaningful ways (28, 29). Given the marked sensitivity to both genetic depletion and pharmacologic inhibition of LDHA in Ewing sarcoma cells, as well as the recent findings that EWS-FLI1 regulates a shift away from oxidative metabolism toward glycolysis, which is NAD+-dependent (30, 31), we sought to determine whether EWS-FLI1 might play a role in the regulation of LDHA and/or LDHB. Depletion of EWS-FLI1 with multiple siRNA sequences resulted in a decrease in LDHA protein, as well as a decrease in NROB1 protein, a known direct target of EWS-FLI1, in each of the four Ewing sarcoma cell lines tested (Fig. 2A; Supplementary Fig. S3A), suggesting that LDHA expression is regulated, at least in part, by EWS-FLI1. In contrast, depletion of EWS-FLI1 had no effect on LDHB expression in each of the four Ewing sarcoma cell lines tested (Supplementary Fig. S3B).
To evaluate whether LDHA might be a direct EWS-FLI1 target, we examined publicly available ChIP-seq data generated using the shFLI1-transfected Ewing sarcoma cell line SK-N-MC (32). In the FLI1 knockdown, EWS-FLI1 and H3K27-acetylation deposition at the LDHA locus were decreased, and a corresponding decrease in LDHA mRNA expression was observed (Fig. 2B). An expected decrease in LDHA protein with EWS-FLI1 depletion in SK-N-MC was similarly observed (Fig. 2C). Furthermore, an analysis of publicly available ChIP-seq data on primary Ewing sarcoma tumors (33) also demonstrated H3K27ac deposition at the LDHA enhancer (Supplementary Fig. S3C). Analysis of ChIP-seq data from the same shFLI1-transfected Ewing sarcoma cell line SK-N-MC revealed that there was neither EWS-FLI1 nor H3K27-acetylation present at the LDHB locus (Supplementary Fig. S3D). Similarly, there was no change in LDHB protein with EWS-FLI1 depletion in this cell line (Supplementary Fig. S3E). Collectively, these data suggest that LDHA, but not LDHB, is a direct target of EWS-FLI1, and supports further investigation of the translational potential of pharmacologic LDH inhibition in Ewing sarcoma.
LDHi act through impairment of glycolysis in Ewing sarcoma cells
To evaluate on-target activity of these LDHi, we characterized the enzymatic inhibition and downstream metabolic effects of these agents. First, we assessed expression levels of LDHA and LDHB, and baseline LDH activity level in Ewing sarcoma cell lines, and determined there was uniform expression and enzymatic activity across the panel (Supplementary Fig. S4A and S4B). Next, we examined the change in basal LDH activity with increasing doses of NCI-737 or NCI-006 and found a dose-dependent inhibition of LDH activity in Ewing sarcoma cell lines for each of the compounds (Fig. 3A). For each of the cell lines tested (TC71, TC32, and EW8), the IC50 values for LDH inhibition were similar for both agents, at approximately 100 nmol/L, consistent with the observation that LDH appears to be uniformly expressed and active in these cell lines. Notably, the LDHi resulted in rapid inhibition of LDH upon exposure to Ewing sarcoma cells, with LDH activity decreasing within 30 seconds of drug exposure and reaching maximal inhibition by approximately 10 minutes in vitro (Supplementary Fig. S4C).
Next, biochemical analyses of glucose and lactate content in the culture media of treated and untreated cells were performed using YSI analysis and demonstrated a significant decrease in glucose consumption and lactate production in four Ewing sarcoma cell lines treated with NCI-737 (Fig. 3B). Furthermore, LDHi treatment induced an increase in intracellular pyruvate concentration in all four cell lines (Fig. 3C). Confirmatory biochemical studies, performed using mass spectrometry–based analysis of intracellular pools of pyruvate and lactate in Ewing sarcoma cells treated with LDHi, demonstrated a similar increase in steady-state levels of pyruvate and a decrease in steady-state levels of lactate in Ewing sarcoma cells treated with NCI-737 (Supplementary Fig. S5A and S5B).
We next examined the impact of LDHi treatment on trace labeling of intracellular pyruvate pools and lactate release using 13C-labeled glucose. In both Ewing sarcoma cell lines tested, we observed a significant increase in accumulation of 13C-labeled intracellular pyruvate (M+3) after 6 hours of treatment with NCI-737 (Supplementary Fig. S5C and S5D). We also noted a significant decrease in the amount of 13C-labeled lactate released into the media of treated cells, consistent with the data generated by YSI analysis (Supplementary Fig. S5E). Taken together, these data are consistent with NCI-737–mediated inhibition of LDH, resulting in accumulation of pyruvate that cannot be converted to lactate, resulting in a reduction of overall glucose consumption.
Because LDH is a major cytosolic regulator of redox homeostasis, we next examined the effect of NCI-737 on the cellular NAD+/NADH ratio. Treatment with NCI-737 resulted in a dose-dependent decrease in the NAD+/NADH ratio in all four Ewing sarcoma cell lines tested, compared with control (Fig. 3D), indicating that loss of NAD+/NADH homeostasis is a key outcome of treatment with these LDHi.
To further understand the bioenergetic effects of LDH inhibition on Ewing sarcoma cells, we utilized extracellular flux analysis to perform a glycolytic stress test (24, 25) on cells treated with NCI-737 to examine changes in extracellular acidification rate (ECAR). In both TC71 and TC32 cell lines, NCI-737 treatment for 6 hours resulted in a dose-dependent decrease in both glycolysis (change in ECAR upon glucose addition) and glycolytic capacity (change in ECAR upon oligomycin treatment), with the effect on glycolytic capacity being more pronounced (Fig. 3E; Supplementary Fig. S6A). Notably, upon stimulation of glycolysis by addition of oligomycin, untreated cells were able to increase ECAR, whereas this effect was blunted or lost completely in cells treated with NCI-737. A comparison of the effects of NCI-006 with those of NCI-737 on glycolytic flux demonstrated that the two agents inhibited ECAR in a nearly identical manner across a range of doses (Supplementary Fig. S6B).
In addition, we examined the effect of short-term NCI-737 treatment on oxidative phosphorylation by measuring the oxygen consumption rate (OCR) of Ewing sarcoma cells. When treated with NCI-737 at doses that affected ECAR, no changes in OCR were observed in Ewing sarcoma cells (Supplementary Fig. S7A and S7B). Further, when cells were exposed to higher doses of LDHi, concentrations between 10 and 30 μmol/L were required to produce a decrease in OCR, whereas decreases in ECAR were noted at concentrations starting at less than 1 μmol/L (Supplementary Fig. S7C and S7D). Taken together with the proliferation data, these findings suggest that at doses under 10 μmol/L these LDHi induce metabolic changes within the glycolytic pathway that directly affect the ability of the cells to proliferate and survive.
Pharmacologic effects on bioenergetics predict effect on proliferation in vitro
Although all Ewing sarcoma cell lines tested exhibited sensitivity to LDHi, there were slight differences noted in the degree of the antiproliferative effects. To understand the basis for this differential sensitivity, we interrogated the Ewing sarcoma cell lines for differences in various characteristics. As previously described, no correlations were noted between cell line sensitivity to LDHi and basal LDH activity, LDHA or LDHB expression, basal glucose consumption, basal pyruvate level, or basal NAD+/NADH ratio (Fig. 3B and D; Supplementary Fig. S4A and S4B). We did observe that the more sensitive cell lines had higher basal lactate production and a greater change in NAD+/NADH ratios upon drug treatment, suggesting that LDH may play a greater role in maintaining the redox balance in sensitive cells (Fig. 3B and D). In addition, further evaluation of extracellular flux data revealed a dose-dependent decrease in both glycolysis (Supplementary Fig. S8A) and glycolytic capacity (Supplementary Fig. S8B) in all cell lines tested, with the extent of change for each parameter varying across the cell lines (Fig. 4A and B). For example, when treated with NCI-737 at 250 nmol/L, TC71 cells underwent an 87% decrease in glycolysis and a 100% decrease in glycolytic capacity, whereas EW8 cells underwent a 19% decrease in glycolysis and a 38% decrease in glycolytic capacity. Notably, cell lines exhibiting larger decreases in glycolysis and glycolytic capacity with drug treatment were more sensitive to the effects of the drug on proliferation, whereas cell lines exhibiting smaller decreases required higher doses of NCI-737 for growth suppression (Fig. 4C). Thus, the magnitude of the biochemical effect of the LDHi on ECAR was predictive of functional response.
In addition, we examined cell lines for potential differences in their use of oxidative phosphorylation, because metabolic plasticity has been shown to affect sensitivity to LDH inhibition (34). To elucidate the relative contributions of the basal rates of glycolysis and oxidative phosphorylation, an energy map of basal and on-treatment ECAR and OCR was generated for several cell lines following 24-hour treatment with NCI-737 at 100 nmol/L (Fig. 4D). Interestingly, although all cell lines experienced an expected decrease in ECAR with treatment, the OCR response varied. The more sensitive cell lines (TC71 and TC32) experienced a relative decrease in OCR, whereas the less sensitive cell lines (RDES and EW8) experienced an increase in OCR, suggesting that increased oxidative phosphorylation may compensate, in part, for the loss of glycolysis in a subset of Ewing sarcoma cell lines, reflecting differences in sensitivity. In addition, data from the in vitro study using 13C-labeled glucose revealed a more robust increase of 13C-labeling of TCA cycle intermediates in EW8 compared with TC71 cells, consistent with a greater ability of EW8 cells to redirect pyruvate toward the TCA cycle upon inhibition of LDH (Fig. 4E).
LDH inhibition impairs glycolysis and affects cell survival in aggressive xenograft models of Ewing sarcoma
To evaluate the translational potential of NCI-737 and NCI-006, we treated several orthotopic xenograft models of Ewing sarcoma with the compounds to examine on-target activity, intratumoral drug concentration, toxicity, and efficacy. For all xenograft studies described, treatment was initiated after tumors became palpable. Initial in vivo studies were aimed at establishing the optimal route and dose of LDHi in orthotopic xenografts. Oral administration of the agents given at the MTD of 75 mg/kg total daily dose for NCI-737 and 50 mg/kg total daily dose for NCI-006 given for 3 weeks either once or twice daily to mice bearing TC71, TC32, or EW8 tumors resulted in minimal efficacy, although mice in the TC71 and TC32 groups treated with NCI-737 on the twice-daily schedule experienced a slightly decreased tumor growth rate (Supplementary Fig. S9A). Analysis of the effect of the compounds on LDH activity in the tumors revealed an inconsistent pattern of suppression of enzyme activity (Supplementary Fig. S9B). Furthermore, intratumoral drug levels were noted to be variable, establishing that oral dosing was insufficient to achieve adequate intratumoral drug levels and clinically relevant inhibition of LDH in these xenograft models (Supplementary Fig. S9C).
Given the limitations of oral dosing, follow-up studies were performed to evaluate IV dosing in TC71 tumor–bearing mice. NCI-737 was selected for the IV studies due to the slightly more efficacious antitumor effect noted in the oral dosing studies. Initial pilot studies were performed using doses of 25 and 40 mg/kg of NCI-737, given IV on a M/W/F schedule. At 25 mg/kg, no impact on tumor growth rate was observed, and at 40 mg/kg, a minimal change in tumor growth rate was noted (Supplementary Fig. S10A). Samples of plasma and tumor drug levels taken 4 hours after dosing revealed no differences in plasma concentrations (2.9 μmol/L for 25 mg/kg and 2.8 μmol/L for 40 mg/kg) and a slightly greater tumor concentration for the 40 mg/kg group (9 μmol/L vs. 10.3 μmol/L; Supplementary Fig. S10B and S10C). With 25 mg/kg dosing, no inhibition of LDH activity was noted in the tumors 4 hours after dosing; with 40 mg/kg dosing, 60% inhibition of LDH activity was seen at this time-point (Supplementary Fig. S10D).
Based on the lack of LDH inhibition seen at 25 mg/kg and the minimal efficacy observed with the 40 mg/kg dose given on the M/W/F schedule, we next examined doses of 40, 50, and 75 mg/kg given on a more frequent schedule of 5 consecutive days per week by IV to animals bearing TC71 tumors. At these doses, a dose-dependent reduction in tumor growth rates was noted, although the differences did not reach statistical significance (Supplementary Fig. S11A). However, at 75 mg/kg on this schedule, the mice displayed evidence of toxicity, so this dose could not be pursued further.
Pharmacokinetic and pharmacodynamic studies were performed on the animals in the 40 and 50 mg/kg dosing groups after tumor-bearing animals were treated with 3 consecutive days of IV NCI-737. Samples of tumor and plasma drug levels taken 1 hour after dosing revealed average intratumoral concentrations of 12.3 μmol/L (40 mg/kg group) and 18.5 μmol/L (50 mg/kg group) and average plasma concentrations of 8.9 μmol/L (40 mg/kg group) and 11.6 μmol/L (50 mg/kg group). (Fig. 5A; Supplementary Fig. S11B). Pharmacodynamic studies indicated 82% and 93% inhibition of intratumoral LDH activity in the 40 mg/kg and 50 mg/kg groups, respectively (Fig. 5B). Notably, intratumoral drug concentration and LDH inhibition rapidly diminished over a short interval. When animals treated at the same doses underwent tumor sampling 4 hours after dose (vs. 1 hour), a rapid loss (28%) of intratumoral compound and a corresponding decrease in the degree of LDH inhibition (82% to 58%) were observed between the 1- and 4-hour sampling times (Fig. 5C and D). These data were consistent with the known pharmacologic properties and full pharmacokinetic profile of NCI-737, which indicate a half-life of approximately 5 hours in plasma.
Given the rapid loss of LDH inhibition in the tumor, we hypothesized that the tumor growth response might be blunted due to inability to maintain continuous inhibition of tumor LDH on the 5 days/week schedule. To test this hypothesis, we evaluated a continuous 7 days/week dosing schedule. A daily dose of 60 mg/kg was chosen to attempt to minimize toxicity and maximize efficacy. NCI-737 treatment for 7 days/week resulted in statistically significant tumor growth suppression in mice bearing TC71 xenografts (Fig. 6A). In addition, histopathologic analysis of tumors revealed that treated tumors had a nearly 2-fold increase in necrosis over vehicle-treated tumors (59% vs. 33%). Similarly, tumors from EW8-bearing xenografts treated with NCI-737 at 60 mg/kg displayed a statistically significant increase in necrosis compared with control (43% vs. 28%; Fig. 6B and C; Supplementary Fig. S12A). However, treatment at this dose was not sufficient to affect overall tumor growth in the EW8 model, suggesting that the relative sensitivity differences noted in vitro may be relevant in vivo (Supplementary Fig. S12B).
Finally, to confirm the in vivo on-target activity of the LDHi, we performed hyperpolarized MR spectroscopy, using hyperpolarized 13C-pyruvate, of orthotopic xenograft Ewing sarcoma tumors in mice to quantify the ratio of 13C-lactate generated from 13C-pyruvate 30 minutes after a single IV dose of NCI-737 at the efficacious dose of 60 mg/kg. NCI-737 treatment resulted in a 57% and 49% reduction in the labeled lactate/pyruvate ratio in TC71 and EW8 tumors, respectively (Fig. 6D; Supplementary Fig. S13). Taken together, these findings verified the on-target activity of the agent in vivo and were consistent with the in vitro results. Furthermore, the in vivo data indicate that to achieve tumor growth inhibition, sustained inhibition of LDH through frequent dosing at the highest tolerable dose is required.
Hemolysis is the primary dose-limiting toxicity of LDH inhibition
Assessments of toxicity via observations of general appearance, blood parameters, and necropsy were performed on animals treated via a 5 days/week IV dosing schedule with NCI-737 at 40, 50, or 75 mg/kg. Toxicity assessments were performed at day 3 and at tumor or humane endpoint. Although mice tolerated treatments without weight loss (Supplementary Fig. S14A), a dose-dependent intolerance of NCI-737 was noted. All mice treated at 40 mg/kg displayed no toxicity, whereas 1 of 3 mice treated at 50 mg/kg and 2 of 3 mice treated at 75 mg/kg displayed decreased physical activity by day 2 of treatment. Laboratory and pathologic assessments of all mice revealed that hemolysis was the primary dose-limiting toxicity observed, with a dose-dependent decrease in hemoglobin noted within 3 days of starting treatment (Supplementary Fig. S14B). Within 3 days, hemoglobin values began to fall below the lower limit of the normal range of 11 g/dL in mice receiving the lowest (40 mg/kg) dose. At higher doses, hemoglobin values were below 8 g/dL. By endpoint, average hemoglobin levels had decreased further to 9.1, 4.6, and 5.3 g/dL in the 40, 50, and 75 mg/kg treated groups, respectively, indicating that toxicity was cumulative with continued dosing.
A corresponding rise in total bilirubin was also noted both at day 3 of treatment and at endpoint (Supplementary Fig. S14C). Although measurements of direct bilirubin were not obtained, normal liver studies were noted in all mice, suggesting that the rise in total bilirubin was the result of an increase in the indirect bilirubin fraction, which is consistent with red blood cell breakdown. In addition, mice treated at all dose levels were noted to have an increase in splenic weight at endpoint, further supporting hemolysis as the major toxicity (Supplementary Fig. S14D). Red blood cells lack nuclei and mitochondria and therefore rely entirely on glycolysis for bioenergetics (35). A previous study describing the phenotypes associated with genetic loss of LDHA in murine models reported nonlethal hemolytic anemia as a main finding, supporting the idea that this toxicity is on-target (4). Of note, no other lab abnormalities were seen on complete blood counts or chemistry panels performed as part of the toxicity assessment.
In this study, we have demonstrated that Ewing sarcoma cells display a marked dependency on LDHA, due, in part, to the regulation of LDHA by the oncogenic transcription factor EWS-FLI1. We have shown that genetic or pharmacologic inhibition of LDHA reduces proliferation and induces apoptosis in Ewing sarcoma cells, and that this is associated with suppression of glycolytic flux and perturbation of the NADH/NAD+ ratio. In addition, we have explored the translational potential of this target through in vivo characterization of two novel LDHi, performing analyses of drug delivery routes, pharmacokinetics, pharmacodynamics, toxicity, and efficacy.
We show that LDHA is an important enzyme for the survival of Ewing sarcoma cells, which are remarkably sensitive to both genetic and pharmacologic inhibition of LDH. Pharmacologic targeting of LDH using NCI-737 and NCI-006 inhibited proliferation of Ewing sarcoma cells both in vitro and in vivo through inhibition of glycolysis and induction of apoptosis. This finding is consistent with published literature describing targeting of LDH in other sensitive cancer types (9, 36). Notably, the effect of these agents on proliferation of Ewing sarcoma cells exceeded the effects seen on other types of cancer cells. Our work suggests that the biology of Ewing sarcoma, in particular the presence of the driving EWS-FLI1 fusion oncogene (17, 18), contributes to the sensitivity of these cells to glycolytic inhibition. Specifically, we have shown that the EWS-FLI1 fusion oncoprotein directly regulates LDHA, but not LDHB, expression. This finding is supported by the recent report that EWS-FLI1 regulates metabolic pathways in Ewing sarcoma, where it shifts glucose consumption away from oxidative metabolism and toward glycolysis (30), and by evidence that Ewing sarcoma cells are highly glycolytic and highly susceptible to glucose deprivation and disruption of glucose metabolism (31, 37–40). In addition, there are clinical data to suggest that outcomes for Ewing sarcoma patients are worse for those who exhibit higher plasma levels of LDH (41–43).
The metabolic consequences of NCI-737 and NCI-006 on Ewing sarcoma were suppressed glycolytic flux through inhibition of the conversion of pyruvate to lactate, which was observed in both in vitro and in vivo models, concomitant with disruption of the NAD+/NADH ratio. By extracellular flux analysis, NCI-737 treatment resulted in decreased ECAR, the magnitude of which was predictive of the effect on cellular viability across multiple Ewing sarcoma cell lines. The ability of the cells to engage oxidative phosphorylation through the TCA cycle, which has been reported as a resistance mechanism in other cancer types (7, 34), also emerged as a potential factor contributing to differential sensitivity to LDHi.
Given the critical need for novel therapies for patients with Ewing sarcoma (19–21), the remainder of our work was focused on determining the translational potential of LDHi for this disease. Although suppression of glycolysis was profound in the single treatment MR spectroscopy experiments, the long-term in vivo studies revealed that effect on xenograft tumor growth was dependent on achievement of consistently high and sustained levels of the inhibitors intratumorally, which required frequent IV dosing. This requirement may also explain why a maximal intratumoral drug concentration in the micromolar range could result in less antitumor activity than would be expected based on the in vitro dosing range. Although an increase in the delivered dose or a more frequent dosing interval may have been able to overcome this, the on-target dose-dependent hemolytic anemia observed in the mice limited further dose-escalation. Thus, there appears to be a narrow therapeutic window for these LDHi to move forward into clinical development as single agents. Strategies to overcome these challenges include development of novel delivery methods for LDHi, or combinatorial approaches that might allow for lower doses of LDHi to be effective. Based on published data, kinase inhibitors, HSP90 inhibitors, oxidative phosphorylation inhibitors, and inducers of reactive oxygen species represent several potential combination partners worthy of further study (5, 44–47). In conclusion, our findings suggest that further translational preclinical work is necessary to optimize the potential of LDHi as anticancer agents, but that due to their exquisite sensitivity to glycolytic impairment, Ewing sarcoma may be one of the best indications for their use.
Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
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Conception and design: C. Yeung, A.E. Gibson, S.H. Issaq, A. Mendoza, L.J. Helman, V. Darley-Usmar, L.M. Neckers, C.M. Heske
Development of methodology: C. Yeung, A.E. Gibson, S.H. Issaq, J.T. Baumgart, D.J. Urban, G.A. Benavides, G.L. Squadrito, T. Dowdy, V. Darley-Usmar, L.M. Neckers, C.M. Heske
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): C. Yeung, A.E. Gibson, S.H. Issaq, N. Oshima, J.T. Baumgart, D.J. Urban, M.S. Johnson, G.A. Benavides, S. Eldridge, T. Dowdy, V. Ruiz-Rodado, A. Mendoza, J.F. Shern, G.M. Stott, M.C. Krishna, M.D. Hall, C.M. Heske
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): C. Yeung, A.E. Gibson, S.H. Issaq, N. Oshima, J.T. Baumgart, D.J. Urban, M.S. Johnson, G.A. Benavides, G.L. Squadrito, M.E. Yohe, H. Lei, S. Eldridge, J. Hamre III, T. Dowdy, V. Ruiz-Rodado, A. Lita, A. Mendoza, J.F. Shern, M. Larion, L.J. Helman, G.M. Stott, M.D. Hall, V. Darley-Usmar, L.M. Neckers, C.M. Heske
Writing, review, and/or revision of the manuscript: C. Yeung, A.E. Gibson, S.H. Issaq, J.T. Baumgart, G. Rai, D.J. Urban, M.S. Johnson, G.A. Benavides, G.L. Squadrito, M.E. Yohe, V. Ruiz-Rodado, A. Lita, J.F. Shern, M. Larion, L.J. Helman, M.D. Hall, V. Darley-Usmar, L.M. Neckers, C.M. Heske
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): S.H. Issaq, L.D. Edessa, G.L. Squadrito, T. Dowdy, A. Mendoza, G.M. Stott, C.M. Heske
Study supervision: C. Yeung, M.D. Hall, V. Darley-Usmar, C.M. Heske
Other (conception, design, and synthesis of the inhibitors used in this study): G. Rai
Other (I performed the computational analysis for quantifying percent necrosis in the tumors and wrote the M&M for that specific part only): J. Hamre III
Authors were supported by grants from the Intramural Research Program of the NIH as follows: C. Yeung, A.E. Gibson, S.H. Issaq, N. Oshima, J.T. Baumgart, L.D. Edessa, M.E. Yohe, H. Lei, T. Dowdy, V. Ruiz-Rodado, A. Lita, A. Mendoza, J.F. Shern, M. Larion, L.J. Helman, M.C. Krishna, L.M. Neckers, and C.M. Heske. N. Oshima, J.T. Baumgart, D.J. Urban, M.S. Johnson, G.A. Benavides, G.L. Squadrito, M.E. Yohe, H. Lei, S. Eldridge, J. Hamre III, T. Dowdy, V. Ruiz-Rodado, A. Lita, A. Mendoza, J.F. Shern, M. Larion, L.J. Helman, G.M. Stott, M.D. Hall, V. Darley-Usmar, L.M. Neckers, and C.M. Heske were supported by the NCI, Center for Cancer Research; G. Rai, D.J. Urban, and M.D. Hall were supported by the National Center for Advancing Translational Sciences; S. Eldridge was supported by the Developmental Therapeutics Program, Division of Cancer Treatment and Diagnosis. This project has also been funded in part with Federal funds from the NCI, NIH, under Contract No. HHSN261200800001E (J. Hamre III and G.M. Stott) and with funding from the Chemical Biology Consortium, NCI Experimental Therapeutics (NExT) Program (N. Oshima, G. Rai, D.J. Urban, M.S. Johnson, G.A. Benavides, G.L. Squadrito, S. Eldridge, M.C. Krishna, M.D. Hall, V. Darley-Usmar, and L.M. Neckers).
The authors would like to thank Devorah Gallardo, Rebekah Madrid, Dr. Daniel Crooks, Kristin Beebe, Dr. David Venzon, and Matilda Culp for their assistance with this study.
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