MCT1 Inhibitor AZD3965 Increases Mitochondrial Metabolism, Facilitating Combination Therapy and Noninvasive Magnetic Resonance Spectroscopy

Metabolic “re-wiring” is a hallmark of cancer that enables malignant cells to meet their growth demands, survive, and thrive in a hostile tumor microenvironment. The Warburg effect describes tumor cell preferential use of the glycolytic pathway to generate lactate under aerobic conditions. This process leads to excess production of lactate, which is removed from cells via monocarboxylate transporters (MCT), a family of transmembrane proteins (including MCT1, MCT2, MCT3, and MCT4) that mediate the proton-linked bidirectional movement of lactate (as well as pyruvate and ketone bodies) in and out of cells (1). The excreted lactate not only prevents intracellular acidification-driven cell toxicity but is believed to aid in tumor invasion, immune suppression as well as providing lactate as a fuel source for the more oxidative neighboring tumor cells (2–4).

MCT1 and MCT4 are overexpressed in cancer, and disrupting lactate transport represents a promising approach for antineoplastic therapeutic targeting (2, 4–6). Indeed, several inhibitors of MCT1 have recently been described that have shown promising preclinical activity, including the dual MCT1/MCT2 inhibitor AR-C155858 (5) and the MCT1 inhibitors SR13800 (7) and AZD3965 (8).

AZD3965 is a selective MCT1 inhibitor that exhibits some activity against MCT2 (but not MCT3 or MCT4), showing growth-inhibitory activity in human gastric and small cell lung cancer tumor xenografts in vivo (8, 9). AZD3965 is currently in phase I and progressing to phase II clinical trials in the UK (www.clinicaltrials.gov), and understanding its impact on cellular metabolism as well as identifying pharmacodynamic (PD) biomarkers for MCT1 modulation is much needed for furthering the clinical development of this and other MCT1-targeted drugs.

In this context, metabolic profiling approaches enable the interrogation of tumor metabolism and, as such, lend themselves to PD biomarker discovery and evaluation of anticancer drugs targeting metabolism, including AZD3965. NMR spectroscopy is one technique that enables the steady-state and dynamic assessment as well as in vivo study of cellular metabolism. The technique has shown great promise for mechanism-based PD metabolic biomarker discovery following targeted cancer therapy, facilitating the translation of studies from experimental cancer models through to patients (10–12). For example, depletion of hexokinase-2 and CMYC following treatment with an MEK inhibitor in BRAF-driven human melanoma cells and xenograft tumors resulted in decreased lactate levels as observed with ¹H NMR (13). Further, PI3K inhibitor–mediated depletion of lactate dehydrogenase expression led to a fall in hyperpolarized ¹³C-pyruvate–lactate exchange, assessed by dynamic ¹³C NMR of human cancer cells and tumors (14, 15).

MCT1 inhibition with AZD3965 has been shown to activate glycolytic metabolism, increasing glycolytic intermediate levels (e.g., glucose 6-phosphate and fructose 6-phosphate) as well as enzyme activity (e.g., hexokinase and pyruvate kinase) concomitant with intracellular lactate (Lactate_I) accumulation (9). However, a recent study in human breast cancer cells showed that AZD3965 has no impact on glycolytic activity or Lactate_I and that its antitumor effects are due to inhibition of pyruvate export (16). It is not clear whether the discrepancies in reported effects are due to variations in baseline MCT4 levels, how AZD3965 affects downstream metabolic pathways and fluxes, and whether any of the drug-induced metabolic changes could have potential as noninvasive PD biomarkers of target inhibition.

In this study, using human lymphoma and colon carcinoma cells with varying baseline MCT4 expression, we show that AZD3965 inhibits monocarboxylate transport in and out of cells concomitant with increased bioenergy-related metabolite levels in MCT4⁻ as well as MCT4⁺ cells. The latter effect is attributed to activation of mitochondrial metabolism via oxidative pyruvate dehydrogenase (PDH) flux, which was accompanied by a relative increase in anaplerotic pyruvate carboxylase (PC) flux. The increased mitochondrial metabolism enabled continued cell survival under drug treatment, which was prevented by coadministration of the mitochondrial complex I inhibitor metformin and pyruvate carrier inhibitor UK5099. Our data also show that the improved bioenergetics are detectable in a tumor xenograft model, using in vivo³¹P NMR spectroscopy, indicating potential for noninvasive evaluation of drug action.

Raji and Hut78 human lymphoma cells were purchased from the ATCC. Kelly and IMR-32 human neuroblastoma and HT29 human colon carcinoma cells were originally obtained from the ATCC and authenticated in our laboratory using STR profiling on October 16, 2015, and March 13, 2017, respectively. Lymphoma and neuroblastoma cells were grown in RPMI, whereas HT29 cells were grown in DMEM as previously described (17). These cell lines were chosen to provide a range of baseline expression of MCT4, with lymphomas being also a target for the phase I trial expansion cohort and neuroblastoma (MYCN-amplified) representing a solid tumor type expected to be MCT1-dependent (7). AZD3965 was provided by AstraZeneca. Metformin and UK5099 were purchased from Sigma-Aldrich.

Cell counts were obtained using a Beckman Coulter Vi-Cell Cell Viability Analyzer. Drug treatment effects on growth were determined using a CellTiter-Blue Viability assay (Promega) following 72-hour incubation, or by cell counting after 24-, 48-, and 72-hour treatment using Trypan blue exclusion of dead cells.

Cells were treated with either DMSO (0.01%) or AZD3965 at the indicated concentrations in fresh media and cells and/or spent media collected at the indicated time points for analysis. To assess changes in glucose metabolism, cells were incubated in media containing 100% 5 mmol/L [1-¹³C]glucose for 6 hours in drug or DMSO, followed by cell and culture media collection for ¹³C NMR.

Raji xenograft tumors were established by subcutaneous injection of 5 × 10⁶ Raji cells in 0.1 mL of a 1:1 medium:matrigel solution into the flank of female SCID mice (6–8 weeks old). Tumors were measured as previously described (17), and once they reached approximately 450 mm³ in volume (after ∼3–4 weeks), mice were randomized into two groups; one group was treated with vehicle (85% saline, 10% DMSO and 5% tween, n = 7) and the other with 50 mg/kg AZD3965 dissolved in vehicle (n = 6) administered orally twice daily, a schedule based on a previously published report (8). For in vivo NMR spectroscopy, mice received three doses in total: two doses on day 1 and one dose on day 2. For growth inhibition measurements, a separate cohort of mice bearing tumors of approximately 350 mm³ in volume was treated with either vehicle (n = 4) or AZD3965 (n = 3) as above for a total of 5 days and tumor volume monitored. All experimental protocols were monitored and approved by the ICR Animal Welfare and Ethical Review Body Animal Welfare and Ethical Review Body, in accordance with UK Home Office regulations under the Animals (Scientific Procedures) Act 1986 and UK National Cancer Research Institute Guidelines for the Welfare and Use of Animals in Cancer Research (18).

Mice were anesthetized as previously described (17) and placed at the isocenter of a horizontal Bruker 7 Tesla microimaging system (Bruker Instruments) with the tumors positioned in the center of a 15 cm dual tune¹H/³¹P surface coil. After shimming (achieving a linewidth on the unsuppressed water peak of ∼20 Hz on average), localized image-selected ³¹P NMR spectroscopy measurements of the tumors were performed prior to day 0 and 2 hours following the last dose (on day 2). Data were processed using jMRUI and AMARES fitting to determine chemical shift positions and peak integrals as previously described (19). Intracellular tumor pH was calculated based on the chemical shift difference between the inorganic phosphate (Pi) and α-nucleotide triphosphate (α-NTP) peaks as previously reported (19). After the last scan, the experiment was terminated with a schedule 1 method, and tumors were excised, immediately snap-frozen in liquid nitrogen, and then stored at −80°C until further processing.

Cells and xenograft tumor tissue were extracted as previously described (20), and freeze-dried samples were reconstituted in 540 μL of D₂O containing 3-(trimethylsilyl) propionic-2,2,3,3-d₄acid (TSP) as an internal ¹H NMRquantitation and chemical shift reference. For extracellular metabolite analysis, 1 mL of media was freeze-dried and reconstituted in D2O+TSP as above. ¹³C NMR data were semiquantified (AU/cell) relative to the TSP integral, using the [3-¹³C]lactate peak at 21.4 ppm for chemical shift referencing. For ³¹P NMR of cell extracts, 60 μL of a D₂O solution containing EDTA and methylenediphosphonic acid (internal standard) to a final concentration of 10 and 0.43 mmol/L respectively (pH 8.2) was added. Further details of data acquisition and processing are provided in the Supplementary Methods.

¹³C-pyruvate–lactate exchange was monitored in intact Raji, Hut78, and HT29 human cancer cells exposed to DMSO or AZD3965 for 1 or 24 hours. For this, cell pellets (10–15 × 10⁶ cells/sample) were washed in saline, incubated at 37°C in FBS-free medium, and ¹³C spectra acquired for 4 minutes with 2-second intervals and a 10° pulse immediately after the addition of 10 mmol/L hyperpolarized [1-¹³C]pyruvic acid and 10 mmol/L unlabeled lactate in a total volume of 0.5 mL. Similar cell handling conditions were previously shown to be adequate for maintaining cell viability and metabolic activity (21) and to produce comparable data to perfused cell studies when looking at drug effects (14, 22, 23).

Dynamic hyperpolarized ¹³C time-series spectra were summed, and the ratio of the areas under the curve for the lactate and pyruvate peaks (Lactate_AUC/Pyruvate_AUC), a parameter shown to accurately reflect pyruvate–lactate exchange (22), was calculated.

Protein expression was performed using Western blotting under standard conditions. Details of the primary and secondary antibodies are provided in Supplementary Methods.

A single comparison unpaired Student t test (for comparison of cell number and metabolite levels between control and treated cells) or a Wilcoxon matched-pairs signed rank test (for tumor volume and in vivo metabolite ratio changes prior to and following treatment) was used, with P < 0.05 considered statistically significant. Data represent the mean ± SE.

The direction of transport of lactate follows a lactate/H+ concentration gradient (1). In glycolytic cells incubated in fresh media, increased intracellular production of lactate favors export out of cells, leading to accumulation upon MCT1 blockade (MCT1 being the dominant transporter in the absence of alternative transporters such as MCT4). Thus, prior to investigating the downstream metabolic effects of MCT1 inhibition, we assessed the impact of AZD3965 on Lactate_I, as an indicator of MCT1 blockade, in MCT4⁻ Raji human lymphoma cells. We also examined the temporal dynamics of the response as well as its dependence on drug concentration.

Our data indicate that at 24 hours, AZD3965 induced accumulation of Lactate_I that was observed with as little as 2 nmol/L AZD3965 (P = 0.01) and became more pronounced with increasing drug concentrations, reaching a maximum at 25 nmol/L AZD3965 (Fig. 1A) and plateauing thereafter. Time course analysis with 25 nmol/L AZD3965 revealed rapid build-up of Lactate_I, showing a trend toward an increase as early as 15 minutes, becoming significant after 90 minutes (P = 0.02), peaking at around 3 hours (up to 1,890 ± 230%, P = 0.002) and then showing a gradual decrease from 6 up to 72 hours after treatment (P ≤ 0.03; Fig. 1B). Extracellular lactate (Lactate_E) levels also decreased compared with control at 24 hours and up to 72 hours after treatment (Fig. 1C). Raji cell counts were reduced relative to control with exposure to AZD3965; however, cell growth persisted despite the increase in Lactate_I (Fig. 1D), possibly suggesting activation of adaptive pathways.

We next tested whether AZD3965 also inhibited MCT1-mediated substrate transport into cells, and the relationship between Lactate_I accumulation and MCT1 inhibition in the presence or absence of MCT4, an alternative mediator of lactate efflux and a predictor of resistance to AZD3965 (8). For this, we used MCT4⁻ Raji cells and two additional human cancer cell lines expressing varying levels of MCT4: Hut78 lymphoma (low MCT4: MCT4⁺) and HT29 colon carcinoma (high MCT4: MCT4⁺⁺⁺; Fig. 2A).

Twenty-four-hour exposure to a low concentration of AZD3965 (5 nmol/L) led to a 10-fold build-up in Lactate_I in MCT4⁺ Hut78 human lymphoma cells, which was sustained at higher drug concentrations (25 and 500 nmol/L; Fig. 2B). In contrast, MCT4⁺⁺⁺ HT29 human colon carcinoma cells showed a trend toward an increase in Lactate_I relative to controls with 5 and 25 nmol/L AZD3965, which only became significant with exposure to 500 nmol/L AZD3965 reaching up to 4-fold, relative to controls, i.e., much smaller in amplitude compared with Raji and Hut78 cells (Fig. 2B). Lactate_E levels were only marginally affected in MCT4⁺ Hut78 and MCT4⁺⁺⁺ HT29 cells (Fig. 2C), in keeping with the presence of the alternative lactate transporter MCT4.

To assess the effect of AZD3965 on monocarboxylate transport dynamics, we used hyperpolarized [1-¹³C]pyruvate, also transported into cells via MCT1 (1), and monitored its exchange with lactate in our three cell lines.

Figure 2D shows representative ¹³C spectra of summed data with signals from hyperpolarized ¹³C-lactate, pyruvate, and pyruvate hydrate and the time-dependent changes in hyperpolarized ¹³C-lactate signals obtained from control- and AZD3965-treated Raji cells. Representative time-series spectra are also provided in Supplementary Fig. S1. Following 24 hours of treatment, the Lactate_AUC/Pyruvate_AUC ratio, reflective of ¹³C-pyruvate–lactate exchange (22), decreased in a concentration-dependent manner to 31 ± 6% in 5 nmol/L AZD3965-treated and 19 ± 2% in 25 nmol/L AZD3965-treated Raji cells relative to controls (P < 0.001, n = 6), indicating inhibition of [1-¹³C]pyruvate uptake with AZD3965 (Fig. 2E). Incubation of Raji cells for 1 hour with 25 nmol/L AZD3965 also reduced Lactate_AUC/Pyruvate_AUC consistent with the immediate action of the drug on MCT1 (Fig. 2E). Further, AZD3965 inhibited pyruvate–lactate exchange in MCT4⁺ Hut78 cells and in MCT4⁺⁺⁺ HT29 cells even at drug concentrations that had no significant effect on intracellular lactate (Fig. 2B and E), in line with MCT1-mediated transport of [1-¹³C]pyruvate being a rate-limiting step in the ¹³C NMR-observed reaction (24). MCT4⁺⁺⁺ HT29 cells did however exhibit the smallest inhibition of pyruvate–lactate exchange, suggesting that the experimental conditions used may be permissive of some MCT4-mediated [1-¹³C]pyruvate transport into cells.

Assay of the activity of lactate dehydrogenase, the intracellular enzyme that catalyzes the conversion of pyruvate to lactate, showed no significant differences between control- and AZD3965-treated Raji cells (Supplementary Fig. S2). Thus, the reduction in pyruvate–lactate exchange observed with AZD3965 appears to be driven primarily by reduced ¹³C-pyruvate transport following MCT1 blockade.

These data confirm inhibition of MCT1 by AZD3965 in Raji, Hut78, and HT29 cells and indicate that the observed intracellular lactate build-up is a result of its decreased transport out of cells (and not increased influx), which is highly dependent on the near or complete absence of MCT4 expression.

To examine the downstream consequences of MCT1 inhibition on additional aspects of metabolism, we analyzed the ¹H NMR spectra of control- and AZD3965-treated Raji cell extracts. The data revealed significantly increased levels of branched-chain amino acids (BCAA), acetate, NADH + NAD⁺, ribose, succinate, fumarate, and ADP+ATP following AZD3965 exposure. Conversely, glutamate and phosphocholine levels were significantly reduced in AZD3965-treated Raji cells relative to controls (Table 1).

The increase in bioenergetic metabolites was further confirmed with ³¹P NMR, demonstrating increased β-NTP levels in Raji cells (Table 1). ³¹P NMR also revealed an increase in glycerol 3-phosphate (G 3-P) levels, consistent with earlier reports (7) indicating inhibition of the glycolytic pathway after exposure to AZD3965 treatment (Table 1).

We also evaluated changes in the metabolic profile of MCT4⁺⁺⁺ HT29 cells, given that they too displayed a build-up (up to 4-fold) in Lactate_I with 500 nmol/L AZD3965. ¹H NMR analysis revealed increased BCAA, alanine, methionine, glutamine, malate, ribose, creatine+phosphocreatine, and ATP+ADP levels in AZD3965-treated relative to control cell extracts (Table 1). Thus, in both MCT4⁻ Raji and MCT4⁺⁺⁺ HT29 cells, treatment with AZD3965 at concentrations that led to Lactate_I build-up was associated with improved bioenergetics and increased TCA cycle intermediates. Elevated β-NTP and G 3-P levels were also observed in MCT4⁺ Hut78 cells treated with 500 nmol/L AZD3965 for 24 hours [up to 129 ± 11% (P = 0.02) and 148 ± 12% (P = 0.01) of controls, respectively]. A bioluminescent assay of ATP confirmed the increased levels following exposure of Raji and Hut78 cells to AZD3965 (Supplementary Fig. S3).

The metabolic basis underlying the improved bioenergetic state and increased TCA cycle intermediates observed with AZD3965 was next investigated by initially assessing mitochondrial abundance using confocal microscopy. MitoTracker Red staining showed no difference between control- and AZD3965-treated Raji cells, indicating no changes in functioning mitochondrial density (Supplementary Fig. S4).

Next, we monitored the fate of [1-¹³C]glucose in Raji cells grown in the presence or absence of drug. ¹³C NMR analysis of culture media following incubation in [1-¹³C]glucose indicated that AZD3965 decreased glucose uptake and Lactate_E, consistent with reduced glycolytic metabolism (Fig. 3A). ¹³C NMR of intracellular metabolites in treated relative to control cells revealed a significant increase in [1-¹³C]glucose, indicating impaired utilization, alongside a substantial rise in [3-¹³C]lactate, consistent with the rapid blockade of its transport, thus reducing export out of cells. Interestingly, these changes were accompanied by significantly elevated ¹³C-labeling of glutamate at the C4 position, reflecting PDH flux. ¹³C-labeling of glutamate at the C2 and C3 positions (reflecting PC flux) remained unchanged (Fig. 3B and C), but given the decreased consumption and incorporation of ¹³C-glucose into the pathway following treatment, the flux through PC is therefore relatively increased to maintain [2-¹³C] and [3-¹³C]glutamate at the same level as in control cells. There was a trend toward a fall in [3-¹³C]alanine, but this was not statistically significant (P = 0.1). Taken together, these results show that AZD3965 treatment blocks lactate transport out of cells, impairs glycolytic activity, and increases routing of glucose toward mitochondrial metabolism through oxidative energy producing PDH flux and, to a lesser extent, anaplerotic PC flux.

The fact that AZD3965 stimulates mitochondrial oxidative PDH flux, leading to increased cellular bioenergy levels, suggests acquisition of a metabolic advantage enabling cells to continue growing under MCT1 blockade (Fig. 1D). Thus, we hypothesized that stemming this capability, by blocking mitochondrial metabolism, may enhance the antiproliferative effects of AZD3965.

Growth inhibition data from Raji cells (Fig. 4A) revealed that the GI₅₀ value of AZD3965 over 72 hours decreased from 3.91 ± 1.37 nmol/L, when given as single agent, to 1.03 ± 0.42 nmol/L when combined with the mitochondrial complex I inhibitor metformin (n = 4, P = 0.005), consistent with greater potency. Cell counts over 72 hours confirmed this effect and indicated that the reduced numbers are due to increased cell death (Fig. 4B). The efficacy of AZD3965 was also increased when it was combined with metformin in MCT4⁺ Hut78 cells, although the effects were less pronounced relative to MCT4⁻ Raji cells (Fig. 4C).

In neuroblastoma cells harboring MYCN amplification and as such representing a solid tumor type expected to be MCT1- dependent (7), AZD3965′s therapeutic activity was increased with metformin cotreatment in both MCT4⁺ Kelly and MCT4⁻ IMR-32 cells (Supplementary Fig. S5A–S5C). IMR-32 had a very slow doubling time (40–48 hours compared with 20–24 hours in Kelly cells), which may account for the relatively smaller changes in cell counts over the 72-hour treatment. The potentiation of AZD3965′s activity by metformin was, at least in Kelly cells, also underpinned by a shift toward mitochondrial metabolism following MCT1 inhibition (as shown by increased fumarate and succinate; Supplementary Fig. S5D). AZD3965 caused a relatively modest increase in Lactate_I in Kelly cells, likely due to MCT4 coexpression (Supplementary Fig. S5A and S5D).

Furthermore, blockade of pyruvate transport into mitochondria by the mitochondrial pyruvate carrier (MPC) inhibitor UK5099 dose-dependently enhanced the potency of AZD3965 in Raji cells, leading to markedly increased cell killing when the two agents were combined relative to when AZD3965 was given alone (Fig. 4D). Similar, albeit less profound, effects were also observed in Hut78 after 72 (Fig. 4E) and 96 hours (Fig. S6) of treatment. In both metformin and UK5099 combination treatments of Raji cells, we observed increased PARP cleavage consistent with induction of apoptosis (Fig. 4F).

Thus, the AZD3965-induced increase in mitochondrial metabolism is required for maintaining cell survival under therapeutic stress, enabling combination therapy with mitochondrial metabolism inhibitors.

Next, to assess the relevance of the in vitro–observed changes to in vivo tumor metabolism and their potential as noninvasive biomarkers of AZD3965 activity, we evaluated changes in Raji xenograft tumor bioenergetic metabolism using in vivo³¹P NMR spectroscopy.

Over an acute 2-day treatment schedule (Fig. 5A), AZD3965 had no significant effect on tumor growth relative to vehicle (percentage change in day 2/day 0 tumor volumes was 122 ± 6% vs. 120 ± 9% for AZD3965 and vehicle groups, respectively (P = 0.85)]. However, after 5 days of therapy, the treated tumor volumes remained at 123 ± 12% of pretreatment values (P = 0.2) as observed in the acute protocol, consistent with cytostasis, whereas the vehicle-treated tumors grew up to 171 ± 19% (P = 0.036) relative to pretreatment volumes (Fig. 5B).

We observed no differences in percentage of necrotic fractions or perfusion between the 350 and 450 mm³-sized tumors (representative of the acute vs. 5-day treatment experiments; Supplementary Fig. S7), and the effect of drug on posttreatment tumor size in the two schedules was also similar, indicating that the two settings are comparable.

In vivo³¹P NMR spectra obtained before and 2 days after treatment with AZD3965 are shown in Fig. 5C where resonances from phosphomonoesters, Pi and α-NTP, β-NTP, and γ-NTP can be distinguished. Phosphocreatine was not observed in most cases and thus was not included in the analysis.

Our results show that compared with day 0, treatment with AZD3965 led to a significant increase in the ratio of β-NTP/total P and β-NTP/Pi up to 178 ± 48% and 198 ± 41%, respectively (P = 0.01, Fig. 5D) in Raji xenograft tumors, indicating an improved bioenergetic state. No significant differences in β-NTP/total P or β-NTP/Pi were detected in vehicle-treated tumors (112 ± 8% and 145 ± 43%, respectively, P ≥ 0.31; Fig. 5D). Further, the data revealed no significant changes in tumor pH (as indicated by the relative shift in the α-NTP-Pi resonances) in the AZD3965- or vehicle-treated tumors, indicating that tumor acidification was unaffected (Fig. 5E).

High-resolution ¹H NMR analysis of resected tumor tissue following in vivo NMR spectroscopy showed a 1.7-fold increase in lactate levels in the AZD3965-treated relative to the vehicle-treated tumors, confirming MCT1 blockade (Fig. 5F). Thus, at the dose and treatment schedule used in our in vivo study, AZD3965 caused an increase in tumor lactate accumulation concomitant with improved bioenergetic levels but no changes in tumor acidification. These data indicate that in addition to tumor lactate, bioenergetic status, as determined by ³¹P NMR measurements of the β-NTP/Pi and β-NTP/total P ratios, could have potential as an early noninvasive PD biomarker of MCT1 blockade with AZD3965 that precedes gross changes in tumor volumes.

Much attention has been focused on disrupting lactate homeostasis for cancer therapy (2–4). As such, understanding the impact of MCT1-targeted drugs on downstream metabolic activity may help identify potential combinatorial strategies as well as metabolic biomarkers for monitoring drug action, which are key for furthering the clinical development of these agents. This is particularly important because MCT1 is ubiquitously expressed (1), and inhibitor administration dose/schedule during the ongoing clinical trials needs to be carefully balanced with potential normal tissue toxicity

In this study, we focus on lymphoma models as representative of an MCT1-dependent disease (7) and a target patient population within the AZD3965 phase I expansion cohort, with the MYCN-amplified MCT4⁻ IMR-32 and MCT4⁺ Kelly neuroblastoma cell lines representing a solid tumor model expected to be MCT1-dependent (7), and MCT4⁺⁺⁺ HT29 cells as a drug-resistant model. Using steady-state metabolic profiling and flux analyses with hyperpolarized and conventional ¹³C-labeling, we show that the MCT1 inhibitor AZD3965 blocks lactate transport out of cells with consequences that affect downstream metabolism and treatment efficacy.

AZD3965 caused a marked time- and concentration-dependent accumulation in Lactate_I that was more pronounced in cells with no or low expression of MCT4, an alternative transporter of lactate that is predictive of resistance to AZD3965 (8). Using hyperpolarized [1-¹³C]pyruvate, we show that AZD3965 blocks pyruvate–lactate exchange despite the large increases in Lactate_i, which are expected to facilitate the process (25). These results are consistent with MCT1-mediated transport of [1-¹³C]pyruvate being rate limiting for the reaction (24). Our data also show that inhibition of pyruvate–lactate exchange by AZD3965 is inversely proportional to MCT4 expression and that the action of this agent is bidirectional, i.e., it inhibits transport both into and out of cells.

At concentrations that led to accumulation of Lactate_I, AZD3965 triggered several other changes in cell metabolism, notably increased TCA cycle metabolic intermediates in MCT4⁻ Raji, MCT4⁺ Hut78 and Kelly, and MCT4⁺⁺⁺ HT29 cells that translated to improved bioenergetics in Raji, Hut78, and HT29 cells. Raji and HT29 cells displayed nonuniform effects on alanine, acetate, glutamate, NADH + NAD⁺, formate, and phosphocholine. Whether such differences are related to MCT4 expression remains to be established.

The AZD3965-induced improvement in bioenergetics is of interest since a study carried out with the alternative MCT1 inhibitor SR13800 in Raji cells reported decreased bioenergetic levels and TCA cycle intermediates concomitant with Lactate_I accumulation (7). In contrast, knockout of MCT1 and MCT4 or their molecular chaperone BASIGIN resulted in an increase in Lactate_I pool but no change in ATP levels (26). Similarly, despite the increase in TCA metabolism in Kelly cells in our study, we did not observe a significant increase in ATP+ADP levels. Thus, the effect of MCT1 inhibition on bioenergetics appears to be context- and drug chemotype–dependent.

Next, and to investigate the metabolic basis underlying the AZD3965-induced changes in bioenergetics, we resorted to [1-¹³C]glucose metabolic flux analysis. Through ¹³C NMR analysis of growth media, we found that AZD3965 reduced ¹³C-glucose uptake and ¹³C-lactate production (further confirming Lactate_I build-up is due to transport blockade, reducing export rather than increased uptake). These effects were concomitant with decreased cellular ¹³C-glucose utilization possibly reflecting negative feedback regulation following the rapid and substantial build-up in cellular lactate. Importantly, we also observed a rise in cellular [4-¹³C]glutamate, indicating increased oxidative PDH flux. Levels of [2-¹³C]glutamate and [3-¹³C]glutamate, reflective of PC flux, remained unchanged; however, taking into account the fall in ¹³C-glucose incorporated into the glycolytic pathway, the data suggest that PC flux is relatively increased to maintain the labeling in [2-¹³C]glutamate and [3-¹³C]glutamate at a similar level to the control cells. Thus, AZD3965 stimulates oxidative PDH flux and, concomitantly, a relative increase in anaplerotic PC flux. The flux of pyruvate to alanine appears to be unchanged given the overall fall in ¹³C label incorporation into the glycolytic pathway, indicating that MCT1 inhibition leads to preferential routing of pyruvate toward mitochondrial metabolism. The upregulated PDH flux is consistent with enhanced oxidative phosphorylation leading to increased bioenergy-related metabolite levels observed with AZD3969 and is in agreement with a previous report showing increased oxygen consumption rates following MCT1 inhibition with AZD3965 (16). The significance of the increased PC flux is potentially multifold as the enzyme can participate in many biosynthetic reactions, but one plausible consequence of its activation here is provision of oxaloacetate to support the increased TCA cycle activity (27).

The enhanced mitochondrial metabolism leading to improved cancer cell bioenergetics and biomass production capacity suggests metabolic adaptation to enable improved survival and fitness (28). Indeed, chemoresistance has previously been shown to depend on increased intracellular ATP levels, which can provide metabolic support for key survival pathways (29). We found that although the mitochondrial complex I metformin had little effect on cell counts on its own, it significantly increased the potency of AZD3965. Coadministration of the two agents resulted in increased cell death, indicating that the shift to oxidative mitochondrial metabolism is required to sustain cell survival under MCT1 inhibition. These data are in agreement with recent reports showing that combining AZD3965 with metformin or its analogue phenformin is more effective for tumor targeting than single-agent therapy (7, 16). Furthermore, blockade of pyruvate mitochondrial transport with the MPC inhibitor UK5099 significantly increased the efficacy of AZD3695 and cell death when the two agents were combined. The increased cell death observed with coadministration of AZD3965 + metformin or AZD3965+UK5099 in Raji cells was concomitant with increased PARP cleavage indicative of apoptosis. Thus, curbing reactivation of mitochondrial metabolism through blockade of pyruvate entry or complex I activity provides a promising strategy for enhancing the antitumor effects of AZD3965.

Finally, using in vivo ³¹P NMR spectroscopy of Raji xenograft tumors showed that at a dose and treatment schedule that increased tumor lactate (confirming MCT1 blockade), acute exposure to AZD3965 triggered an increase in β-NTP/Pi and β-NTP/total P, indicating improved tumor bioenergetics, consistent with our in vitro observations.

Similar increases in bioenergetic levels have previously been attributed to improved blood flow and tumor reoxygenation following radiotherapy as well as chemotherapy (30–32). The AZD3965-triggered reactivation of oxidative mitochondrial metabolism observed in vitro suggests that this mechanism is likely to be a key contributor to the tumor energization observed here, although further work is required to assess the effect of the therapy on tumor perfusion.

In vivo³¹P NMR spectroscopy is clinically applicable with many studies reporting its use for evaluating human tumor metabolism (19, 33, 34). Thus, in addition to direct assessment of tumor lactate with lactate-selective ¹H NMR spectroscopy methods (which are also clinically applicable; ref. 35), the β-NTP/Pi and β-NTP/total P ratios could have potential as metabolic biomarkers for reporting on the action of AZD3965 in tumors. In this regard, and given the relatively low sensitivity of in vivo³¹P NMR spectroscopy, good quality spectra are a prerequisite for obtaining meaningful clinical data.

It is noteworthy that despite triggering an increase in tumor lactate, our data show that AZD3965 treatment had no effect on intracellular tumor pH. One possible explanation for this could be the relatively smaller increase in lactate observed in tumors compared with cells, where others previously reported cell acidification resulting from an increase in the Lactate_I pool following knockout of MCT1/MCT4 or BASIGIN (26). Other potential reasons could be the involvement of additional pH-regulatory mechanisms, such as carbonic anhydrase 9, shown to be upregulated with Lactate_I accumulation (26), although further work is necessary to test this possibility.

Of relevance to monitoring response to therapy, our data also show that AZD3965 blocks hyperpolarized ¹³C-pyruvate–lactate exchange due primarily to impaired ¹³C-pyruvate transport following MCT1 inhibition. Although our measurements were not conducted to monitor therapeutic response, but rather to study transport kinetics and further confirm the drug's mechanism of action, the evaluation of hyperpolarized ¹³C-pyruvate–lactate exchange has shown potential for tumor metabolic imaging in many preclinical studies (36), and their clinical feasibility has recently been demonstrated in prostate cancer patients (37, 38). When the technology is validated for clinical cancer imaging and becomes more widely available, monitoring hyperpolarized ¹³C-pyruvate–lactate exchange could provide a complementary biomarker of MCT1 inhibition in tumors in vivo, with ³¹P NMR measurements enabling the assessment of downstream consequences of treatment. Studies such as the one described here are thus of particular value for predicating the design of future clinical trials incorporating much-needed noninvasive biomarkers of drugs targeting cancer metabolism. Following successful completion of a clinical dose finding phase I trial, AZD3965 is now entering the expansion phase of the trial, and the data reported here underpin aspects of this trial design and the NMR spectroscopy PD biomarkers included in the protocol.

In summary, we show that MCT1 inhibition with AZD3965 impairs bidirectional monocarboxylate transport, causes build-up of Lactate_i under low/no MCT4 expression, and leads to inhibition of glycolytic activity and reactivation of mitochondrial pyruvate metabolism necessary to maintain cell survival. These findings highlight the potential to combine MCT1 inhibitors with mitochondrial-targeted therapy in the clinic and incorporate noninvasive PD magnetic resonance spectroscopy to evaluate drug action in patients.

P.D. Smith has an ownership interest (including patents) in AstraZeneca. No potential conflicts of interest were disclosed by the other authors.

Conception and design: M. Beloueche-Babari, P.D. Smith, M.O. Leach

Development of methodology: M. Beloueche-Babari, S. Wantuch, H.G. Parkes, Y.-L. Chung, T.R. Eykyn, M.O. Leach

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): M. Beloueche-Babari, S. Wantuch, T. Casals Galobart, M. Koniordou, H.G. Parkes

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): M. Beloueche-Babari, S. Wantuch, T. Casals Galobart, H.G. Parkes, T.R. Eykyn

Writing, review, and/or revision of the manuscript: M. Beloueche-Babari, H.G. Parkes, Y.-L. Chung, T.R. Eykyn, M.O. Leach

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): M. Beloueche-Babari, S. Wantuch, T. Casals Galobart, V. Arunan

Study supervision: M. Beloueche-Babari, P.D. Smith, M.O. Leach

We thank Dr. Louise Howell for help with confocal microscopy and the staff of the Biological Services Unit at ICR for animal care and maintenance.

M. Beloueche-Babari, S. Wantuch, T. Casals Galobart, M. Koniordou, H.G. Parkes, V. Arunan, Y.-L. Chung, T.R. Eykyn, and M.O. Leach are supported by a CRUK Centre for Cancer Imaging grant C1090/A16464. We also acknowledge grant C1060/A10334 from CRUK and EPSRC Cancer Imaging Centre in association with the MRC and Department of Health (England), NHS funding to the NIHR Biomedical Research Centre, and the Clinical Research Facility in Imaging. M.O. Leach is a NIHR Emeritus Senior Investigator.

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