Endothelial cells (ECs) require glycolysis for proliferation and migration during angiogenesis; however, the necessity for the mitochondrial respiratory chain during angiogenesis is not known. Here we report that inhibition of respiratory chain complex III impairs proliferation, but not migration, of ECs in vitro by decreasing the NAD+/NADH ratio. To determine whether mitochondrial respiration is necessary for angiogenesis in vivo, we conditionally ablate a subunit of the respiratory chain complex III (QPC) in ECs. Loss of QPC decreases respiration, resulting in diminished EC proliferation, and impairment in retinal and tumour angiogenesis. Loss of QPC does not decrease genes associated with anabolism or nucleotide levels in ECs but diminishes amino acid levels. Our findings indicate that mitochondrial respiration is necessary for angiogenesis and that the primary role of mitochondria in ECs is to serve as biosynthetic organelles for cell proliferation.
While largely quiescent in healthy adult tissues, ECs are dynamic, undergoing rapid migration and proliferation during angiogenesis, thus increasing their biosynthetic and bioenergetic demand. Recent findings indicate that metabolism is a critical regulator of EC function during angiogenesis. ECs are classically described as being highly glycolytic, and numerous studies have illustrated the necessity of glycolysis in ECs during vessel sprouting1. Despite having abundant access to oxygen in the bloodstream, ECs generate up to 85% of their ATP from glycolysis2. When vessel sprouting is stimulated by pro-angiogenic signals, such as vascular endothelial growth factor A, glycolysis is enhanced owing to upregulation of glycolytic enzymes, nearly doubling flux through glycolysis2,3,4. This increased glycolytic flux is critical, as decreased expression of the glycolytic activator phosphofructokinase-2/fructose-2,6-bisphosphatase (PFKFB3), severely impairs both migration and proliferation of ECs in vitro and in vivo2. Furthermore, inhibition of PFKFB3 in ECs decreased pathological neovascularization in ocular and inflammatory models, as well as cancer cell metastasis5,6. Moreover, decreased levels of other glycolytic enzymes, including hexokinase 2, pyruvate kinase M2, and glucose transporter 1, cause impaired angiogenesis7,8,9. Collectively these data indicate that glycolytic flux in ECs is necessary for angiogenesis.
Since ECs undergo high levels of aerobic glycolysis and thus oxidize only small amounts of glycolysis-derived pyruvate within the mitochondria, mitochondrial function was previously surmised to play a marginal role in EC metabolism2. In addition to importing glucose, ECs take up fatty acids from their environment and break them down into acetyl-CoA in the mitochondria in a process known as fatty acid oxidation. Fatty acid–derived acetyl-CoA is then utilized to fuel the tricarboxylic acid (TCA) cycle10. Fatty acid oxidation has been shown to be critical in ECs, as knockdown of fatty acid binding protein 4 or carnitine palmitoyltransferase 1A leads to diminished angiogenesis11,12,13. Glutamine is also readily imported into ECs where it is metabolized into alpha-ketoglutarate (α-KG), which is used to replenish the mitochondrial TCA cycle (namely, anaplerosis). Limiting EC glutamine utilization results in diminished angiogenesis both in vitro and in vivo14,15. While fatty acid oxidation and glutamine utilization fuel metabolism during vessel sprouting, loss of either of these two pathways results in a relatively modest decrease in angiogenesis compared with inhibition of PFKFB3. It is likely that restricting one particular carbon source to fuel the TCA cycle will not markedly impair mitochondrial metabolism.
In the present study, we directly tested the necessity for mitochondrial metabolism during angiogenesis by inhibiting the mitochondrial respiratory chain complex III. Respiration is initiated by donation of electrons to complexes I and II by the TCA cycle–generated reducing equivalents NADH and FADH2, respectively. Mitochondrial complexes I and II pass electrons to complex III and finally to oxygen via complex IV. This electron flow drives the pumping of hydrogen ions into the intermembrane space to generate the membrane potential required for ATP production by ATP synthase (complex V). Respiration is linked to three distinct mitochondrial functions: (1) oxidative phosphorylation for ATP generation; (2) oxidative TCA cycle flux to produce metabolites for macromolecule biosynthesis; and (3) release of reactive oxygen species and metabolites to determine cell fate and/or function. Thus, loss of mitochondrial complex III could have multiple effects on cells, including death due to a bioenergetic collapse, decreased proliferation due to diminished oxidative TCA cycle function for macromolecule synthesis, and loss of cell fate and/or function due to changes in angiogenic gene expression. Thus, we utilized mitochondrial complex III inhibitors in vitro, and an EC-specific mitochondrial complex III-deficient mouse model to test which mechanisms linked to mitochondrial respiration in ECs are required for angiogenesis.
Here, we conclude that in vitro mitochondrial complex III is necessary for EC proliferation but not migration, and is critical for maintenance of the NAD+/NADH ratio. Conditional loss of complex III (QPC) in ECs in vivo diminishes retinal, lung, and tumour angiogenesis by decreasing EC proliferation. While anabolic gene signalling is maintained, loss of QPC in ECs in vivo results in reduced amino acid levels. Overall, we conclude that mitochondrial complex III is required for angiogenesis, and in ECs the principal role of mitochondria is to act as signalling organelles to support proliferation.
Mitochondrial complex III is required for endothelial cell proliferation in vitro
To study the effects of mitochondrial respiratory chain inhibition on EC behaviour, we cultured human umbilical vein endothelial cells (HUVECs) with the mitochondrial complex III inhibitor antimycin A to induce loss of mitochondrial respiration (Fig. 1a). Since respiratory capacity was abolished, we confirmed that HUVECs were performing maximal glycolysis basally (Fig. 1b). Importantly, even after 96 h of treatment with antimycin A in culture, no significant decrease in cell viability was observed, indicating that respiratory chain inhibition does not cause HUVEC death or apoptosis (Fig. 1c and Supplementary Fig. 1a,b). Furthermore, antimycin A treatment does not lead to mitochondrial demise in vitro, as we observed no loss of mitochondrial membrane potential, mitochondrial content, or mitochondrial DNA (Supplementary Fig. 1c–i). Two important processes activated in ECs in response to angiogenic signalling are sprouting (that is, invasion and migration) and proliferation16. We therefore utilized HUVECs in culture to explore how mitochondrial inhibition affects these key angiogenic functions. First, we used an in vitro sprouting assay where HUVEC spheroids sprout and migrate through a three-dimensional collagen matrix. After 24 h, spheroids treated with antimycin A showed no changes in sprout length or number (Fig. 1d–f). Additionally, respiration-deficient HUVECs migrated similarly to controls in a two-dimensional scratch-wound migration assay (Fig. 1g). Interestingly, HUVECs treated with antimycin A displayed a striking defect in proliferation, as they completely failed to grow over a period of 96 h (Fig. 1h). Similar results were obtained when HUVECs were treated with the mitochondrial complex I inhibitor piericidin (Supplementary Fig. 2). Together, these data clearly show that the mitochondrial respiratory chain is necessary for HUVEC proliferation in vitro, while it is dispensable for migration and sprouting.
Mitochondrial complex III maintains NAD+/NADH ratio, necessary for EC proliferation
The mitochondrial TCA cycle is critical for cellular proliferation, as it produces metabolic intermediates that are used as building blocks for biosynthetic macromolecules17. Metabolic profiling revealed that respiration-deficient HUVECs displayed decreased levels of TCA cycle intermediates (Fig. 2a). Broadly, amino acid levels in HUVECs treated with antimycin A were not impaired, with the exception of aspartate, which was significantly decreased (Fig. 2b). Nucleotide metabolite abundance remained relatively unchanged in respiration-deficient HUVECs (Supplementary Fig. 3a,b). To test specificity of antimycin A, we expressed the Ciona intestinalis alternative oxidase (AOX) in HUVECs, which is refractory to antimycin treatment18,19. AOX has the capability to accept electrons from ubiquinol and transfer them to oxygen, bypassing complex III and IV functions, and restoring electron transport chain activity in the presence of complex III inhibition18,19. We found that HUVECs expressing AOX maintain oxygen consumption and NAD+/NADH in the presence of antimycin A, indicating that AOX is functioning properly (Fig. 2c,d). Importantly, AOX was able to rescue proliferative capacity after antimycin A treatment, as it restores both TCA cycle metabolites and aspartate levels (Fig. 2e–g). Together, these data highlight the specificity of antimycin A and demonstrate that its effects on HUVECs are due to its inhibition of complex III.
Cells use complex I of the electron transport chain to oxidize NADH to NAD+; thus, after complex III inhibition, there is a decreased capacity to regenerate NAD+, causing decreased TCA cycle flux. Previously, it has been demonstrated that restoration of NAD+/NADH is sufficient to rescue proliferation in respiration-deficient cancer cells20,21. To test this hypothesis in HUVECs, we expressed an NADH oxidase from Lactobacillus brevis (LbNOX), which generates NAD+ from NADH independent of electron transport chain function21. We found that expressing LbNOX in the cytosol did not restore oxygen consumption rate (OCR) but was able to rescue NAD+/NADH, and proliferation in antimycin A-treated HUVECs (Fig. 2h–j). Additionally, respiration-deficient HUVECs expressing LbNOX accumulate large amounts of succinate, due to complex III inhibition (Fig. 2k). Finally, expressing LbNOX in antimycin A-treated HUVECs was able to increase aspartate levels relative to empty vector (Fig. 2l). These data indicate that an essential function of the electron transport chain in HUVECs is to maintain NAD+/NADH to support cellular proliferation.
Numerous studies have reported that cells lacking a functional respiratory chain are able to proliferate in vitro on supplementation with supra-physiological levels of pyruvate in the culture medium20,22,23. Similarly, we observed that addition of methyl pyruvate, while not rescuing OCR, was able to restore proliferative capacity in antimycin A–treated HUVECs (Supplementary Fig. 3c,d). Additionally, antimycin A–treated HUVECs with methyl pyruvate accumulated succinate owing to complex III inhibition and displayed restored aspartate levels (Supplementary Fig. 3e,f). As aspartate levels are decreased in antimycin A–treated HUVECs and are rescued on expression of AOX, LbNOX, or addition of methyl pyruvate, we hypothesized that supplementation with aspartate would allow proliferation after electron transport chain inhibition. Previous studies have reported that in respiration-deficient cancer cells, aspartate is sufficient to support proliferation in the absence of pyruvate20,22. Surprisingly, aspartate was not able to rescue proliferation in antimycin A–treated HUVECs (Supplementary Fig. 3g). Additionally, neither cell-permeable methyl aspartate nor asparagine (which is converted to aspartate by the enzyme asparaginase) were able to restore proliferation (Supplementary Fig. 3h,i). Together, these data reveal that in HUVECs, mitochondrial complex III fulfils biosynthetic requirements by supporting NAD+ regeneration. It is likely that NAD+-dependent aspartate synthesis is necessary but not sufficient for cell proliferation.
Inhibiting mitochondrial complex III respiration in HUVECs does not broadly disrupt histone modifications
In addition to fulfilling a biosynthetic demand, mitochondrial metabolites have recently been shown to be necessary to maintain histone acetylation24,25. Accumulation of the TCA cycle intermediates succinate and fumarate as well as the metabolite l-2-hydroxyglutarate (2HG), leads to competitive inhibition of α-KG-dependent dioxygenases, including JmJC domain-containing histone lysine demethylases and the ten-eleven translocation (TET) family of 5-methlycytosine (5mC) hydroxylases26,27. Recently, we reported that build-up of these three metabolites in mitochondrial complex III knockout (KO) haematopoietic stem cells resulted in hypermethylation of DNA and histones24. In the present study, we unexpectedly observed a decrease in 2HG as well as the expected increase in 2HG:α-KG, succinate:α-KG, and fumarate:α-KG (Supplementary Fig. 3j–m). Others have reported that fatty acid oxidation is necessary to maintain H3K9 acetylation in lymphatic endothelial cells28. However, we found that histone modifications in HUVECs treated with antimycin A were largely unchanged (Supplementary Fig. 4a). Specifically, we did not observe changes in H3K9 acetylation or methylation and saw minimal alterations in other major histone marks linked to gene expression (Supplementary Fig. 4b–f). Together, these data indicate that mitochondrial complex III in HUVECs in vitro is not required for maintenance of chromatin modifications.
Mitochondrial complex III respiration in ECs is required for postnatal retinal angiogenesis
The microenvironment and nutrient availability of ECs differs greatly between cell culture and a living organism, prompting us to ask what the requirements are for EC mitochondrial complex III function in vivo. To explore the effect of mitochondrial respiratory chain inhibition in vivo, we crossed mice with a floxed Uqcrq gene, which encodes for ubiquinol-binding protein (a critical subunit of the mitochondrial respiratory chain complex III) (QPCfl/fl), with Cdh5CreERT2 mice, to allow for tamoxifen-inducible, endothelial-specific loss of mitochondrial respiration (QPC-KO)29.
To understand the role of EC mitochondrial complex III respiration in vessel sprouting in vivo, we studied angiogenesis in the postnatal mouse retina. Mouse pups were administered five doses of tamoxifen on postnatal days 0–4 (P0–P4) to induce Cre recombination and loss of respiration in ECs. To ensure recombination, QPC-WT (wild type) and QPC-KO mice were crossed to mice harbouring a lox-stop-lox TdTomato allele (Supplementary Fig. 5). Retinal whole mounts from P7 pups stained with isolectin-B4 (IB4) revealed a dramatic impairment in retinal angiogenesis in QPC-KO mice (Fig. 3a). By P7, vessels should nearly reach the outer edge of the retina; however, radial expansion was massively impaired in QPC-KO mice, as vessels only reached about half the distance to the outer retinal edge (Fig. 3a,b). Additionally, a striking decrease in vascular density could be observed in QPC-KO retinas, which showed substantially fewer branchpoints per mm2 (Fig. 3a,c). Further staining revealed that QPC-KO retinas had significantly fewer phosphohistone 3 (pH3)-positive ECs, indicating a proliferative impairment congruent with that observed in vitro (Figs. 1h and 3d,e). Moreover, filopodia of QPC-KO ECs on the outer retinal edge were unremarkable in both appearance and number compared with QPC-WT, suggesting no defect in migration, which is again consistent with our in vitro data (Figs. 1d–1g and 3f,g). Additionally, we did not observe an increase in retinal EC apoptosis (cleaved caspase 3) or vessel regression (empty collagen IV sleeves) in QPC-KO mice compared with WT, indicating that decreased retinal vascularity was not due to EC death or regression (Supplementary Fig. 5b–e). We conclude that complex III inhibition in vivo decreases EC proliferation but does not alter migration, apoptosis, or regression of vessels during postnatal retinal angiogenesis.
Mitochondrial complex III function in ECs is necessary for postnatal developmental angiogenesis
Next, we aimed to uncover whether mitochondrial respiration in ECs is necessary for postnatal pup survival. To answer this question, QPC-WT and QPC-KO pups were given five doses of tamoxifen (P0–P4) to induce Cre recombination and loss of QPC in ECs. We observed a striking decrease in survival of QPC-KO pups, the majority of which died between P15 and P30 (Fig. 4a). To further explore vascular defects in this model, lungs were collected from P15 QPC-WT and QPC-KO pups. We chose to investigate the lung as it contains a large population of ECs, which are actively undergoing angiogenesis at this age, a critical process in postnatal lung alveolarization30. QPC-KO lung ECs had diminished expression of Uqcrb (Qpc) messenger RNA (mRNA) and displayed a decreased OCR and NAD+/NADH ratio, indicating a loss of respiratory chain function (Supplementary Fig. 6a–c). QPC-KO pups showed no obvious signs of distress or decline in body weight at P15 (Fig. 4b). However, flow cytometric analysis of homogenized P15 lungs revealed a significant reduction in both the percentage and total number of ECs, consistent with a decrease in lung angiogenesis (Fig. 4c,d and Supplementary Fig. 6d). Recall that in vitro, as well as in the retina, mitochondrial respiratory chain complex III inhibition caused reduced proliferation (Figs. 1h and 3d,e). Likewise, pH3 staining of P15 lungs revealed a decrease in EC proliferation (Fig. 4e,f). Taken together, these data show that loss of mitochondrial complex III leads to a significant defect in lung angiogenesis.
Loss of mitochondrial complex III function in ECs increases anabolic-associated gene expression
Although we did not observe profound changes in histone methylation or acetylation in respiration-deficient HUVECs (Supplementary Fig. 4), we wondered whether loss of mitochondrial respiration in ECs would lead to gene deregulation in vivo. Thus, we collected lung ECs from QPC-WT and QPC-KO P15 pups treated with tamoxifen (P0–P4) and performed RNA-sequencing (RNA-seq) analysis. We observed modest alterations in gene expression, with 237 upregulated and 142 downregulated genes in QPC-KO ECs compared with QPC-WT (false discovery rate (FDR) ≤ 0.01) (Fig. 5a). Surprisingly, gene set enrichment analysis (GSEA) revealed a significant upregulation in pathways associated with anabolism and cellular proliferation, including MYC, MTORC, E2F, and G2M target signalling (Fig. 5b). We also observed an increase in the unfolded protein response, a cellular stress response that has been shown to be activated after loss of mitochondrial respiratory function31 (Fig. 5b). Ribosomal biosynthesis genes are critical targets of cellular pro-growth signalling32. Indeed, we found that a substantial percentage (21%) of the significantly upregulated genes in QPC-KO ECs were ribosomal genes, further corroborating intact proliferative signalling (Fig. 5c). QPC-KO ECs show several significantly increased metabolic genes, including those regulating glycolysis, one-carbon metabolism, and the urea and TCA cycles (Fig. 5d). Additionally, QPC-KO ECs display a trend towards increased oxidative phosphorylation genes, probably a compensatory mechanism due to massive loss of mitochondrial function (Fig. 5e). These data indicate that although QPC-KO ECs have decreased proliferation, they maintain anabolic gene expression.
Interestingly, RNA-seq revealed a handful of key angiogenic signalling genes that were significantly altered in QPC-KO ECs (Fig. 5f). Specifically, hairy/enhancer-of-split related with YRPW motif protein 1 (Hey1), delta-like 4 (Dll4), and TEK receptor tyrosine kinase (Tek, also called Tie2) were significantly downregulated, while neuropilin-2 (Nrp2), and angiopoitin-2 (Angpt2) were significantly upregulated in QPC-KO ECs (Fig. 5f). Validation of our data at the protein level revealed increased metabolic protein expression but no change in angiogenic protein expression (Supplementary Fig. 7). Our data suggest that ECs lacking mitochondrial complex III have impaired proliferation and angiogenesis yet retain anabolic gene expression.
Mitochondrial complex III in ECs is necessary to maintain amino acid levels in vivo
As anabolic signalling remained intact in QPC-KO ECs, we hypothesized that a metabolic deficiency was preventing proliferation. Metabolite analysis revealed decreased levels of numerous metabolites in the QPC-KO ECs compared with QPC-WT (Fig. 6a). We did not observe an accumulation of 2HG; however, the TCA cycle metabolites fumarate and malate were significantly lower in QPC-KO ECs, as were several glycolytic intermediates (Fig. 6b and Supplementary Fig. 8). Intriguingly, the majority of the metabolites decreased in QPC-KO ECs were amino acids (Fig. 6a). In contrast to HUVECs in vitro, where amino acid levels were maintained with the exception of aspartate, QPC-KO lung ECs displayed significantly diminished levels of nearly all amino acids (Fig. 6a,c). Purine and pyrimidine nucleotide levels, however, remained largely unchanged after loss of EC respiration (Fig. 6d,e). Together, these results suggest that mitochondrial complex III in ECs is required to maintain amino acid levels, but not nucleotides, which could lead to the impaired proliferation observed in QPC-KO ECs.
Mitochondrial complex III function in ECs is required for tumour angiogenesis
To further investigate the role of EC respiration and determine its requirement in adult mice, we asked whether mitochondrial respiration in ECs is required for tumour angiogenesis. Eight-week-old adult QPC-WT and QPC-KO mice were fed tamoxifen chow for 2 weeks to induce Cre recombination and loss of Qpc mRNA expression and respiration in ECs (Fig. 7a,b). Next, we injected syngeneic B16-F10 melanoma cells subcutaneously into the mice and measured tumour volume over the course of 21 d. QPC-KO mice showed a decrease in tumour growth compared with QPC-WT, and by days 19 and 21, they harboured significantly smaller tumours (Fig. 7c). On day 21, when tumours were collected, QPC-KO mice also displayed a reduction in tumour weight (Fig. 7d). Histological analysis revealed that QPC-KO tumours had fewer vessels per area, indicating decreased tumour angiogenesis (Fig. 7e,f). Consistent with data from the retina and lung, tumour ECs proliferated less, suggesting that impaired tumour angiogenesis is probably due to reduced EC proliferation (Fig. 7g,h). These data suggest that mitochondrial complex III function is required in adult ECs to sustain tumour angiogenesis and tumour growth.
Endothelial cells exhibit high levels of flux through glycolysis, only oxidizing a small fraction of glucose-derived carbons in the mitochondria2,15. Accordingly, limiting glycolysis profoundly impairs angiogenesis2,5,6,7,8,9. By contrast, the function of mitochondrial metabolism in ECs is not fully understood. Here we report that pharmacological inhibition of mitochondrial respiratory transport chain complex III in ECs impairs cell proliferation by decreasing NAD+/NADH in vitro. Conditional loss of respiratory chain complex III in ECs in vivo diminished postnatal retinal, lung, and tumour angiogenesis. Our results conclusively demonstrate that mitochondrial respiratory chain complex III is necessary for angiogenesis by controlling EC cell proliferation. These results fill a critical gap in knowledge about the role of the mitochondrial respiratory chain in ECs, a classically glycolytic cell type in which mitochondrial metabolism has largely been underappreciated. Our results indicate that angiogenesis requires coordination of both glycolysis and mitochondrial respiratory chain–linked metabolism. It is likely that this coordination in ECs occurs through MYC, since loss of MYC specifically in ECs impairs glycolysis, mitochondrial metabolism, and proliferation33.
Mitochondria serve three main functions within a cell: (1) they generate ATP via oxidative phosphorylation for cell survival; (2) the TCA cycle generates metabolic intermediates that produce critical macromolecules required for cell growth, including amino acids, nucleotides, and lipids; and (3) mitochondria act as signalling organelles, generating reactive oxygen species to activate transcriptional networks and produce metabolic intermediates that control epigenetics. The major phenotype we observed due to mitochondrial complex III function impairment was diminished cell proliferation in vitro and in vivo. This defect is probably not due to lack of mitochondrial ATP production, as it has been suggested that ECs generate up to 85% of their ATP through glycolysis alone2. Additionally, complex III loss in ECs did not impair ATP-demanding activities such as sprouting and migration in vitro or filopodia formation in vivo, further highlighting that glycolysis can alone provide sufficient ATP for these processes. Our data are consistent with previous studies showing that glycolysis drives filopodia formation and migration in ECs2.
The proliferative impairment observed in complex III–deficient ECs in vitro and in vivo is probably due to the inability to generate the necessary metabolites for macromolecule synthesis. Previous studies in cancer cells in vitro have demonstrated that loss of complex III decreases NAD+ levels, resulting in diminished aspartate, which is necessary for cancer cell proliferation20,22. In cancer cells treated with complex III inhibitors, restoration of NAD+ levels, either by supplementation with pyruvate or by genetic expression of NAD+-regenerating enzymes, restores aspartate levels and cell proliferation in vitro20,21. Congruently, in vitro, we observed decreased NAD+/NADH and diminished aspartate levels in respiration-deficient HUVECs. As previous studies have shown, supplementation with pyruvate restored proliferation after electron transport chain inhibition as well as aspartate levels. Additionally, we found that genetically increasing the ratio of NAD+/NADH was sufficient to rescue proliferation and aspartate in antimycin A–treated HUVECs, which is again consistent with in vitro cancer cell data21. Intriguingly, unlike cancer cells, aspartate was not able to restore proliferation in respiration-deficient ECs in vitro, perhaps due to distinct metabolic programming between cancer and primary cells. It is important to note that proliferation of not all cancer cell lines is sensitive to aspartate limitation34. Additionally, cancer cells differ in their ability to uptake aspartate, and have varied asparaginase activity35. We suggest that perhaps aspartate is not sufficient to support proliferation in HUVECs. However, NAD+/NADH ratio, which is linked to other metabolic functions beyond restoring aspartate levels, is sufficient to support HUVEC proliferation.
Additionally, we found that ECs in vivo lacking complex III function not only had diminished NAD+/NADH and aspartate levels, but also decreased abundance of the majority of amino acids. It is not clear why amino acid levels were markedly diminished from complex III-deficient ECs. GSEA revealed an unexpected increase in anabolic gene expression profiles in QPC-KO ECs, with an increase in genes linked to MYC and mTORC1, including upregulated ribosomal gene expression. This increase in anabolic programmes would impose a high demand for amino acids. Thus, in the absence of a functional respiratory chain, the TCA cycle metabolites are not able to maintain amino acid levels to keep up with the increased anabolic demand, resulting in diminished growth. Previously, we observed that GSEA from mitochondrial complex III–deficient haematopoietic stem cells (HSCs) similarly displayed upregulation of MYC- and mTORC1-linked genes24. Going forward it will be important to decipher how loss of mitochondrial complex III function causes an increase in MYC- and mTORC1-related genes in vivo.
Although our present data on ECs indicate that respiratory chain–linked metabolism is necessary for cell proliferation, it is not a universal feature of proliferating cells. For example, foetal mouse HSCs do not require mitochondrial complex III for cell proliferation but to generate sufficient progenitor populations in vivo24. Moreover, adult mouse complex III–deficient HSCs lose quiescence and undergo stem cell exhaustion; thus, complex III is required for HSCs to function properly24. Mitochondrial complex III–deficient HSCs display deregulated expression of approximately 1,000 genes, concomitant with histone H3K4, H3K9, and H3K79 hypermethylation24. These changes were accompanied by increased levels of succinate, fumarate, and 2HG, metabolites known to inhibit α-KG-dependent dioxygenases, including KDMs and TETs24,26,27. We hypothesized that we would see these same trends in ECs, however we did not observe accumulation of neither succinate, fumarate, nor 2HG in vivo.
Although, the mitochondrial complex III–deficient ECs did not display widespread deregulation of gene expression, there were a few angiogenic associated genes altered, including those involved in the Notch signalling pathway (Dll4 and the two downstream targets Hey1 and Hes1) in vivo. Previous studies have shown that Notch signalling is critical for angiogenesis both in vitro and in vivo2,36. Specifically, decreased Notch signalling in HUVECs accelerated sprouting, while constitutive Notch activation had the opposite effect2. In a mouse model, pharmacological inhibition of Notch signalling with the inhibitor DAPT or genetic ablation through conditional knockout of the Notch ligand Dll4 in ECs increased branching and tip cell formation in the retina36. In our in vivo model, loss of complex III in ECs lead to decreased Notch signalling; however, we observed decreased branching, contrary to what has been observed after loss of Notch signalling in ECs. As loss of Notch signalling has been found to have the opposite phenotype to diminished respiration in ECs, we conclude that decreased Notch signalling is probably not the dominant factor that leads to impaired angiogenesis in our model. Additionally, mitochondrial complex III–deficient ECs display decreased mRNA expression of Tie2, along with overexpression of the Tie2-negative regulator Angpt2, suggesting decreased signalling through Tie2 in QPC-KO ECs37,]38. However, we observed no concomitant change in Tie2 or Angpt2 protein levels, suggesting no alterations in Tie2 signalling. Overall, we conclude that mitochondrial complex III’s dominant function in ECs is to sustain amino acid availability for cell proliferation in vivo.
Our observation that complex III–linked metabolism in ECs is necessary for cell proliferation is similar to observations in cancer cells. Previously, both ourselves and others have demonstrated that pharmacological inhibition or genetic ablation of the respiratory chain within cancer cells diminishes tumourigenesis in part by decreasing cell proliferation39,40,41,42,43,44,45,46,47,48. This has led to the idea of targeting the respiratory chain for cancer therapy49. We observed that inhibition of the respiratory chain, by diminishing complex III in ECs, impaired tumourigenesis. Consequently, administration of respiratory chain inhibitors could work as an anticancer therapy through decreasing proliferation of both cancer and endothelial cells. Interestingly, increasing mitochondrial function diminishes prostate tumour vascularization, highlighting that mitochondrial homeostasis is crucial for maintaining tumour angiogenesis50.
Cell culture and drug treatment
HUVECs (Lonza) were cultured in endothelial basal medium (MCDB 131) without pyruvate (USBiological) and used at a low passage number (1–6) for in vitro assays. Medium was supplemented with EGM-2 SingleQuot growth factors (Lonza), 1% GlutaMax (Gibco), and 400 µM uridine (supplemented MCDB 131 media). B16-F10 melanoma cells were cultured in RPMI (Corning), with 10% FBS (Corning), 1% sodium pyruvate (Gibco), 1% non-essential amino acids (Gibco), 1% GlutaMAX (Gibco), 1% antibiotic/antimycotic (Corning), and 0.05 mM β-mercaptoethanol (Sigma). Cells were maintained at 37 °C with 5% CO2. HUVECs were treated with: 25 nM antimycin A (Sigma), 250 nM piericidin (Sigma), 1 mM methyl pyruvate (Sigma), and/or indicated doses of l-aspartic acid (aspartate) (Sigma), l-aspartic acid dimethyl ester hydrochloride (methyl aspartate) (Sigma), or l-asparagine (asparagine) (Sigma).
Oxygen consumption and extracellular acidification rate measurements
OCR and ECAR were measured using an XF96 extracellular flux analyser (Seahorse Bioscience). In vitro: 30,000 HUVECs per well were plated onto XF96 cell culture plates in supplemented MCDB 131 medium and allowed to attach for 4 h. Cells were then treated with mitochondrial respiratory chain inhibitor for 2 h. Basal respiration was measured by subtracting the OCR values after treatment with 2 μM antimycin A (Sigma) and 2 μM rotenone (Sigma). Coupled respiration was determined by treatment with 2.5 μM oligomycin A (Sigma) and subtracting oligomycin A values from basal respiration. To measure ECAR, cells were treated in supplemented MCDB 131 medium without sodium bicarbonate. Basal ECAR was measured by subtracting the ECAR rate after treatment with 20 mM 2-deoxyglucose (Sigma). Maximum ECAR rate was measured by subtracting the rate after after 2-deoxyglucose treatment from the rate after treatment with 2.5 μM oligomycin A.
In vivo: 100,000 lung ECs were plated onto XF96 cell culture plates in supplemented MCDB 131 medium and attached using Cell-Tak following the manufacturer’s protocol (Corning). Basal respiration was measured by subtracting the OCR values after treatment with 2 μM antimycin A (Sigma) and 2 μM rotenone (Sigma). Coupled respiration was determined by treatment with 2.5 μM oligomycin A (Sigma) and subtracting oligomycin A values from basal respiration.
HUVEC proliferation and viability assays
For proliferation curves, HUVECs were plated in supplemented MCDB 131 medium and allowed to attach overnight. The next day (time 0), medium was replaced with supplemented MCDB 131 medium with or without drug treatments. Cells were then counted at each time point (24, 48, 72, or 96 h) via flow cytometry using AccuCount Fluorescent Particles (Spherotech). For viability, HUVECs were plated and treated as described above. Viability was assessed at 96 h post-treatment via flow cytometry by adding 100 ng ml−1 of DAPI and assessing the percentage of DAPI-negative cells. To measure apoptosis and viability, HUVECs were plated and treated as described above. After 48 or 96 h, cells were collected and stained with AnnexinV and propidium iodide according to the manufacturer’s protocol (eBioscience AnnexinV apoptosis detection kit).
Measurement of HUVEC mitochondrial membrane potential and content
HUVECs were plated and treated as described above and collected after 48 or 96 h. To measure membrane potential, cells were stained with 50 nM TMRE (tetramethylrhodamine ethyl ester perchlorate, Molecular Probes) for 20 min at 37 °C. Median fluorescence intensity (PE) was measured by flow cytometry and corrected by subtracting the median fluorescence intensity of each sample after addition of 25 μM carbonyl cyanide m-chlorophenyl hydrazine. To measure mitochondrial content, cells were stained with 25 nM MitoTracker Green (Molecular Probes) for 20 min at 37 °C. Median fluorescence intensity (using fluorescein isothiocyanate) was measured by flow cytometry.
qRT–PCR and western blot analysis
To measure QPC expression, RNA was isolated using the E.Z.N.A Total RNA Kit I by following the manufacturer’s protocol (Omega). CYBRFast 1-Step RT–qPCR Lo-ROX Kit (Tonbo) was used to measure QPC expression with the following primers: QPC-F: 5′-GAGACTGAGGATATCGATTG-3′, and QPC-R: 5′-GGATGCGCTCGCGAGTGCGG-3′. To measure mitochondrial DNA content, genomic DNA was purified using the QIAamp DNA Mini Kit (Qiagen), and qRT–PCR was performed using iQ SYBR Green Supermix (Bio-Rad) using the following primers: ND2-F: 5′-GCCCTAGAAATAAACATGCTA-3′; ND2-R: 5′-GGGCTATTCCTAGTTTTATT-3′; COX2-F: 5′-CTGAACCTACGAGTACACCG-3′; COX2-R: 5′-TTAATTCTAGGACGATGGGC-3′; CYTB-F: 5′-CATTTATTATCGCGGCCCTA-3′; CYTB-R: 5′-TGGGTTGTTTGATCCTGTTTC-3′; SDHA-F: 5′-TCCACTACATGACGGAGCAG-3′; SDHA-R: 5′-CCATCTTCAGTTCTGCTAAACG-3′; β-actin-F: 5′-TCCACCTTCCAGCAGATGTG-3′; β-actin-R: 5′-GCATTTGCGGTGGACGAT-3′. For western blot analysis, cells were lysed in 1× cell lysis buffer (Cell Signaling) containing PMSF. Relative protein abundance was measured through the Wes system (ProteinSimple) using the manufacturer’s protocol and the following antibodies: SHMT2 (Cell Signaling, 12762), PHGDH (Abcam, Ab211365), GAPDH (Sigma, G9545), Tie2/TEK (Millipore, 05-584), and Angpt2 (Invitrogen, PA5-23612).
HUVEC sprouting assay
HUVECs were first resuspended in 20% methylcellulose (Sigma, 4 Pa. s viscosity) at a concentration of 1,000 cells per 25 μl. This suspension was then dropped onto non-adherent plates (25-μl drops) and flipped upside-down, and HUVECs were allowed to form spheroids overnight in hanging drops. Spheroids were collected in PBS + 10% FBS and spun down at 300g for 5 min and then 500g for 3 min at room temperature with no brake. Spheroids were then resuspended in 40% FBS in methylcellulose, combined with collagen type I (EMD Millipore), NaHCO3, and NaOH, plated, and allowed to polymerize at 37 °C. Wells were then topped with an equal volume of supplemented MCDB 131 medium with or without drug treatments. Medium contained 2 μg ml−1 mitomycin C (Sigma) to inhibit proliferation. Spheroids were allowed to sprout for 24 h before fixation with 4% paraformaldehyde.
HUVEC scratch-wound cell migration assay
Scratch-wound cell migration assays were performed using the IncuCyte ZOOM 96-well scratch-wounds cell migration system (Essen BioScience). HUVECs were plated onto 96-well Image Lock tissue culture plates (Essen BioScience, 4379) at a density of 30,000 cells per well in supplemented MCDB 131 medium and allowed to attach overnight. Cells were then treated with 5 μg ml−1 mitomycin C for 2 h to inhibit proliferation. Monolayers were then wounded using the 96-well WoundMaker (Essen BioScience) following the manufacturer’s protocol. Medium was then replaced with supplemented MCDB 131 with or without drug treatments and images of wounds were taken every 4 h until closure.
Empty vector (EV) and AOX pWPI GFP51 or EV and LbNOX pLV-EF1 RFP21 (VectorBuilder, VB160708-1059xrd) lentiviral constructs were transfected into COS1 cells using jetPRIME transfection reagent (Polyplus), along with pMD2.G and psPAX2 packaging vectors to produce EV-GFP, AOX-GFP, EV-RFP, and LbNOX-RFP lentiviruses. HUVECs were transduced with each lentivirus, and either GFP+ or RFP+ cells were sorted using FACS after 48 h. Cells were allowed to recover overnight and then plated for each assay.
Mice and tamoxifen administration
C57BL/6 mice harbouring a loxP-flanked exon 1 of the Uqcrq gene (encodes QPC) were generated by Ozgene. Mice were genotyped using the following primers: (1) 5′-CTTCCGCTCCTCCCGGAAGT-3′, (2) 5′-TTCCCAAACTCGCGGCCCATG-3′, and (3) 5′-CAATTCCAGCCAACAGTCCC-3′ which allow identification of the QPC wild-type, loxP-flanked, and excised alleles. QPC-floxed mice were crossed with Cdh5CreERT2 mice, which have a tamoxifen-inducible Cre allele under the endothelial-specific Cadherin 5 (Cdh5, aka VE-cadherin) promoter29. The Cre allele was detected by using the following primers: 1. 5′-AATCTCCCACCGTCAGTACG-3′, and 2. 5′-CGTTTTCTGAGCATACCTGGA-3′. Lox-stop-lox TdTomato mice were purchased from Jackson (stock no. 007914). Tamoxifen stock was prepared by dissolving tamoxifen (Sigma) in corn oil by shaking at 37 °C for 2 h. Tamoxifen was administered to mouse pups orally by dropping 2.5 µl of 40 mg ml−1 tamoxifen stock into the mouth using a p10 pipet. Pups were given five doses of tamoxifen from P0–P4 at a dose of 100 µg per pup per day. For tumour angiogenesis assays, adult mice were fed tamoxifen chow (Envigo) for 2 weeks prior to tumour injection and remained fed with tamoxifen chow for the entirety of the experiment. All animal procedures were approved by the Institutional Animal Care and Use Committee at Northwestern University.
Postnatal retinal angiogenesis assay
Mouse pups were dosed with tamoxifen as described above. Eyeballs were collected and fixed, and retinas were dissected on P7 as previously described52. Retinas were blocked/permeabilized in PBS + 5% BSA (OmniPur) + 0.5% TritonX-100 (Sigma) for 4 h at room temperature while rocking. Retinas were then stained with fluorescently labelled primary antibodies overnight at 4 °C, cut into clover leaves, and whole-mounted onto slides using Prolong Diamond Antifade Mounting Medium (Molecular Probes).
Isolation of endothelial cells from mouse lungs
Whole lungs were collected from QPC-WT and QPC-KO mice dosed with tamoxifen. Lungs were perfused with PBS and then with dispase (Corning). Lungs were then cut into small pieces using scissors and digested while shaking at 37 °C in 5 mg ml−1 collagenase type I (Gibco) and 1 mg ml−1 DNase I (Roche). Lung tissue was homogenized by passing through a 28-G needle several times and filtering through a 70-μm filter. Red cell lysis was performed by resuspending homogenized lung pellets in 1 ml of RBC lysis buffer for 2 min. For RNA-seq, homogenized lungs were stained with biotinylated CD45 and depleted of CD45+ cells using EasySep Mouse Streptavidin RapidSpheres Isolation Kit (Stemcell) by using the manufacturer’s protocol. CD31+ cells were then sorted by FACS. For metabolomics, homogenized lungs were stained with both biotinylated CD45 and biotinylated CD326/EpCAM and depleted of CD45+ and EpCAM+ cells using EasySep Mouse Streptavidin RapidSpheres Isolation Kit (Stemcell) by using the manufacturer’s protocol. Cells were then stained with CD31-FITC, and CD31+ cells were enriched using the Mouse FITC Selection Kit (Stemcell).
Tumours were fixed in 4% paraformaldehyde for 24 h and submitted for histological analysis. Briefly, organs were paraffin embedded and sectioned onto slides. Following standard deparaffinization, antigen retrieval was performed using pH 6 sodium citrate buffer at 110 °C for 20 min in a Biocare decloaking chamber. Slides were blocked and incubated with primary antibodies overnight at 4 °C. Blocking and secondary antibody steps were performed using an automated Intellipath staining system made by BioCare. Lungs were dissected and perfused through the trachea with a 1:1 solution of optimal cutting temperature medium (Fisher), embedded in optimal cutting temperature medium, and frozen in 2-methylbutane on dry ice. Frozen tissue was cryosectioned (7 µm), placed on slides, and fixed with acetone. Briefly, sections were blocked and then incubated with primary conjugated antibodies for 1 h at room temperature.
Antibodies, flow cytometry, and cell sorting
Lung homogenates were stained with the following antibodies: anti-mouse CD45 biotin (eBioscience, 13-0451-82, clone 30-F11, 1:100), anti-mouse CD326/EpCAM biotin (Invitrogen, 2020-10-01, clone G8.8, 1:100), anti-mouse CD31-FITC (BioLegend, 102406, clone 390, 1:100). For flow cytometry, live cells were gated by staining with GhostDye viability stain (Tonbo Biosciences, 1:200). Retinas were stained with the following antibodies/stains: GS isolectin-B4 (IB4) (Invitrogen, I21411, Alexa Fluor 488, 1:100), anti-phosphohistone 3 (Ser10) (Alexa Fluor 647, EMD Millipore, 06-570, polyclonal, 1:400), collagen IV (Goat, EMD Millipore, AB769, polyclonal, 1:25), cleaved caspase-3 (Rabbit, Cell Signaling, 9661, polyclonal, 1:100), Alexa Fluor 647 AffiniPure Bovine Anti-Goat (Jackson Immuno., 805-605-180, 1:500), and DyLight 405 AffiniPure Donkey Anti-Rabbit (Jackson Immuno., 711-475-152, 1:500). Sections were stained with the following antibodies: anti-CD31 (Goat, Santa Cruz, sc-1506, M-20, 1:250), anti-phosphohistone 3 (Ser10) (Rabbit, Abcam, ab5176, polyclonal, 1:1000), anti-goat Cy3 (Jackson Immuno., 1:1000), and anti-rabbit Alexa Fluor 488 (Jackson Immuno., 1:1000). Lung sections were stained with the following antibodies: anti-CD31 (PE-CF594, BD Horizon, 563616, MEC 13.3, 1:100) and anti-phosphohistone 3 (Ser10) (Alexa Fluor 647, EMD Millipore, 06-570, polyclonal, 1:100). Tumours and lung section nuclei were stained with 100 ng ml−1 of DAPI. Flow cytometry samples were run on LSR Fortessa flow cytometers (BD), and data were analysed using FlowJo software. Cell sorting was performed using the FACSAria II (BD).
Microscopy and image analysis
Images of HUVEC spheroids were taken using a Nikon AZ100 Multi-purpose Zoom Microscope and Nikon imaging software. Images were analysed using ImageJ software. Images of scratch-wound assays were taken every 4 h using the automated IncuCyte live-cell analysis system (Essen BioScience). Percentage wound closure was analysed and calculated using IncuCyte scratch-wound software (Essen BioScience). Retinal, lung, and tumour images were taken using the Nikon A1 Confocal Laser Microscope System and analysed using ImageJ software. Branchpoint analysis was performed using AngioTool software.
In vitro: HUVECs were plated in supplemented MCDB 131 medium and allowed to attach overnight. Medium was then replaced with supplemented MCDB 131 medium with or without drug and treated for 24 h. Cells were collected, and pellets were flash frozen in liquid nitrogen in cryovials. In vivo: endothelial cells were isolated from P15 mouse lungs as described above, and pellets were flash frozen in liquid nitrogen. Frozen pellets were stored at −80 °C until extraction. Metabolite extraction and sample analysis: samples were thawed and resuspended in 500 μL of ice cold 80% methanol (HPLC grade) and subjected to three freeze–thaw cycles alternating between liquid nitrogen and a 37 °C water bath. Next, samples were centrifuged at 18,000g for 10 min at 4 °C. The resulting supernatants were transferred to fresh tubes and dried. Samples were resuspended in 10 μl per 200,000 cells. Samples were analysed by high-performance liquid chromatography and high-resolution mass spectrometry and tandem mass spectrometry (HPLC–MS/MS). Specifically, the system consisted of a Thermo Q-Exactive in line with an electrospray source and an Ultimate 3000 (Thermo) series HPLC consisting of a binary pump, degasser, and auto-sampler outfitted with an Xbridge Amide column (Waters; dimensions of 4.6 mm × 100 mm and a 3.5 μm particle size). The mobile phase A contained 95% (vol./vol.) water, 5% (vol./vol.) acetonitrile, 20 mM ammonium hydroxide, 20 mM ammonium acetate, pH 9.0; B was 100% acetonitrile. The gradient was as following: 0–1 min, 15% A; 18.5 min, 76% A; 18.5–20.4 min, 24% A; 20.4–20.5 min, 15% A; 20.5–28 min, 15% A with a flow rate of 400 μl min−1. The capillary of the electron spray ionisation source was set to 275 °C, with sheath gas at 45 arbitrary units, auxiliary gas at 5 arbitrary units, and the spray voltage at 4.0 kV. In positive/negative polarity switching mode, an m/z scan range from 70 to 850 was chosen, and MS1 data were collected at a resolution of 70,000. The automatic gain control target was set at 1 × 106 and the maximum injection time was 200 ms. The top five precursor ions were subsequently fragmented, in a data-dependent manner, using the higher energy collisional dissociation cell set to 30% normalized collision energy in MS2 at a resolution power of 17,500. The sample volumes of 10 μl, which contained 200,000 cells, were injected. Data acquisition and analysis were carried out by Xcalibur 4.0 software and Tracefinder 2.1 software, respectively (both from ThermoFisher Scientific). NAD+/NADH ratio was measured using the NAD/NADH-glo Assay (Promega) by following the manufacturer’s protocol.
Mouse pups were dosed with tamoxifen and endothelial cells were isolated from P15 mouse lungs as described above. Cells were lysed with RLT buffer + 0.1% β-mercaptoethanol from the RNeasy Plus Micro Kit (Qiagen) and stored at −80 °C until RNA was extracted. RNA isolation was performed using the RNAeasy Plus Micro Kit (Qiagen) following the manufacturer’s instructions with an additional on-column DNase treatment using RNase-free DNase Set (Qiagen). RNA quality and quantity were measured using Agilent 4200 Tapestation using high Sensitivity RNA ScreenTape System (Agilent Technologies). NEBNext Ultra RNA (New England Biolabs) was used for full-length cDNA synthesis and library preparation. Libraries were pooled, denatured, and diluted, resulting in a 2.0 pM DNA solution. PhiX control was spiked at 1%. Libraries were sequenced on an Illumina NextSeq 500 instrument (Illumina) using NextSeq 500 High Output reagent kit (Illumina) (1 × 75 cycles) with a target read depth of approximately 8–16 million aligned reads per sample. FASTQ reads were trimmed using Trimmomatic to remove end nucleotides with a PHRED score less than 30, and a minimum length of 20 bp was required. Reads were then aligned to the mm10 genome using tophat version 2.1.04 using the following options: –no-novel-juncs –read-mismatches 2 –read-edit-dist 2 –max-multihits 20 –library-type fr- unstranded. The generated bam files were then used to count the reads only at the exons of genes using htseq-count5 with the following parameters: -q -m intersection-nonempty -s no -t exon. Differential expression analysis was done using the R package edgeR6. Bigwig tracks of RNA-seq expression were generated by using the GenomicAlignments package in R to calculate the coverage of reads in counts per million normalized to the total number of uniquely mapped reads for each sample in the library. GSEA analysis was done using the Broad Institute GSEA software7. In brief, the gene list output from edgeR was ranked by calculating a rank score of each gene as –log10(P value) × sign(logFC). A pre-ranked GSEA analysis was done using 3000 permutations and the Hallmark pathway database.
Tumour angiogenesis assay
Eight-week-old mice were dosed with tamoxifen as described above. The right flank was shaved, and 100,000 B16-F10 melanoma cells (ATCC) were injected subcutaneously. Tumour size was measured using calipers every 2 to 3 d for 21 d. According to the approved protocol, tumours were not permitted to exceed 2 cm in diameter (approximately 4 cm3). Tumour volume was calculated by first determining the geometric mean between the length and width, then using the equation: (4 / 3) × (π) × (geometric mean / 2)3. On day 21, tumours were collected and weighed, tumour size was measured, and volume was calculated using the equation: (4/3) × (π) × (length / 2) × (width / 2) × (depth / 2).
Mass spectrometry to identify histone modifications
Nuclei were isolated using gentle detergent treatment (0.3% NP-40 in NIB-250 buffer) of cells and centrifugation at 0.6g. Detergent was removed by 2× washing of obtained pellets with NIB-250 without NP-40 buffer. Histones from isolated nuclei were acid extracted and derivatized with propionic anhydride both before and after trypsin digestion as previously described53. Propionylated histone peptides were resuspended in 50 μl water with 0.1% trifluoroacetic acid, and 3 μl was injected in three technical replicates on a nanoLC/triple quadrupole MS that consisted of a Dionex UltiMate 3000 coupled to a ThermoFisher Scientific TSQ Quantiva triple quadrupole mass spectrometer. Buffer A was 100% LC–MS-grade water with 0.1% formic acid and buffer B was 100% acetonitrile. The propionylated peptides were loaded onto an in-house packed C18 trapping column (4 cm × 150 μm; Magic AQ C18, 3 μm, 200 Å-Michrom) for 10 min at a flow rate of 2.5 μl min−1 in 0.1% trifluoroacetic acid loading buffer. The peptides were separated by a gradient from 1 to 35% buffer B from 5 to 45 min. The analytical column was 10 cm × 75 μm PicoChip (1PCH7515-105H253-NV New Objective) and consisted of the same C18 material as the trapping column. The triple quadrupole settings were as follows: collision gas pressure of 1.5 mtorr; Q1 peak width of 0.7 (full-width at half-maximum); cycle time of 3 s; skimmer offset of 10 V; electrospray voltage of 2.5 kV. Selected reaction monitoring mass spectrometer transitions were developed as described previously54,55. Data were analysed using Skyline software (v3.5; MacCoss Lab, University of Washington) with Savitzky–Golay smoothing of peaks56. Automatic peak assignment and retention times were checked manually.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
All data from this manuscript are available from the corresponding author upon request. RNA sequence data that support the findings of this study have been deposited in GEO with the accession code GSE121770.
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This work was supported by the National Institutes of Health (NIH) grant nos R35CA197532, 5P01AG049665 and 5P01HL071643-13 (to N.S.C.) and T32 GM08061 (to L.P.D.). We thank R. Adams for Cdh5CreERT2 mice. Imaging work was performed at the Northwestern University Center for Advanced Microscopy generously supported by NCI CCSG P30 CA060553 awarded to the Robert H Lurie Comprehensive Cancer Center. Histology services for tumour tissue was provided by the Northwestern University Research Histology and Phenotyping Laboratory, which is supported by NCI P30 CA060553 awarded to the Robert H. Lurie Comprehensive Cancer Center. Analysis of histone modifications was performed by the Northwestern Proteomics Core Facility, generously supported by NCI CCSG P30 CA060553 awarded to the Robert H. Lurie Comprehensive Cancer Center and the National Resource for Translational and Developmental Proteomics supported by P41 GM108569. Flow cytometric analysis was supported by the Northwestern University Flow Cytometry Core Facility supported by NCI CCSG P30 CA060553. Flow cytometry cell sorting was performed on a BD FACSAria SORP system, purchased through the support of NIH grant 1S10OD011996-01. Metabolomics services were performed by the Metabolomics Core Facility at Robert H. Lurie Comprehensive Cancer Center of Northwestern University. We would like to thank H. Abdala-Valencia and K. Nam for RNA sequencing. Finally, we would like to thank G. Oliver at Northwestern University for his helpful intellectual input.
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Nature Metabolism (2019)