Control of endothelial quiescence by FOXO-regulated metabolites

Andrade, Jorge; Shi, Chenyue; Costa, Ana S. H.; Choi, Jeongwoon; Kim, Jaeryung; Doddaballapur, Anuradha; Sugino, Toshiya; Ong, Yu Ting; Castro, Marco; Zimmermann, Barbara; Kaulich, Manuel; Guenther, Stefan; Wilhelm, Kerstin; Kubota, Yoshiaki; Braun, Thomas; Koh, Gou Young; Grosso, Ana Rita; Frezza, Christian; Potente, Michael

doi:10.1038/s41556-021-00637-6

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Published: 01 April 2021

Control of endothelial quiescence by FOXO-regulated metabolites

Nature Cell Biology volume 23, pages 413–423 (2021)Cite this article

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Abstract

Endothelial cells (ECs) adapt their metabolism to enable the growth of new blood vessels, but little is known how ECs regulate metabolism to adopt a quiescent state. Here, we show that the metabolite S-2-hydroxyglutarate (S-2HG) plays a crucial role in the regulation of endothelial quiescence. We find that S-2HG is produced in ECs after activation of the transcription factor forkhead box O1 (FOXO1), where it limits cell cycle progression, metabolic activity and vascular expansion. FOXO1 stimulates S-2HG production by inhibiting the mitochondrial enzyme 2-oxoglutarate dehydrogenase. This inhibition relies on branched-chain amino acid catabolites such as 3-methyl-2-oxovalerate, which increase in ECs with activated FOXO1. Treatment of ECs with 3-methyl-2-oxovalerate elicits S-2HG production and suppresses proliferation, causing vascular rarefaction in mice. Our findings identify a metabolic programme that promotes the acquisition of a quiescent endothelial state and highlight the role of metabolites as signalling molecules in the endothelium.

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Main

Homeostasis of the blood vasculature relies on a single layer of endothelial cells (ECs) forming the inner surface of all vessels. In most adult tissues, ECs reside in a non-cycling, quiescent state, which is critical for their function as a barrier and signalling interface^1,2. This resting state is reversible, and ECs can (re-)activate, enter the cell cycle and expand to meet physiological or stress-induced demands. While the processes of endothelial activation and proliferation are being defined with increasing molecular resolution, the mechanisms leading to the acquisition of a quiescent phenotype remain poorly understood.

Previous studies have demonstrated that the forkhead box O (FOXO) transcription factor FOXO1 is a critical driver of endothelial quiescence^3,4. FOXO1 activity is controlled by the phosphatidylinositol-3-OH kinase–AKT pathway that inhibits FOXO1 transcriptional responses through AKT-mediated phosphorylation^5,6. ECs are highly sensitive to changes in FOXO1 activity, as both EC-specific deletion and forced activation are early embryonic lethal in mice^3,4,7,8. Loss of FOXO1 leads to unregulated endothelial proliferation and vascular overgrowth, whereas forced activation causes premature quiescence and vascular rarefaction. FOXO1 promotes quiescence, in part, by suppressing MYC signalling, which leads to a reduction in endothelial metabolic activity⁴. Yet, the FOXO1-regulated metabolic programmes in ECs are largely unknown. Given the importance of endothelial metabolic regulation for vascular growth and function^{9,10,11,12,13,14,15,16,17,18,19} and the sensitivity of ECs towards changes in FOXO1 activity, we sought to investigate the global metabolic profile of FOXO1-induced ECs.

Results

FOXO1 activation induces 2-hydroxyglutarate generation in ECs

We performed untargeted metabolomics of human umbilical vein endothelial cells (HUVECs) that were transduced with adenoviruses encoding a constitutively active FOXO1 mutant (AdFOXO1^A3) or green fluorescent protein (GFP) as a control (AdCtrl). This mutant has the three AKT phosphorylation sites replaced by alanine (→FOXO1^A3), which renders FOXO1 in the nucleus^5,6. ECs expressing FOXO1^A3 were in a reversible proliferation arrest and exhibited a molecular signature characteristic of cellular quiescence, including suppression of the proliferation markers MYC and proliferating cell nuclear antigen (PCNA), as well as induction of the cell cycle inhibitor p27 (CDKN1B) and the repressive histone mark histone H3 lysine 27 trimethylation (H3K27me3)^20,21,22,23 (Extended Data Fig. 1a–c,f).

Principal component analysis (PCA) of the intracellular metabolites collected 24 h post-transduction revealed a separate clustering of control and FOXO1^A3-transduced ECs (Fig. 1a). ECs with nuclear FOXO1 exhibited increased levels of several branched-chain amino acid (BCAA) metabolites and of early tricarboxylic acid (TCA) cycle intermediates (Supplementary Table 1). Notably, the most increased metabolite (7.6-fold) was 2-hydroxyglutarate (2HG) (Fig. 1b and Extended Data Fig. 1d), which is a chiral molecule derived from 2-oxoglutarate (2OG) that functions as a competitive inhibitor of many 2OG-dependent dioxygenases. Among these are Jumonji C (JmjC) domain-containing lysine demethylases and hypoxia-inducible factor (HIF)-regulating prolyl hydroxylases (PHDs)^{24,25,26,27,28,29}, which suggests that FOXO1 may exert profound effects on gene expression via regulation of this metabolite.

**Fig. 1: Activation of FOXO1 signalling induces the generation of S-2HG in ECs.**

We therefore confirmed the regulation of 2HG using a lentiviral system for doxycycline-inducible expression of FOXO1^A3 (Fig. 1c and Extended Data Fig. 1e) and contact inhibition of endothelial proliferation—a stimulus for nuclear translocation and activation of endogenous FOXO1 (Fig. 1e–i and Extended Data Fig. 1g). Indeed, we observed an 8.4-fold increase in 2HG abundance in dense (contact-inhibited) culture conditions, whereby FOXO1 is an induced and predominantly nuclear protein and in which canonical FOXO1 target genes are highly transcribed (for example, MAX interactor 1 (MXI1), pyruvate dehydrogenase kinase 1 (PDK1), pyruvate dehydrogenase kinase 4 (PDK4) and CD36 molecule (CD36)) (Fig. 1f–i and Extended Data Fig. 1g). Thus, ECs generate 2HG when FOXO1 signalling is activated.

FOXO1-activated ECs produce S-2HG, which inhibits 2OG-dependent dioxygenases

Two conformations of 2HG exist, the R-2HG and S-2HG enantiomers, which have similar but not identical functions^30,31,32. Chiral derivatization followed by liquid chromatography–mass spectrometry (LC–MS) revealed that FOXO1 induces the preferential generation of S-2HG, which is also the more abundant enantiomer in ECs (Fig. 1d,j). To explore whether 2HG regulates the activity of endothelial 2OG-dependent dioxygenases, we first analysed HIFs, which are marked by PHDs for proteasomal degradation under normoxic conditions. In line with an inhibitory effect of 2HG on PHDs^{30,31,33,34,35}, incubation of HUVECs with cell-permeable forms of R-2HG or S-2HG increased HIF1α and HIF2α protein abundance and enhanced HIF target gene expression (Extended Data Fig. 2a–c). Interestingly, S-2HG had more profound effects on HIF responses than R-2HG (Extended Data Fig. 2a–c). Similar results were obtained for H3K27me3, which is targeted by the 2OG-dependent JmjC histone demethylases (Extended Data Fig. 2d), which suggests that S-2HG is the more potent inhibitor of 2OG-dependent dioxygenases in ECs. Therefore, we focused our further analysis on this enantiomer.

S-2HG promotes a quiescent endothelial state

To characterize the role of S-2HG in ECs, we incubated HUVECs with different S-2HG concentrations and noted a time- and dose-dependent suppression of endothelial proliferation (Fig. 2a–c and Extended Data Fig. 3a). S-2HG-treated ECs were arrested in the G0/G1 phase of the cell cycle, and markers of cell growth and proliferation, including MYC, PCNA and phosphorylated ribosomal protein S6, were diminished (Fig. 2d,e and Extended Data Fig. 3b). S-2HG did not cause metabolic distress (depletion of a particular metabolite), senescence or apoptotic cell death, as reporters of these processes were unaltered or not detectable (Fig. 2f and Extended Data Fig. 3c–f). To further characterize this antiproliferative effect, we transduced HUVECs with a lentiviral vector encoding the fluorescent ubiquitination-based cell cycle indicator (FUCCI). This dual-colour reporter labels nuclei of cells in the G0/G1 phase in red and those in the S/G2/M phases in green (Extended Data Fig. 3g,h). S-2HG caused a progressive increase in ECs with red fluorescence, thereby confirming the G0/G1 arrest (Fig. 2g and Supplementary Videos 1 and 2). However, S-2HG withdrawal led to the reappearance of ECs with green fluorescent nuclei (Fig. 2g and Supplementary Videos 1 and 2), which indicates that the S-2HG-induced proliferation arrest is reversible.

**Fig. 2: S-2HG supports a quiescent endothelial phenotype.**

RNA sequencing (RNA-seq) of HUVECs treated with vehicle or S-2HG for 24 h revealed that S-2HG regulates genes strongly linked to proliferation (Fig. 2h, Extended Data Fig. 4a–c and Supplementary Table 2). A total of 43 out of the 50 most downregulated genes are involved in cell cycle progression and cell division, including DNA topoisomerase II alpha (TOP2A), anillin (ANLN), marker of proliferation Ki-67 (MKI67) and cyclin-dependent kinase 1 (CDK1) (Extended Data Fig. 4b). Gene set enrichment analysis (GSEA) corroborated this notion (Fig. 2i and Extended Data Fig. 4c). Moreover, numerous quiescence-associated genes³⁶, including those that are not directly linked to cell proliferation, were enriched in S-2HG-treated ECs (Fig. 2i). These data suggest that S-2HG regulates not only endothelial proliferation but also, more generally, promotes a quiescent endothelial state. Consistent with this idea, S-2HG-treated ECs showed a number of cellular phenotypes that are characteristic for quiescent cells²³, including lower metabolic activity and reduced RNA and protein synthesis (Fig. 2j–m and Extended Data Fig. 4d).

S-2HG limits the angiogenic behaviour of ECs

To seek further evidence of a quiescence-promoting function of S-2HG, we visualized endothelial angiogenic behaviour in an in vitro scratch assay and found that S-2HG-treated ECs were less motile (Fig. 3a,b and Supplementary Videos 3 and 4). Moreover, analysis of three-dimensional endothelial spheroid cultures revealed that S-2HG-treated spheroids formed fewer and shorter sprouts compared to controls (Fig. 3c and Extended Data Fig. 5a). We validated the relevance of these findings by studying blood vessel growth in the mouse retina, in which blood vessels develop postnatally in a highly stereotypical manner. Postnatal day 5 (P5) pups received a single intraocular injection of S-2HG in one eye and vehicle in the other (Extended Data Fig. 5b). Analysis of the retinal vasculature 2 days later (at P7) showed a poorly connected and hypocellular endothelial network in S-2HG-treated retinas (Fig. 3d,e,i and Extended Data Fig. 5c). This phenotype was particularly evident at the angiogenic front, where most of the endothelial growth and proliferation occurs, while central parts of the retinal vasculature that formed before the injection were less affected (Fig. 3d,e). The sparse network was not due to increased vessel pruning or apoptotic cell death, since analysis of endothelial-less basement membrane sleeves or cleaved caspase-3-positive ECs did not reveal significant changes (Fig. 3g–i). Instead, we found a marked suppression of EC proliferation, which was indicated by a reduction in 5-ethynyl-2′-deoxyuridine (EdU)-positive endothelial nuclei (Fig. 3f,i and Extended Data Fig. 5c). These data are consistent with an anti-angiogenic activity of S-2HG in ECs and suggest that S-2HG enforces the pro-quiescent function of FOXO1.

**Fig. 3: S-2HG restrains the angiogenic activity of ECs.**

FOXO1 induces S-2HG generation by inhibiting the OGDH enzyme

Previous studies have shown that S-2HG is generated in conditions of reduced mitochondrial function in which 2OG becomes promiscuously reduced to S-2HG^{33,34,35,37,38,39,40,41}. To understand how ECs generate S-2HG in response to FOXO1 activation, we examined mitochondrial metabolism. This analysis revealed altered mitochondrial respiration in FOXO1^A3-expressing ECs (Extended Data Fig. 6a), which suggests that there is diminished activity of the 2OG dehydrogenase (OGDH) enzyme—a multisubunit complex that catalyses the conversion of 2OG to succinyl-CoA (Extended Data Fig. 6b). Indeed, we observed that FOXO1^A3-expressing ECs accumulated the OGDH substrate 2OG, while succinyl-carnitine—a surrogate metabolite of succinyl-CoA—was depleted (Fig. 4a and Supplementary Table 1). Of note, recent genetic studies identified OGDH inhibition as a mechanism for S-2HG generation^33,42,43. We therefore used the clustered regularly interspaced short palindromic repeats (CRISPR)–Cas9 method to specifically inactivate OGDH in ECs (Fig. 4b). Transduction of HUVECs with lentiviral constructs encoding FLAG-tagged Cas9 (Cas9^FLAG) and guide RNAs (gRNAs) targeting OGDH (gOGDH) led to the expected accumulation of 2OG and a decline in the mitochondrial oxygen consumption rate (OCR) (Fig. 4c and Extended Data Fig. 6c). Importantly, OGDH deficiency increased endothelial S-2HG levels and abrogated the ability of ECs to proliferate (Fig. 4d–g). The growth inhibitory effect was not caused by the increased 2OG levels, because treatment of HUVECs with (cell permeable) 2OG did not influence their capacity to sprout or divide (Fig. 4h and Extended Data Fig. 6d,e). Moreover, it was not a generic response to TCA cycle perturbation, because inactivation of two other TCA cycle enzymes, succinate dehydrogenase (SDH) and fumarate hydratase (FH), neither increased S-2HG levels nor inhibited endothelial proliferation (Fig. 4i–k and Extended Data Fig. 6f–k). These data suggest a crucial role of OGDH in endothelial cell-cycle control, which relies on the signalling metabolite S-2HG.

**Fig. 4: OGDH inactivation induces S-2HG generation and limits endothelial proliferation.**

Endothelial OGDH is crucial for angiogenic growth

To determine the physiological relevance of these findings, we generated mice in which exon 3 and 4 of the Ogdh gene are flanked by loxP sites (Ogdh^fl) (Extended Data Fig. 7a). Deleting these exons in ECs (and some haematopoietic cells) using a constitutive Tie2-cre transgene (Ogdh^EC-KO) caused severe defects in the yolk sac vasculature that led to developmental retardation and embryonic death (Fig. 5a,b and Extended Data Fig. 7b,c). To further characterize the functional consequences of Ogdh deletion, we analysed blood vessel growth in the postnatal retina. To this end, we bred Ogdh^fl mice with mice expressing the tamoxifen-inducible recombinase creERT2 from the endothelial-restricted Cdh5 promoter (Ogdh^iECKO). 4-Hydroxytamoxifen (4-OHT)-induced depletion of OGDH caused a sparse vascular network that mimicked the defective vasculature in the constitutive Ogdh knockout embryos (Fig. 5c–e and Extended Data Fig. 7d–g). Ogdh mutant ECs had elevated S-2HG levels and exhibited diminished proliferation, but lacked signs of increased cell death (Fig. 5e–i and Extended Data Fig. 7f,g). This result underscores the importance of OGDH function for endothelial proliferation and vascular growth control.

**Fig. 5: Loss of endothelial OGDH restricts vascular growth.**

FOXO1 inhibits OGDH function via the generation of BCAA catabolites

We then investigated the mechanisms through which FOXO1 regulates OGDH function. First, we examined whether FOXO1 regulates the expression of components of the OGDH complex. For this purpose, we performed RNA-seq analysis in control and FOXO1^A3-expressing HUVECs. While FOXO1 governed the expression of prototypical FOXO1 target genes in the expected manner, we failed to detect changes in components of the OGDH complex that would explain a reduction in enzymatic activity (Fig. 6a). Similar data were obtained at the protein level (Extended Data Fig. 8a). We therefore considered alternative mechanisms of regulation. Our initial metabolomics analysis showed that, besides 2HG, FOXO1 induces the accumulation of 3-methyl-2-oxovalerate (KMV), 3-methyl-2-oxobutyrate (KIV) and 4-methyl-2-oxopentanoate (KIC), and other intermediates of BCAA catabolism (Fig. 6b–d). Strikingly, these metabolites are known inhibitors of OGDH^34,38,44,45, which suggests that the FOXO1-induced increase in these branched chain α-keto acids (BCKAs) leads to inhibition of the OGDH complex. Consistent with this model, KMV reduced mitochondrial oxygen consumption and OGDH activity, and led to a 9.7-fold increase in endothelial S-2HG levels (Fig. 6e–h).

**Fig. 6: FOXO1 induces S-2HG generation by regulating BCAA catabolism.**

Genes involved in BCAA metabolism are direct FOXO1 target genes

We probed further to elucidate how FOXO1 regulates BCKA generation. Interrogation of the transcriptional signature of FOXO1^A3-transduced HUVECs demonstrated that FOXO1 induced several genes coding for enzymes of the BCAA catabolic pathway, including dihydrolipoamide branched chain transacylase E2 (DBT), branched chain keto acid dehydrogenase E1 subunit beta (BCKDHB), acyl-CoA dehydrogenase short/branched chain (ACADSB) and methylmalonyl-CoA mutase (MUT), among others (Fig. 7a,b and Extended Data Fig. 8b). Notably, DBT and BCKDHB are subunits of the branched-chain α-ketoacid dehydrogenase (BCKD) complex, which oxidizes KMV, KIC and KIV to branched-chain acyl-CoAs and constitutes the rate-limiting step in BCAA metabolism⁴⁶. Moreover, expression of protein phosphatase, Mg²⁺/Mn²⁺-dependent 1K (PPM1K), a phosphatase that activates the BCKD complex⁴⁶, was also increased by FOXO1, which suggests that FOXO1 coordinates the expression of genes involved in BCAA catabolism. Of note, FOXO1 regulates these genes independently of its repressive effects on MYC signalling⁴, since inactivation of MYC by CRISPR–Cas9 did not suppress the levels of these BCAA transcripts (Extended Data Fig. 8c–e).

**Fig. 7: The FOXO1-regulated BCAA catabolite KMV limits endothelial proliferation.**

We therefore studied whether FOXO1 directly targets these genes, and conducted FOXO1 chromatin immunoprecipitation with sequencing (ChIP-seq) in FOXO1^A3-expressing ECs. We found that FOXO1 peaks were enriched for the high-affinity FOXO1-binding motif and were preferentially located at gene promoters close to the transcriptional start site (Extended Data Fig. 9a,b). FOXO1 ChIP-seq identified bona fide FOXO1 targets, including several of the differentially regulated BCAA genes (DBT, DLD, PPM1K, MUT and OXCT1) (Fig. 7c,d, Extended Data Fig. 9c–f and Supplementary Table 3). These FOXO1-bound genes were also in an active state, as indicated by the acetylation of H3K27 and the trimethylation of H3K4 (Fig. 7c,d and Extended Data Fig. 9c–f). These results demonstrate that FOXO1 drives a transcriptional programme involved in BCAA catabolism and suggest a mechanism for the generation of BCKAs such as KMV.

BCAA metabolites restrict endothelial proliferation and vascular growth

Finally, we assessed the impact of BCKAs on ECs by treating HUVECs with KMV. Similar to S-2HG, KMV suppressed the proliferative activity of ECs without affecting their viability (Fig. 7e,f). Intraocular injection of KMV in neonatal pups (at P5) also reduced endothelial proliferation, giving rise to a sparse vascular network that resembled the vasculature in S-2HG-treated mice (Fig. 7g–i). Although we cannot exclude that KMV might also signal via alternative pathways, these data strongly suggest a pivotal role for KMV (and other BCKAs) in FOXO1-induced quiescence signalling.

Discussion

In summary, we identified a network of FOXO1-regulated metabolites that promote the acquisition of a quiescent endothelial state, and in which the metabolic signalling molecule S-2HG plays a critical role. We propose that endothelial S-2HG levels increase in response to FOXO1 activation to induce a potent but reversible cell cycle arrest. Our data suggest that FOXO1 stimulates S-2HG generation by increasing BCKAs such as KMV, which inhibit the activity of the mitochondrial OGDH complex. Inhibition of this TCA cycle enzyme results in the accumulation of its substrate 2OG, which is subsequently reduced to S-2HG via promiscuous substrate usage^34,35,37,38. Changes in endothelial FOXO1 activity therefore determine whether 2OG is used in the TCA cycle to support EC proliferation or is converted to S-2HG to signal quiescence. Our data are consistent with studies of several cancer cell lines showing that OGDH activity is necessary for cell proliferation⁴⁷ and that S-2HG can mount antiproliferative responses^48,49.

The precise mechanisms by which S-2HG limits endothelial cell-cycle progression remain to be addressed. We found that S-2HG leads to increased HIF abundance and a robust HIF transcriptional response, which is in accordance with the inhibitory function of S-2HG on PHD enzymes^30,31,34,35 and the cell-cycle-arresting activity of HIFs^50,51. Importantly, partial inactivation of PHD2 in ECs has also been shown to cause a resting (phalanx) phenotype via stabilization of HIF proteins⁵², which suggests that PHDs are quantitatively sensitive targets of S-2HG that may contribute to its endothelial effects. In line with these reports, we found that genetic PHD inactivation lowers the proliferative capacity of ECs (Extended Data Fig. 10).

Yet, S-2HG inhibits a variety of other 2OG-dependent dioxygenases, including JmjC histone lysine demethylases, ten-eleven translocation (TET) DNA hydroxylases and members of the AlkB family of proteins, which control histone, DNA and RNA methylation, respectively^{33,37,38,39,48,53}. Although the vascular functions of these epigenetic regulators are largely unknown, it seems probable that they contribute to the effects of S-2HG in ECs. Future studies should aim to elucidate the role of these enzymes in the endothelium and assess their regulation by S-2HG and other small-molecule metabolites. Understanding such metabolic signalling mechanisms will provide insight into how blood vessels develop and are maintained, and how their functions become abnormal in disease states.

Methods

Cells and cell culture

Pooled HUVECs were purchased from Lonza (CC-2519) and cultured in endothelial basal medium (EBM; Lonza) supplemented with hydrocortisone (1 µg ml⁻¹), bovine brain extract (12 µg ml⁻¹), gentamicin (50 µg ml⁻¹), amphotericin B (50 ng ml⁻¹), human recombinant epidermal growth factor (10 ng ml⁻¹) and 10% fetal bovine serum (FBS; Life Technologies). Human embryonic kidney cells (HEK293FT) were purchased from Life Technologies (R70007) and cultured in DMEM supplemented with 10% FBS (Life Technologies) and geneticin (500 µg ml⁻¹; Invitrogen). Cells were tested negative for mycoplasma and maintained at 37 °C in a humidified atmosphere with 5% CO₂.

The isolation of mouse lung ECs was performed as previously described^4,54. In brief, lungs from adult Ogdh^fl/fl mice were washed with ice-cold Hanks buffer (Gibco, 14175-053), dissociated into smaller fragments and incubated with dispase II (Gibco, 17105-041). The homogenate was filtered through a cell strainer, collected by centrifugation and washed with PBS (Gibco, 14190-094) containing 0.1% BSA (PBSB). The resulting cell suspension was incubated with rat anti-mouse VE-cadherin antibody-coated (BD Pharmingen, 555289) magnetic beads (Dynabeads, Invitrogen, 11035). Beads were washed with PBSB and then resuspended in D-MEM/F12 (Invitrogen) supplemented with 20% fetal calf serum (FCS), endothelial growth factor (Promocell, c-30140), penicillin and streptomycin. The isolated cells were seeded on gelatin-coated culture dishes and repurified with the VE-cadherin antibody during the first three passages.

Cell treatments

Cell-permeable (2R)-octyl-α-hydroxyglutarate (R-2HG; 16366) and (2S)-octyl-α-hydroxyglutarate (S-2HG; 16367) from Cayman Biochemical were reconstituted in dimethylsulfoxide (DMSO; Sigma, D4540). Cell-permeable 2OG was obtained from Sigma (349631). For most studies, HUVECs were treated with 800 µM of R-2HG, S-2HG or 2OG, and DMSO was used as a vehicle control^35,37,39,48. (±)-3-Methyl-2-oxovaleric acid sodium salt (KMV; Sigma, K7125) was reconstituted in PBS and used at 10 mM at the indicated time points. To block the PHD-dependent degradation of HIFs, cells were treated with 1 mM DMOG (Sigma, D3695) for 6 h. To inhibit autophagy, HUVECs were treated overnight with 50 µM chloroquine (Sigma, C6628). Endothelial apoptosis was induced by treating HUVECs with 10 µg ml⁻¹ cycloheximide (Merck-Millipore, 239765) and 10 ng ml⁻¹ TNFα (TNF; Invitrogen, PHC3015) for 6 h. For cellular senescence studies, HUVECs were exposed to 100 µM hydrogen peroxide (H₂O₂) for 1 h. To measure changes in protein synthesis by surface sensing of translation (SUnSET)⁵⁵, HUVECs were pulsed with 1 µg ml⁻¹ puromycin (InvivoGen, ant-pr-1) for 15 min. Protein synthesis was blocked by pretreatment with 100 µM cycloheximide (Merck-Millipore, 239765) for 1 h before the puromycin pulse was applied.

Adenoviral transductions

Subconfluent HUVECs were transduced with custom-made adenoviruses (Vector Biolabs) to overexpress the FLAG-tagged human FOXO1^A3 (AdFOXO1^A3) that is constitutively nuclear⁴. Adenoviruses that contained an empty CMV promoter (AdCtrl; 1300, Vector Biolabs) were used as a control. For transductions, HUVECs were incubated in EBM (Lonza) containing 0.1% (v/v) BSA (Sigma, 1595) for 4 h and transduced with AdCtrl or AdFOXO1^A3 for an additional 4 h in the presence of 8 µg ml⁻¹ polybrene (Santa Cruz). Afterwards, HUVECs were washed five times with Hank’s balanced salt solution (Life Technologies) and cultured in EBM with 10% FBS and supplements. Adenoviral transductions of mouse lung ECs isolated from Ogdh^fl/fl mice were performed with Cre-encoding adenovirus (AdCre, 1045, Vector Biolabs) or the respective control (AdCtrl).

Lentiviruses and lentiviral transductions

For doxycycline-inducible lentiviral expression of a constitutively nuclear human FOXO1 (FOXO1^A3), FLAG-tagged FOXO1^A3 was cloned into pLVX-TetOne-Puro (Clontech). For CRISPR–Cas9 genome editing, gene-specific gRNA sequences (Supplementary Table 4) were cloned into a plentiCRISPRv2 plasmid co-expressing FLAG-tagged Cas9 nuclease and a puromycin-selection marker (Addgene, 52961). The FUCCI lentiviral reporters mCherry-hCdt1(30/120)-pCSII-EF and mVenus-hGeminin(1/110)-pCSII-EF⁵⁶ were obtained from the Laboratory for Cell Function Dynamics, CBS, RIKEN, Japan.

Lentivirus production was performed by co-transfection of HEK293FT cells with pMD2.G (Addgene, 12259), psPAX2 (Addgene, 12260) and transfer plasmids, accordingly. Transfections were carried out using Lipofectamine 2000 transfection reagent (Life Technologies) as previously described⁵⁷. Viruses were collected 48 and 72 h after transfection, filtered through a 0.45-μm filter and incubated with HUVECs for 16 h in the presence of 8 µg ml⁻¹ polybrene (Santa Cruz). After transduction, cells were expanded for 48 h and selected with EBM containing 1 µg ml⁻¹ puromycin (InvivoGen, ant-pr-1).

CRISPR–Cas9 genome editing of HUVECs

Gene-specific gRNA sequences were designed using the Genetic Perturbation Platform from the Broad Institute (https://portals.broadinstitute.org/gpp/public/). For each target gene, three independent gRNAs were used and co-transfected with the packaging vectors for lentivirus production. After transduction, puromycin was used to select a pool of CRISPR–Cas9-mediated mutant HUVECs. Knockout efficiency was confirmed by western blot analysis. Single gRNA sequences are provided in Supplementary Table 4.

Proliferation assays

HUVECs were seeded onto 6-well plates at a density of 25,000 cells per well and left to attach overnight. Next day (0 h), the indicated treatments were added to the culture medium and total cell numbers were counted every 24 h. Cell culture medium was replaced every 48 h. For ³H-thymidine incorporation into DNA, HUVECs treated for 48 h with S-2HG or vehicle were pulsed with 1 μCi μl⁻¹ ³H-thymidine (Perkin Elmer) for 6 h before collection. Samples were processed as previously described¹⁸, and scintillation was measured with a liquid scintillation analyser (Tri-Carb 2810R, Perkin Elmer). Data were normalized to the total protein concentration per sample.

RNA and protein synthesis assays

RNA synthesis was measured by ¹⁴C-glucose incorporation into RNA. HUVECs were pulsed with 0.22 μCi μl⁻¹ d-(6-¹⁴C)-glucose (Perkin Elmer) for 48 h before collection. Samples were processed and analysed as previously described⁵⁸. Data were normalized to the total RNA concentration per sample. To measure protein synthesis by ³H-tyrosine incorporation into newly synthesized protein, HUVECs were pulsed with 1 μCi μl⁻¹ ³H-tyrosine (Perkin Elmer) for 6 h before collection. Samples were processed and analysed as previously described¹². Data were normalized to the total protein concentration per sample.

Cell cycle analysis by flow cytometry and live-cell imaging

Cell cycle analysis of HUVECs was performed using a BrdU Flow kit (BD Pharmingen BrdU-APC Flow kit, 557892). HUVECs were pulsed with 10 µM BrdU in culture medium for 45 min before cell collection. Samples were further processed according to the manufacturer’s protocol and analysed by flow cytometry on a LSR Fortessa instrument (BD Biosciences). Analysis of endothelial cell-cycle dynamics was performed in HUVECs expressing the FUCCI reporter⁵⁶ using an IncuCyte Live-Cell imaging system (Essen BioScience).

Western blotting

Cell lysates, subcellular fractionations and western blot analyses were performed as previously described⁵⁹. The antibodies and amounts used are listed in Supplementary Table 5.

Spheroid assay

HUVECs resuspended in 80% EBM and 20% Methocel (v/v) were seeded onto a 96-well U-bottomed suspension cell culture plate for 24 h. Spheroids were collected, centrifuged at 200 × g for 5 min and resuspended in 80% (v/v) Methocel with 20% (v/v) FBS (Life Technologies). The spheroid–Methocel solution was mixed with equal volumes of collagen matrix and seeded onto a 24-well plate. The plate was incubated for 30 min to allow polymerization and then supplemented with 100 µl of EBM with control vehicle (DMSO), 2OG or S-2HG. Endothelial sprouting was allowed for 24 h, after which the spheroid-containing gels were washed with PBS and fixed for 30 min at room temperature in 10% formaldehyde. Bright-field images were acquired, and angiogenic parameters assessed from at least five spheroids per condition. Spheroids were further labelled with Phalloidin-iFluor 488 (Abcam, 176753) overnight at 4 °C, and representative z-stack images were acquired by confocal microscopy.

Scratch-wound assay

Scratch-wound assays were performed using an IncuCyte Cell Migration kit (Essen BioScience, 4493). HUVECs (10,000 per well) were plated onto 96-well ImageLock microplates (Essen BioScience, 4379) and cultured overnight in EBM with 10% FBS and supplements. A uniform scratch-wound was generated using an IncuCyte 96-well WoundMaker (Essen BioScience) following the manufacturer’s instructions. Cells were washed three times with culture medium and cultured with EBM with 10% FBS, supplements and treatments. Images were taken every hour until wound closure. The wound area at 0 and 12 h was measured and the results are expressed as percentage of scratch closure.

Senescence-associated β-galactosidase staining

Senescence-associated β-galactosidase staining was performed using a kit from Cell Signaling Technology (9860S). For each sample, five representative images were used to quantify the number of cells positive for senescence-associated β-galactosidase. H₂O₂-treated HUVECs were used as a positive control. Images were acquired using a LSM800 microscope (Zeiss) with a ×20 objective. Volocity (Perkin Elmer), Fiji/ImageJ, Photoshop (Adobe) and Illustrator (Adobe) software were used for image quantification and processing.

Immunohistochemistry of cell cultures

HUVECs were seeded on glass-bottom culture dishes (Mattek) coated with fibronectin (50 μg ml⁻¹) and cultured at 37 °C and 5% CO₂. To detect FOXO1 subcellular localization, sparse and dense HUVEC cultures were washed and fixed with 4% paraformaldehyde (PFA) for 20 min at room temperature. Permeabilization was performed in 1% (w/v) BSA, 10% (v/v) donkey serum and 0.5% (v/v) Tween-20 in PBS. Cells were labelled with anti-FOXO1 (Cell Signaling Technology, 2880; 1:100), anti-PECAM-1/CD31 (R&D Biosystems, AF3628; 1:200) and 4,6-diamidino-2-phenylindole (DAPI; 1 µg ml⁻¹) diluted in incubation buffer (0.5% BSA, 5% donkey serum and 0.25% Tween-20 in PBS). Following washes in PBS with 0.1% (v/v) Tween-20 (PBST), HUVECs were incubated with Alexa-Fluor-conjugated secondary antibodies (Invitrogen; 1:1,000) diluted in incubation buffer for 2 h at room temperature. Cells were washed in PBST, mounted in VectaShield (Vector Laboratories, H-1000) and confocal images acquired using a SP8 confocal microscope (Leica). Labelling of proliferating cells with EdU was performed using a Click-iT EdU imaging kit (Invitrogen). HUVECs were pulsed with 10 µM EdU in culture medium for 3 h before collection and further processed according to the manufacturer’s instructions.

Metabolomics analyses

Untargeted biochemical profiling of intracellular metabolites was measured using gas chromatography (GC)–MS and LC–MS/MS platforms from Metabolon. Samples were normalized using the cell number and rescaled to set the median to 1. Missing values were imputed with the minimum value, and analyses performed with hierarchical clustering and PCA. For the analysis of 2HG in HUVECs transduced with the doxycycline-inducible lentiviruses, cells were seeded onto 60-mm dishes and treated with 100 ng ml⁻¹ doxycycline (Sigma) for 48 h before collection. For studies using sparse and dense cultures, HUVECs were seeded onto 60-mm dishes and cultured in EBM for 10 days (dense cultures). Culture medium was regularly replaced every 2 days. To generate sparse cultures, dense cultures at day 8 were trypsinized and re-seeded to reinitiate proliferation 36 h before metabolite extraction. After the indicated experimental treatments, HUVECs were washed three times with ice-cold PBS (Life Technologies), and metabolites were collected with extraction buffer (50% (v/v) methanol, 30% (v/v) acetonitrile, 20% (v/v) ultrapure water and 100 ng ml⁻¹ HEPES). Thereafter, plates were incubated for 15 min over dry ice and methanol, samples collected by scraping onto Eppendorf tubes and incubated on dry ice for an additional 15 min. Tubes were incubated at 4 °C and shaken at 1,400 r.p.m. for 15 min, and metabolite extracts were cleared by centrifugation at 14,000 r.p.m., 4 °C for 10 min. Finally, the supernatants were transferred into autosampler vials and stored at −80 °C before analysis. LC–MS analysis of sample extracts was performed on a QExactive Orbitrap mass spectrometer coupled to a Dionex UltiMate 3000 Rapid Separation LC system (Thermo). The LC system was fitted with a SeQuant Zic-pHILIC (150 × 2.1 mm, 5 μm) with a guard column (20 × 2.1 mm, 5 μm) from Merck. The flow rate was set at 200 µL min⁻¹ with the following gradient: 0 min 80% B, 2 min 80% B, 17 min 20% B, 17.1 min 80% B, and a hold at 80% B for 5 min. The mass spectrometer was operated in full MS and polarity-switching mode. Five independent cell cultures were measured for each condition, and samples were randomized to avoid bias in sample analyses due to machine drift. The acquired spectra were analysed using XCalibur Qual Browser and XCalibur Quan Browser software (Thermo Scientific) by referencing to an internal library of compounds. The R packages muma and gplots were used for data visualization. Quantification of 2HG was performed via interpolation of the corresponding standard curve obtained from serial dilutions of dl-α-hydroxyglutaric acid disodium salt (Sigma, 94577) running with the same batch of samples. Chiral derivatization of 2HG metabolite extracts with (+)-O,O′-diacetyl-l-tartaric anhydride (DATAN; Sigma, 358924) was performed as previously described^38,60, with some modifications. Briefly, for each sample, 300 μl of metabolite extract was evaporated to dryness at ambient temperature in a Savant SpeedVac concentrator (Thermo Scientific). The residue was resuspended in 75 μl of freshly prepared mixture of 80% acetonitrile/20% acetic acid (v/v) plus 50 mg ml⁻¹ DATAN (Acros Organics) and incubated at 75 °C for 45 min. Samples were cooled to room temperature and centrifuged before the addition of 75 μl 80% acetonitrile/20% acetic acid. The derivatized sample (5 μl) was injected into a Acquity UHPLC HSS T3 column (100 × 2.1 mm, 1.8-μm particle size). A Dionex UltiMate 3000 Rapid Separation LC was coupled to a QExactive Orbitrap mass spectrometer, and a gradient elution of 1.5 mM ammonium formate in water (mobile phase A, pH 3.6 adjusted with formic acid) and 0.1% formic acid in acetonitrile (mobile phase B) was used to separate the derivatized R- and S-2HG enantiomers. The target analytes were monitored by in parallel-reaction monitoring in negative electrospray ionization mode. The transition (precursor ion → product ion) of m/z 363 → 147 for derivatized R- and S-2HG was monitored, and an accurate mass of 147.03 was extracted using XCalibur Qual Browser and XCalibur Quan Browser software (Thermo Scientific) and used for relative quantification. Experimental samples were randomized to avoid bias in sample analyses due to machine drift.

Metabolic assays

The OCR was measured using a Seahorse XFe96 analyser (Seahorse Bioscience). In brief, HUVECs (40,000 cells per well) were seeded onto fibronectin-coated XFe96 microplates. After 4 h, the medium was changed to a non-buffered assay medium and cells were maintained in a non-CO₂ incubator for 1 h. Using a Mito Stress test kit (Seahorse Bioscience), the OCR was measured sequentially under basal conditions, after injection of oligomycin (3 µM), the mitochondrial uncoupler carbonyl cyanide-4-(trifluoromethoxy)phenyl-hydrazone (FCCP; 1 µM) and after injection of a mix of the respiratory chain inhibitors antimycin A (1.5 µM) and rotenone (3 µM), as previously described⁴. OGDH activity and ATP levels were measured in HUVEC lysates (1 × 10⁶ cells) using a OGDH activity assay kit (Sigma) and an ATP Bioluminescence Assay kit CLS II (Roche), respectively.

RNA-seq and GSEA

For RNA-seq, RNA was isolated from HUVECs using a miRNeasy micro kit (Qiagen) combined with on-column DNase digestion (DNase-Free DNase Set, Qiagen). RNA and library preparation integrity were verified using a LabChip Gx Touch 24 (Perkin Elmer). For input, 4 μg of total RNA was used for Truseq Stranded mRNA library preparation following the low-sample protocol (Illumina). Sequencing was performed on a NextSeq500 instrument (Illumina) using v2 chemistry, resulting in a minimum of 32 million reads per library with 1 × 75-bp single-end setup. Processing of raw reads was performed as previously described⁶¹. For GSEA⁶² and Gene Ontology terms analysis, gene set collections from the Molecular Signatures Database (MSigDB) 4.0 (http://www.broadinstitute.org/gsea/msigdb/) were used. Heatmaps were generated using Morpheus (https://software.broadinstitute.org/morpheus/).

RT–qPCR

RNA isolation, complementary DNA synthesis and quantitative PCR with reverse transcription (RT–qPCR) were performed as previously described⁴. The human TaqMan Gene Expression assays used are listed in Supplementary Table 6.

ChIP assay

HUVECs were fixed with 1% formaldehyde for 15 min and quenched with 0.125 M glycine. Chromatin was isolated via the addition of lysis buffer, followed by disruption with a Dounce homogenizer. Lysates were sonicated, and the DNA sheared to an average length of 300–500 bp. Genomic DNA (input) was prepared by treating aliquots of chromatin with RNase, proteinase K and heat for reverse-crosslinking, followed by ethanol precipitation. Pellets were resuspended and the resulting DNA was quantified on a NanoDrop spectrophotometer. Extrapolation to the original chromatin volume allowed quantitation of the total chromatin yield. Sheared chromatin (30 μg) was precleared with protein A agarose beads (Invitrogen). Genomic DNA regions of interest were isolated using 4 μg of ChIP-grade antibodies against FOXO1 (Abcam, ab39670), H3K4me3 (Active Motif, 39159) and H3K27ac (Active Motif, 39133). Complexes were washed, eluted from the beads with SDS buffer and subjected to RNase and proteinase K treatment. Crosslinks were reversed by incubation overnight at 65 °C, and ChIP DNAs were purified by phenol–chloroform extraction and ethanol precipitation.

ChIP-seq

Illumina sequencing libraries were prepared from ChIP and input DNAs by the standard consecutive enzymatic steps of end-polishing, dA-addition and adaptor ligation. After a final PCR amplification step, the resulting DNA libraries were quantified and sequenced on Illumina’s NextSeq 500 (75-nt reads, single end). Reads were aligned to the human genome (hg38) using the Burrows–Wheeler algorithm⁶³ (default settings). Duplicate reads were removed, and only uniquely mapped reads (mapping quality ≥ 25) were used for further analysis. Alignments were extended in silico at their 3′-ends to a length of 200 bp, which is the average genomic fragment length in the size-selected library, and assigned to 32-nt bins along the genome. The resulting histograms (genomic ‘signal maps’) were stored in bigWig files. Peak locations were determined using the MACS algorithm (v.2.1.0)⁶⁴ with a cut-off of P value of 1 × 10⁻⁷. Peaks that were on the ENCODE blacklist of known false ChIP-seq peaks were removed. Signal maps and peak locations were used as input data to Active Motifs proprietary analysis program, which creates tables containing detailed information on sample comparison, peak metrics, peak locations and gene annotations. Binding motifs were identified with the findMotifsGenome program of the HOMER package⁶⁵ using default parameters and input sequences comprising ±100 bp from the centre of the top 1,000 peaks. Individual profiles were produced with a window of 5 bp. All profiles were plotted on a normalized reads-per-million basis. The processed data were plotted and visualized using software of the R project for statistical computing.

Animals, genetic experiments and intraocular injections

The conditional Ogdh knockout allele was generated by flanking exons 3 and 4 with loxP sites. Loss of exons 3 and 4 after Cre-mediated recombination results in a frameshift of the Ogdh gene. Mice were on a C57BL/6 genetic background and were generated by Cyagen Biosciences. For constitutive Cre-mediated recombination in ECs, Ogdh^fl/fl mice (Ogdh^EC-KO) were bred with Tie2-cre transgenic mice⁶⁶. To avoid recombination in the female germline, only Tie2-cre-positive male mice were used for intercrosses. Embryos were collected from cre-negative females at the indicated time points and genotyping performed from isolated embryos. For inducible Cre-mediated recombination in ECs, loxP-flanked Ogdh mice (Ogdh^fl/fl) were bred with transgenic mice expressing the tamoxifen-inducible, Cdh5 promoter-driven creERT2 (ref. ⁶⁷) (Ogdh^iEC-KO). For the analysis of angiogenesis in the postnatal mouse retina, Cre-mediated recombination was induced by intraperitoneal injections of 25 µl 4-OHT (2 mg ml⁻¹; Sigma) from P1 to P4. Eyes were collected at P6 for analysis. Control animals were littermate animals without Cre. Mice were genotyped by PCR performed on genomic DNA. Genomic DNA was extracted from tail biopsies using DirectPCR Lysis reagent (Peqlab). Intraocular administration of S-2HG (800 µM) and KMV (10 mM) into newborn mouse retinas was performed as previously described⁶⁸. Briefly, a single dose of metabolites (0.25 µl) was injected into the vitreous cavity of P5 mice using a Nanoliter 2000 microinjector (World Precision Instruments). As controls, all pups were injected with S-2HG or KMV into the one eyeball and with the vehicle (DMSO or PBS, respectively) into the contralateral eye. Eyes were collected at P7 for further analysis by immunohistochemistry. Male and female mice (C57BL/6) were used for this experiment. All mice were maintained under specific pathogen-free conditions, and animal experiments were conducted in accordance with institutional guidelines and laws, following protocols approved by local animal ethics committees and authorities. The genetic experiments were approved by the Regierungspraesidium Darmstadt and the intraocular injections were performed under approval from the Institutional Animal Care and Use Committee of the Korea Advanced Institute of Science and Technology.

Immunohistochemistry of murine retinal and yolk sac vasculature

To analyse blood vessel growth in the postnatal retina, whole mouse eyes were fixed in 4% (w/v) PFA on ice for 2.5 h. Eyes were washed in PBS and retinas were dissected and partially cut into four leaflets. After blocking and permeabilization in PBS containing 2% (v/v) FCS, 1% (w/v) BSA and 0.5% (v/v) Tween-20 for 1 h at room temperature, retinas were incubated overnight at 4 °C with the primary antibody diluted in PBS containing 1% (v/v) FCS, 0.5% (w/v) BSA, 0.25% (v/v) Tween-20 and 0.25% (v/v) Triton X-100. Primary antibodies against the following proteins were used: collagen IV (Bio-Rad, 2150-1470; 1:400); cleaved (Asp175)-caspase3 (Cell Signaling Technology, 9664; 1:100); ERG (Abcam, ab92513; 1:200); ICAM2 (BD Biosciences, 553326; 1:200); and PECAM-1/CD31 (R&D Biosystems, AF3628; 1:200). After four washes in PBST, retinas were incubated with Alexa-Fluor 488-, Alexa-Fluor 555-, Alexa-Fluor 594- or Alexa-Fluor 647-conjugated secondary antibodies (Life Technologies; 1:400) diluted in blocking buffer for 2 h at room temperature. After four washes in PBST, retinas were flat-mounted with ProLong Gold Antifade reagent (Life Technologies) and imaged by high-resolution confocal microscopy (Leica SP8). Labelling of proliferating cells with EdU was performed using a Click-iT EdU imaging kit (Invitrogen). Briefly, mouse pups were intraperitoneally injected with 25 µl of EdU (6 mg ml⁻¹; Invitrogen) for 3 h before retinal collection, and EdU labelling was performed according to the manufacturer’s instructions.

For the analysis of the murine yolk sac vasculature, yolk sacs were flat-pinned on a silicon plate and fixed in 4% PFA on ice for 1 h followed by two washes in PBST. After blocking in 2% (v/v) donkey serum, 0.2% (w/v) BSA in PBST for 2 h at room temperature, the yolk sacs were incubated overnight at 4 °C with the indicated primary antibody diluted in blocking buffer. After five washes in PBST, yolk sacs were incubated with Alexa-Fluor-conjugated secondary antibodies for 2 h at room temperature. Samples were washed four times with PBST, flat-mounted in ProLong Gold Antifade and imaged by confocal microscopy.

Immunostainings were carried out in tissues from littermates and processed under the same conditions. Volocity (Perkin Elmer), Fiji/ImageJ, Photoshop (Adobe) and Illustrator (Adobe) software were used for image acquisition and processing. For all of the images in which the levels of immunostaining were compared, settings for laser excitation and confocal scanner detection were kept constant between groups.

Quantitative analysis of the vasculature

All quantifications were done on high-resolution confocal images of thin z-sections of the sample using Volocity (Perkin Elmer) software. Endothelial coverage and the number of endothelial branch points were quantified behind the angiogenic front in a region between an artery and a vein. All parameters were quantified from a minimum of three vascularized fields per sample. Endothelial coverage was determined by assessing the ratio of the PECAM-positive area to the total area of the vascularized field (400 × 200 μm for murine retinas and 200 × 200 μm for yolk sac vasculature) and expressed as the percentage of the area covered by PECAM-positive ECs. The number of ECs (ERG-positive cells per field), the rate of proliferating ECs (EdU/ERG double-positive cells normalized to the number of ERG-positive cells) or vessel regression (number of ICAM2-negative, collagen-IV-positive basement membrane sleeves per 1,000-μm vessel segment) were also quantified in the same fields. The number of filopodial extensions was quantified at the angiogenic front as previously described⁶⁹. All images shown are representative of the vascular phenotype observed in samples from at least three distinct litters per group.

Statistics and reproducibility

Statistical analysis was performed using unpaired, two-tailed Student’s t-test unless mentioned otherwise. For all bar graphs, data are represented as the mean ± s.e.m. P values of <0.05 were considered significant. All calculations were performed using Prism 8.0 (GraphPad Software).

No randomization or blinding was used for the analyses. Images are representative of at least three independent experiments in mice or cells of the same treatment group or genotype. Western blot data were from the respective experiment, processed in parallel and are representative of at least three independent experiments. Sample sizes were selected on the basis of published protocols⁶⁹ and previous experiments.

Reporting Summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

Deep-sequencing (ChIP-seq and RNA–seq) data that support the findings of this study have been deposited in the Gene Expression Omnibus (GEO) under accession code GSE128636. All other data supporting the findings of this study are available from the corresponding author upon reasonable request. Source data are provided with this paper.

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Acknowledgements

Research in the M.P. laboratory was supported by the Max Planck Society, the European Research Council (ERC) Consolidator Grant EMERGE (773047), the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) – Project-ID 75732319 – SFB 834, the Leducq Foundation, the European Union’s Horizon 2020 research and innovation programme under the Marie Sklodowska-Curie action (814316), the Excellence Cluster Cardio-Pulmonary Institute (EXC 2026, project ID 390649896), the DZHK (German Centre for Cardiovascular Research), the Stiftung Charité, and the European Molecular Biology Organization (EMBO) Young Investigator Programme. Work in the T.B. laboratory was supported by the DFG – Project-ID 268555672 – SFB 1213. Work in the C.F. laboratory was supported by the Medical Research Council (MRC_MC_UU_12022/6). This work was performed with assistance from the CSHL Mass Spectrometry Shared Resource, which is supported by the Cancer Center Support Grant 5P30CA045508.

Author information

Ana S. H. Costa
Present address: Cold Spring Harbor Laboratory, Cold Spring Harbor, NY, USA
Jaeryung Kim
Present address: Department of Oncology and Ludwig Institute for Cancer Research, University of Lausanne and Centre Hospitalier Universitaire Vaudois, Epalinges, Switzerland
These authors contributed equally: Jorge Andrade, Chenyue Shi, Ana S. H. Costa.

Authors and Affiliations

Angiogenesis and Metabolism Laboratory, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
Jorge Andrade, Chenyue Shi, Anuradha Doddaballapur, Toshiya Sugino, Yu Ting Ong, Marco Castro, Barbara Zimmermann, Kerstin Wilhelm & Michael Potente
Medical Research Council Cancer Unit, University of Cambridge, Cambridge, UK
Ana S. H. Costa & Christian Frezza
Graduate School of Medical Science and Engineering, Korea Advanced Institute of Science and Technology (KAIST), Daejeon, Korea
Jeongwoon Choi & Gou Young Koh
Center for Vascular Research, Institute for Basic Science (IBS), Daejeon, Korea
Jeongwoon Choi, Jaeryung Kim & Gou Young Koh
Gene Editing Group, Institute of Biochemistry II, Goethe University, Frankfurt (Main), Germany
Manuel Kaulich
Department of Cardiac Development and Remodeling, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
Stefan Guenther & Thomas Braun
Department of Anatomy, Keio University School of Medicine, Tokyo, Japan
Yoshiaki Kubota
UCIBIO–Unidade de Ciências Biomoleculares Aplicadas, Departamento Ciências da Vida, Faculdade de Ciências e Tecnologia–Universidade Nova de Lisboa Campus de Caparica, Caparica, Portugal
Ana Rita Grosso
Instituto de Medicina Molecular, Faculdade de Medicina, Universidade de Lisboa, Lisboa, Portugal
Ana Rita Grosso
Berlin Institute of Health (BIH) at Charité–Universitätsmedizin Berlin, Berlin, Germany
Michael Potente
Max Delbrück Center for Molecular Medicine (MDC), Berlin, Germany
Michael Potente

Authors

Jorge Andrade
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Chenyue Shi
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Ana S. H. Costa
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Jeongwoon Choi
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Jaeryung Kim
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Anuradha Doddaballapur
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Toshiya Sugino
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Yu Ting Ong
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Marco Castro
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Barbara Zimmermann
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Manuel Kaulich
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Stefan Guenther
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Kerstin Wilhelm
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Yoshiaki Kubota
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Thomas Braun
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Gou Young Koh
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Ana Rita Grosso
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Christian Frezza
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Michael Potente
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Contributions

J.A., C.S., A.S.H.C., J.C., J.K., A.D., T.S., Y.T.O., M.C., B.Z. and S.G. performed experiments. J.C. and J.K. performed intraocular injections. J.A., C.S., A.S.H.C., J.C., J.K., A.D., T.S., Y.T.O., M.C., B.Z., S.G., A.R.G., C.F. and M.P. analysed data. A.S.H.C., J.C., J.K., M.K., K.W., Y.K., T.B., G.Y.K., A.R.G. and C.F. provided essential reagents and protocols. C.S., A.S.H.C. and C.F. contributed to manuscript writing and preparation. J.A. and M.P. wrote the paper. J.A. and M.P. conceived and directed the study. All authors discussed the results and commented on the manuscript.

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Correspondence to Michael Potente.

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The authors declare no competing interests.

Additional information

Peer review information Nature Cell Biology thanks Navdeep Chandel, Lena Claesson-Welsh and the other, anonymous, reviewer for their contribution to the peer review of this work.

a, Immunoblot analysis in AdCtrl and AdFOXO1^A3-transduced HUVECs showing the protein expression of the OGDH complex subunits. A FLAG antibody was used to validate the expression of the FLAG-tagged FOXO1^A3 mutant. Tubulin served as loading control. b, Immunoblot analysis of the BCAA metabolism enzymes ACADSB, MUT and DBT in AdCtrl and AdFOXO1^A3 transduced HUVECs. c, Immunoblot analysis of MYC protein levels in control and MYC-depleted HUVECs (gMYC). Cells were generated by lentiviral transduction with FLAG-tagged Cas9, control (gCtrl) or MYC (gMYC) targeting gRNAs. d, Gene set enrichment analysis (GSEA) showing a suppression of MYC-regulated genes in gMYC HUVECs when compared to controls (gCtrl). ES, enrichment score; NES, normalized enrichment score; FDR, false discovery rate. e, Expression of canonical MYC target genes and FOXO1-regulated BCAA genes in gCtrl and gMYC HUVECs. Analysis was performed by RNA-seq, (n=3 independent samples). Western blot data in a-c were from the respective experiment, processed in parallel, and are representative of at least three independent experiments. Data in e represent mean ± s.e.m.; a two-tailed unpaired t-test was used unless otherwise indicated; *P<0.05; **P< 0.01; ***P<0.001; ****P<0.0001; NS, not significant. The numerical data, unprocessed western blots and P values are provided as source data.

Source data

Extended Data Fig. 9 Identification of FOXO1 target genes by ChIP-seq.

a, Genome-wide analysis of FOXO1 binding peaks revealed the FOXO1 high-affinity binding sequence as the most enriched motif in the immunoprecipitated chromatin of FOXO1^A3-transduced HUVECs. b, Distribution of FOXO1 bindings peaks relative to the transcriptional start site (TSS), showing preferential binding of FOXO1 to genomic regions located at gene promoters near the TSS. c-f, ChIP-seq signals for FOXO1, H3K27Ac and H3K4me3 at the genomic loci of (c) OXCT1, (d) PDK1, (e) PDK4 and (f) MXI1. The FOXO consensus motifs that are bound by FOXO1 are highlighted in orange. ChIP-seq signals are represented as reads per kilobase million (RPKMs).

Extended Data Fig. 10 Genetic PHD inactivation lowers the proliferative activity of ECs.

a, Immunoblot analysis of PHD1/2/3-depleted HUVECs showing regulation of HIF targets and proliferation-associated proteins. Cells were generated by lentiviral transduction with FLAG-tagged Cas9, control- (gCtrl) or PHD1/2/3-targeting gRNAs (gPHD1/2/3). b, EdU-incorporation in PHD1/2/3-depleted HUVECs. DAPI was used to identify ECs nuclei. c, Quantification of EdU-incorporation in gCtrl and gPHD1/2/3-transduced HUVECs. Values represent the percentage of EdU-labelled ECs, (n=20, 24 independent samples for gCtrl and gPHD1/2/3). d, Reduced DNA synthesis in PHD1/2/3-depleted ECs, as assessed by analysing ³H-thymidine incorporation. Values are represented as fold-change relative to control, (n=6 independent samples). Western blot data in a were from the respective experiment, processed in parallel, and are representative of at least three independent experiments. c and d represent mean ± s.e.m.; a two-tailed unpaired t-test was used; **P< 0.01; ****P<0.0001. The numerical data, unprocessed western blots and P values are provided as source data.

Source data

Supplementary information

Reporting Summary

Supplementary Tables

Supplementary Table 1: metabolite changes induced by FOXO1 activation in HUVECs. Supplementary Table 2: genome-wide transcriptome analysis comparing control and S-2HG-treated HUVECs for 24 h. Supplementary Table 3: genomic localization of FOXO1-binding sites at the genomic loci of FOXO1 target genes. Supplementary Table 4: gene-specific gRNA sequences used for CRISPR–Cas9 genome-editing studies in HUVECs. Supplementary Table 5: antibodies used for western blot studies. Supplementary Table 6: human TaqMan Gene Expression assays used for RT–qPCR analyses.

Supplementary Video 1

Time-lapse image sequences of HUVECs transduced with the FUCCI reporter comparing the cell cycle dynamics in control ECs.

Supplementary Video 2

Time-lapse image sequences of HUVECs transduced with the FUCCI reporter comparing the cell cycle dynamics in S-2HG-treated ECs.

Supplementary Video 3

Scratch-wound assay showing the migratory behaviour of control ECs.

Supplementary Video 4

Scratch-wound assay showing the migratory behaviour of S-2HG-treated ECs.

Unprocessed western blots.

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Andrade, J., Shi, C., Costa, A.S.H. et al. Control of endothelial quiescence by FOXO-regulated metabolites. Nat Cell Biol 23, 413–423 (2021). https://doi.org/10.1038/s41556-021-00637-6

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Received: 05 February 2020
Accepted: 21 January 2021
Published: 01 April 2021
Issue Date: April 2021
DOI: https://doi.org/10.1038/s41556-021-00637-6

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Abstract

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Main

Results

FOXO1 activation induces 2-hydroxyglutarate generation in ECs

FOXO1-activated ECs produce S-2HG, which inhibits 2OG-dependent dioxygenases

S-2HG promotes a quiescent endothelial state

S-2HG limits the angiogenic behaviour of ECs

FOXO1 induces S-2HG generation by inhibiting the OGDH enzyme

Endothelial OGDH is crucial for angiogenic growth

FOXO1 inhibits OGDH function via the generation of BCAA catabolites

Genes involved in BCAA metabolism are direct FOXO1 target genes

BCAA metabolites restrict endothelial proliferation and vascular growth

Discussion

Methods

Cells and cell culture

Cell treatments

Adenoviral transductions

Lentiviruses and lentiviral transductions

CRISPR–Cas9 genome editing of HUVECs

Proliferation assays

RNA and protein synthesis assays

Cell cycle analysis by flow cytometry and live-cell imaging

Western blotting

Spheroid assay

Scratch-wound assay

Senescence-associated β-galactosidase staining

Immunohistochemistry of cell cultures

Metabolomics analyses

Metabolic assays

RNA-seq and GSEA

RT–qPCR

ChIP assay

ChIP-seq

Animals, genetic experiments and intraocular injections

Immunohistochemistry of murine retinal and yolk sac vasculature

Quantitative analysis of the vasculature

Statistics and reproducibility

Reporting Summary

Data availability

References

Acknowledgements

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Authors and Affiliations

Contributions

Corresponding author

Ethics declarations

Competing interests

Additional information

Extended data

Supplementary information

Source data

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