Folates enable the activation and transfer of one-carbon units for the biosynthesis of purines, thymidine and methionine1,2,3. Antifolates are important immunosuppressive4 and anticancer agents5. In proliferating lymphocytes6 and human cancers7,8, mitochondrial folate enzymes are particularly strongly upregulated. This in part reflects the need for mitochondria to generate one-carbon units and export them to the cytosol for anabolic metabolism2,9. The full range of uses of folate-bound one-carbon units in the mitochondrial compartment itself, however, has not been thoroughly explored. Here we show that loss of the catalytic activity of the mitochondrial folate enzyme serine hydroxymethyltransferase 2 (SHMT2), but not of other folate enzymes, leads to defective oxidative phosphorylation in human cells due to impaired mitochondrial translation. We find that SHMT2, presumably by generating mitochondrial 5,10-methylenetetrahydrofolate, provides methyl donors to produce the taurinomethyluridine base at the wobble position of select mitochondrial tRNAs. Mitochondrial ribosome profiling in SHMT2-knockout human cells reveals that the lack of this modified base causes defective translation, with preferential mitochondrial ribosome stalling at certain lysine (AAG) and leucine (UUG) codons. This results in the impaired expression of respiratory chain enzymes. Stalling at these specific codons also occurs in certain inborn errors of mitochondrial metabolism. Disruption of whole-cell folate metabolism, by either folate deficiency or antifolate treatment, also impairs the respiratory chain. In summary, mammalian mitochondria use folate-bound one-carbon units to methylate tRNA, and this modification is required for mitochondrial translation and thus oxidative phosphorylation.
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Fox, J. T. & Stover, P. J. Folate-mediated one-carbon metabolism. Vitam. Horm. 79, 1–44 (2008)
Tibbetts, A. S. & Appling, D. R. Compartmentalization of mammalian folate-mediated one-carbon metabolism. Annu. Rev. Nutr. 30, 57–81 (2010)
Ducker, G. S. & Rabinowitz, J. D. One-carbon metabolism in health and disease. Cell Metab. 25, 27–42 (2017)
Lipsky, P. E. et al. Infliximab and methotrexate in the treatment of rheumatoid arthritis. N. Engl. J. Med. 343, 1594–1602 (2000)
Chabner, B. A. & Roberts, T. G. Jr. Timeline: chemotherapy and the war on cancer. Nat. Rev. Cancer 5, 65–72 (2005)
Ron-Harel, N. et al. Mitochondrial biogenesis and proteome remodeling promote one-carbon metabolism for T cell activation. Cell Metab. 24, 104–117 (2016)
Nilsson, R. et al. Metabolic enzyme expression highlights a key role for MTHFD2 and the mitochondrial folate pathway in cancer. Nat. Commun. 5, 3128 (2014)
Kim, D. et al. SHMT2 drives glioma cell survival in ischaemia but imposes a dependence on glycine clearance. Nature 520, 363–367 (2015)
Ducker, G. S. et al. Reversal of cytosolic one-carbon flux compensates for loss of the mitochondrial folate pathway. Cell Metab. 23, 1140–1153 (2016)
Garrow, T. A. et al. Cloning of human cDNAs encoding mitochondrial and cytosolic serine hydroxymethyltransferases and chromosomal localization. J. Biol. Chem. 268, 11910–11916 (1993)
Gohil, V. M. et al. Nutrient-sensitized screening for drugs that shift energy metabolism from mitochondrial respiration to glycolysis. Nat. Biotechnol. 28, 249–255 (2010)
Mullen, A. R. et al. Reductive carboxylation supports growth in tumour cells with defective mitochondria. Nature 481, 385–388 (2011)
Sullivan, L. B. et al. Supporting aspartate biosynthesis is an essential function of respiration in proliferating cells. Cell 162, 552–563 (2015)
Birsoy, K. et al. An essential role of the mitochondrial electron transport chain in cell proliferation is to enable aspartate synthesis. Cell 162, 540–551 (2015)
Iborra, F. J., Kimura, H. & Cook, P. R. The functional organization of mitochondrial genomes in human cells. BMC Biol. 2, 9 (2004)
Brown, S. S., Neal, G. E. & Williams, D. C. Subcellular distribution of some folic acid-linked enzymes in rat liver. Biochem. J. 97, 34C–36C (1965)
Anderson, D. D., Quintero, C. M. & Stover, P. J. Identification of a de novo thymidylate biosynthesis pathway in mammalian mitochondria. Proc. Natl Acad. Sci. USA 108, 15163–15168 (2011)
Kozak, M. Comparison of initiation of protein synthesis in procaryotes, eucaryotes, and organelles. Microbiol. Rev. 47, 1–45 (1983)
Tucker, E. J. et al. Mutations in MTFMT underlie a human disorder of formylation causing impaired mitochondrial translation. Cell Metab. 14, 428–434 (2011)
Saada, A. et al. Mutant mitochondrial thymidine kinase in mitochondrial DNA depletion myopathy. Nat. Genet. 29, 342–344 (2001)
Calvo, S. E. & Mootha, V. K. The mitochondrial proteome and human disease. Annu. Rev. Genomics Hum. Genet. 11, 25–44 (2010)
Agris, P. F., Vendeix, F. A. & Graham, W. D. tRNA’s wobble decoding of the genome: 40 years of modification. J. Mol. Biol. 366, 1–13 (2007)
Van Haute, L. et al. Deficient methylation and formylation of mt-tRNAMet wobble cytosine in a patient carrying mutations in NSUN3. Nat. Commun. 7, 12039 (2016)
Pütz, J., Dupuis, B., Sissler, M. & Florentz, C. Mamit-tRNA, a database of mammalian mitochondrial tRNA primary and secondary structures. RNA 13, 1184–1190 (2007)
Fu, Y. et al. The AlkB domain of mammalian ABH8 catalyzes hydroxylation of 5-methoxycarbonylmethyluridine at the wobble position of tRNA. Angew. Chem. Int. Ed. Engl. 49, 8885–8888 (2010)
Songe-Møller, L. et al. Mammalian ALKBH8 possesses tRNA methyltransferase activity required for the biogenesis of multiple wobble uridine modifications implicated in translational decoding. Mol. Cell. Biol. 30, 1814–1827 (2010)
Yasukawa, T. et al. Defect in modification at the anticodon wobble nucleotide of mitochondrial tRNALys with the MERRF encephalomyopathy pathogenic mutation. FEBS Lett. 467, 175–178 (2000)
Yasukawa, T., Suzuki, T., Ueda, T., Ohta, S. & Watanabe, K. Modification defect at anticodon wobble nucleotide of mitochondrial tRNAsLeu(UUR) with pathogenic mutations of mitochondrial myopathy, encephalopathy, lactic acidosis, and stroke-like episodes. J. Biol. Chem. 275, 4251–4257 (2000)
Suzuki, T. & Suzuki, T. A complete landscape of post-transcriptional modifications in mammalian mitochondrial tRNAs. Nucleic Acids Res. 42, 7346–7357 (2014)
Ghezzi, D. et al. Mutations of the mitochondrial-tRNA modifier MTO1 cause hypertrophic cardiomyopathy and lactic acidosis. Am. J. Hum. Genet. 90, 1079–1087 (2012)
Kopajtich, R. et al. Mutations in GTPBP3 cause a mitochondrial translation defect associated with hypertrophic cardiomyopathy, lactic acidosis, and encephalopathy. Am. J. Hum. Genet. 95, 708–720 (2014)
Moukadiri, I. et al. Evolutionarily conserved proteins MnmE and GidA catalyze the formation of two methyluridine derivatives at tRNA wobble positions. Nucleic Acids Res. 37, 7177–7193 (2009)
Doherty, E. A., Batey, R. T., Masquida, B. & Doudna, J. A. A universal mode of helix packing in RNA. Nat. Struct. Biol. 8, 339–343 (2001)
Rouskin, S., Zubradt, M., Washietl, S., Kellis, M. & Weissman, J. S. Genome-wide probing of RNA structure reveals active unfolding of mRNA structures in vivo. Nature 505, 701–705 (2014)
Kirino, Y., Goto, Y., Campos, Y., Arenas, J. & Suzuki, T. Specific correlation between the wobble modification deficiency in mutant tRNAs and the clinical features of a human mitochondrial disease. Proc. Natl Acad. Sci. USA 102, 7127–7132 (2005)
Grim, J., Chládek, J. & Martínková, J. Pharmacokinetics and pharmacodynamics of methotrexate in non-neoplastic diseases. Clin. Pharmacokinet. 42, 139–151 (2003)
Mayr, J. A. et al. Spectrum of combined respiratory chain defects. J. Inherit. Metab. Dis. 38, 629–640 (2015)
Ran, F. A. et al. Genome engineering using the CRISPR–Cas9 system. Nat. Protocols 8, 2281–2308 (2013)
Szebenyi, D. M ., Musayev, F. N ., di Salvo, M. L ., Safo, M. K. & Schirch, V. Serine hydroxymethyltransferase: role of glu75 and evidence that serine is cleaved by a retroaldol mechanism. Biochemistry 43, 6865–6876 (2004)
Contestabile, R. et al. Role of tyrosine 65 in the mechanism of serine hydroxymethyltransferase. Biochemistry 39, 7492–7500 (2000)
Iurescia, S., Condò, I., Angelaccio, S., Delle Fratte, S. & Bossa, F. Site-directed mutagenesis techniques in the study of Escherichia coli serine hydroxymethyltransferase. Protein Expr. Purif. 7, 323–328 (1996)
Tischner, C. et al. MTO1 mediates tissue specificity of OXPHOS defects via tRNA modification and translation optimization, which can be bypassed by dietary intervention. Hum. Mol. Genet. 24, 2247–2266 (2015)
Sasarman, F. & Shoubridge, E. A. Radioactive labeling of mitochondrial translation products in cultured cells. Methods Mol. Biol. 837, 207–217 (2012)
Clasquin, M. F., Melamud, E. & Rabinowitz, J. D. LC–MS data processing with MAVEN: a metabolomic analysis and visualization engine. Curr. Protoc. Bioinformatics Chapter 14, Unit14.11 (2012)
Afgan, E. et al. The Galaxy platform for accessible, reproducible and collaborative biomedical analyses: 2016 update. Nucleic Acids Res. 44 (W1), W3–W10 (2016)
R Development Core Team. R: A Language and Environment for Statistical Computing; http://www.R-project.org/ (Vienna, Austria, 2016)
Marcel, M. Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet.journal 17, 10–12 (2011)
Kim, D. et al. TopHat2: accurate alignment of transcriptomes in the presence of insertions, deletions and gene fusions. Genome Biol. 14, R36 (2013)
Anders, S., Pyl, P. T. & Huber, W. HTSeq—a Python framework to work with high-throughput sequencing data. Bioinformatics 31, 166–169 (2015)
Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550 (2014)
Bao, X. R. et al. Mitochondrial dysfunction remodels one-carbon metabolism in human cells. eLife 5, e10575 (2016)
Mayr, J. A. et al. Mitochondrial phosphate-carrier deficiency: a novel disorder of oxidative phosphorylation. Am. J. Hum. Genet. 80, 478–484 (2007)
Langmead, B. & Salzberg, S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357–359 (2012)
Ramírez, F. et al. deepTools2: a next generation web server for deep-sequencing data analysis. Nucleic Acids Res. 44 (W1), W160–W165 (2016)
Garrison, E. & Marth, G. Haplotype-based variant detection from short-read sequencing. Preprint at https://arxiv.org/abs/1207.3907 (2012)
Hahne, F. & Ivanek, R. Visualizing genomic data using Gviz and Bioconductor. Methods Mol. Biol. 1418, 335–351 (2016)
Ingolia, N. T., Brar, G. A., Rouskin, S., McGeachy, A. M. & Weissman, J. S. The ribosome profiling strategy for monitoring translation in vivo by deep sequencing of ribosome-protected mRNA fragments. Nat. Protocols 7, 1534–1550 (2012)
Rooijers, K., Loayza-Puch, F., Nijtmans, L. G. & Agami, R. Ribosome profiling reveals features of normal and disease-associated mitochondrial translation. Nat. Commun. 4, 2886 (2013)
Couvillion, M. T., Soto, I. C., Shipkovenska, G. & Churchman, L. S. Synchronized mitochondrial and cytosolic translation programs. Nature 533, 499–503 (2016)
Dunn, J. G. plastid: a positional library for sequencing analysis; http://plastid.readthedocs.io (2016)
Nakahigashi, K. et al. Effect of codon adaptation on codon-level and gene-level translation efficiency in vivo. BMC Genomics 15, 1115 (2014)
Balakrishnan, R., Oman, K., Shoji, S., Bundschuh, R. & Fredrick, K. The conserved GTPase LepA contributes mainly to translation initiation in Escherichia coli. Nucleic Acids Res. 42, 13370–13383 (2014)
Oh, E. et al. Selective ribosome profiling reveals the cotranslational chaperone action of trigger factor in vivo. Cell 147, 1295–1308 (2011)
Dunn, J. G., Foo, C. K., Belletier, N. G., Gavis, E. R. & Weissman, J. S. Ribosome profiling reveals pervasive and regulated stop codon readthrough in Drosophila melanogaster. eLife 2, e01179 (2013)
Wang, Y., Geer, L. Y., Chappey, C., Kans, J. A. & Bryant, S. H. Cn3D: sequence and structure views for Entrez. Trends Biochem. Sci. 25, 300–302 (2000)
Krogh, A., Larsson, B., von Heijne, G. & Sonnhammer, E. L. Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes. J. Mol. Biol. 305, 567–580 (2001)
Del Campo, C., Bartholomäus, A., Fedyunin, I. & Ignatova, Z. Secondary structure across the bacterial transcriptome reveals versatile roles in mRNA regulation and function. PLoS Genet. 11, e1005613 (2015)
Danson, M. J. & Hough, D. W. Citrate synthase from hyperthermophilic Archaea. Methods Enzymol. 331, 3–12 (2001)
Feichtinger, R. G. et al. Low aerobic mitochondrial energy metabolism in poorly- or undifferentiated neuroblastoma. BMC Cancer 10, 149 (2010)
Rustin, P. et al. Biochemical and molecular investigations in respiratory chain deficiencies. Clin. Chim. Acta 228, 35–51 (1994)
Clayton, D. A. & Shadel, G. S. Isolation of mitochondria from tissue culture cells. Cold Spring Harb. Protoc. https://doi.org/10.1101/pdb.prot080002 (2014)
Zheng, G. et al. ALKBH5 is a mammalian RNA demethylase that impacts RNA metabolism and mouse fertility. Mol. Cell 49, 18–29 (2013)
Ogata, T. et al. Chemical synthesis and properties of 5-taurinomethyluridine and 5-taurinomethyl-2-thiouridine. J. Org. Chem. 74, 2585–2588 (2009)
Shevchenko, A., Tomas, H., Havlis, J., Olsen, J. V. & Mann, M. In-gel digestion for mass spectrometric characterization of proteins and proteomes. Nat. Protocols 1, 2856–2860 (2006)
Dorfer, V. et al. MS Amanda, a universal identification algorithm optimized for high accuracy tandem mass spectra. J. Proteome Res. 13, 3679–3684 (2014)
Eng, J. K., McCormack, A. L. & Yates, J. R. An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database. J. Am. Soc. Mass Spectrom. 5, 976–989 (1994)
Bern, M., Kil, Y. J. & Becker, C. Byonic: advanced peptide and protein identification software. Curr. Protoc. Bioinformatics Chapter 13, Unit13.20 (2012)
Spivak, M., Weston, J., Bottou, L., Käll, L. & Noble, W. S. Improvements to the percolator algorithm for Peptide identification from shotgun proteomics data sets. J. Proteome Res. 8, 3737–3745 (2009)
MacLean, B. et al. Skyline: an open source document editor for creating and analyzing targeted proteomics experiments. Bioinformatics 26, 966–968 (2010)
Schilling, B. et al. Platform-independent and label-free quantitation of proteomic data using MS1 extracted ion chromatograms in skyline: application to protein acetylation and phosphorylation. Mol. Cell. Proteomics 11, 202–214 (2012)
Renwick, S. B., Snell, K. & Baumann, U. The crystal structure of human cytosolic serine hydroxymethyltransferase: a target for cancer chemotherapy. Structure 6, 1105–1116 (1998)
We thank T. Pan, W. Lu, L. Chen, L. Parsons, W. Wang and T. Srikumar, and all members of the Rabinowitz laboratory. This work was supported by funding to J.D.R. from the US National Institutes of Health (NIH) (R01CA163591 and DP1DK113643) and StandUp to Cancer (SU2C-AACR-DT-20-16). G.S.D. was supported by a postdoctoral fellowship (PF-15-190-01- TBE) from the American Cancer Society. J.A.M. was supported by the science fund of the Paracelsus Medical University Salzburg (E-12/15/076-MAY). Z.G. was supported by the NIH (DP1AI124669).
J.D.R. is a founder of Raze Therapeutics, which aims to target 1C metabolism for cancer therapy.
Reviewer Information Nature thanks D. Appling, V. Gohil and the other anonymous reviewer(s) for their contribution to the peer review of this work.
Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended data figures and tables
Extended Data Figure 1 SHMT2 deletion-induced respiratory chain dysfunction in different cellular backgrounds and clones.
a, Change in media colour after 48 h cell growth. b, c, Lactate secretion (b) and normalized NAD+/NADH ratio (c) of HCT116 knockout cell lines (n = 6). d, e, Basal respiration as measured by Seahorse XF analyser (n = 3) (d) and normalized NAD+/NADH ratio (n = 3) (e) of HEK293T folate 1C gene CRISPR–Cas9 knockout cell lines. f, Normalized levels of TCA cycle and associated metabolites (n = 3). g, Steady-state labelling fraction into citrate from [U-13C]substrates glutamine (left) and glucose (right) (n = 3). h, Immunoblot of extracted mitochondria for subunits of respiratory chain complexes I–V (CI–CV) and markers of mitochondrial mass. i, Mitochondrial complex I levels (NDUFS4) in independent HCT116 folate 1C gene knockout clones. Data are mean ± s.e.m. n indicates the number of biological replicates. *P < 0.01, two-tailed Student’s t-test (see Supplementary Table 7 for exact P values).
a, Mapping of mutated amino acid residues on human SHMT1 (PDB code 1BJ481) using iCn3D and alignment of E. coli serine hydroxymethyltransferase (GLYA), H. sapiens mitochondrial serine hydroxymethyltransferase 2 (GLYM) and cytosolic serine hydroxymethyltransferase 1 (GLYC). Positions for GLYM are given with reference to GenBank NM_005412.5. b, Sanger sequencing traces of mutant constructs. c, Immunoblot for mitochondrial complex I levels (NDUFS4) in cell lines re-expressing catalytically deficient forms of SHMT2.
Extended Data Figure 3 Restoring SHMT2 catalytic activity normalizes 1C flux, respiratory chain expression, glycolytic activity, and cell growth.
a, Immunoblot of re-expression of catalytically active SHMT2 (left) and the effects of its re-expression on mitochondrial complex I and II levels (right). b–f, Effect of re-expression of catalytically active and inactive forms of SHMT2 in two different ΔSHMT2 clones in the HEK293T background. b, Normalized NAD+/NADH ratio (n = 6). c, Lactate secretion and glucose uptake (n = 6). d, Cell proliferation (n = 6). e, Purine biosynthesis intermediate 5-aminoimidazole-4-carboxamide ribonucleotide (AICAR) levels (n = 4) as an indicator of cytosolic folate 1C status. f, [2,3,3-2H]serine tracing to differentiate cytosolic from mitochondrial folate 1C unit production for incorporation into deoxythymidine triphosphate (n = 3). Data are mean ± s.e.m. n indicates the number of biological replicates. *P < 0.01, two-tailed Student’s t-test (see Supplementary Table 7 for exact P values). Source data
Extended Data Figure 4 Oxidative phosphorylation defect is caused by a post-transcriptional mechanism independent of methionine formylation.
a, Fraction of initiating amino acid (formylmethionine versus methionine) of mitochondrial-expressed COX1 peptide determined by high-resolution LC–MS (wild type n = 4, ∆SHMT2 n = 3, ΔMTHFD2 n = 2). b, Lactate secretion (n = 3) upon sarcosine supplementation (1 mM). c, Relative mtDNA levels in HEK293T cells (n = 3). d, Agarose gel of mtDNA long-range PCR products of HCT116 and HEK293T knockout cell lines. e, Relative mRNA levels of mtDNA-encoded respiratory chain subunits in the HEK293T background (n = 3). f, Gene expression levels in SHMT2-knockout cell lines compared to SHMT2 wild-type re-expressed lines by total RNA sequencing. Each dot represents mean gene expression as derived from two biological replicates of two independent knockout clones and matched re-expressed lines (n = 4). Genes linked to human OXPHOS function37 are highlighted in red. Significantly differentially expressed genes are listed in Supplementary Table 2. g, Position-dependent next-generation sequencing coverage of mtDNA in HEK293T wild-type, SHMT2-knockout and MTHFD2-knockout cell lines supports the absence of deletions due to SHMT2 loss. h, Corresponding variant position and frequency. Variant list is provided in Supplementary Table 1. Data are mean ± s.e.m. n indicates the number of biological replicates. *P < 0.01, two-tailed Student’s t-test (see Supplementary Table 7 for exact P values).
a, SDS–PAGE of [35S]methionine-labelled mitochondrially translated proteins in wild-type (lane 1) and two SHMT2-knockout (lane 2 and 3) HEK293T cell lines. Decreased synthesis of COX1 and COX2/3 are evident upon short exposure and reduced synthesis of ND5 and ND6 is more easily visualized upon longer exposure. b, Absorbance at 254 nm upon sucrose gradient fractionation of cell lysates digested by micrococcal nuclease (Fig. 3a). Fractions corresponding to 4 and 5 were collected for mitochondrial ribosome enrichment as shown on the matched immunoblot for mitochondrial ribosome subunit MRPL11. c, Read length distribution (top) and read length-dependent sub-codon read phasing (bottom) across the 13 mitochondrial protein-coding transcripts. Data in c are based on the mitochondrial ribosome profiling experiment in Fig. 3, and represent the mean of two technical replicates of two independent samples.
Extended Data Figure 6 Mitochondrial ribosome stalling at guanosine-ending split codon box nucleotide triplets suggests deficient 5-taurinomethyluridine modification.
a, Expanded version of Fig. 3b, showing the mean cumulative ribosome protected fragments of all mitochondrial protein-coding genes. b, Mean relative density of actively translating (that is, not stalled) ribosomes for mitochondrial transcripts. Data in a and b represent two technical replicates of two independent samples. c, Enzymatic activities of citrate synthase and individual mitochondrial respiratory chain complexes from mitochondrial extracts (n = 5). Data are mean ± s.e.m. *P < 0.01, two-tailed Student’s t-test (see Supplementary Table 7 for exact P values). d, Mitochondrial genetic code table with split codon boxes depending on taurinomethylated tRNAs for translation highlighted in red. Codons decoded by anticodon formylcytidine-containing tRNAMet are highlighted in blue. e, Mean codon-specific mitochondrial ribosome occupancy of HCT116 SHMT2/MTHFD2 double-knockout cell lines supplemented with sarcosine (1 mM). Codons highlighted in red are decoded by tRNAs carrying a 5-taurinomethyluridine modification. The supplementation with sarcosine prevents the stalling normally observed with SHMT2 deletion (n = 2). Source data
Extended Data Figure 7 tRNA modification status in ∆SHMT2 and effects of 5-taurinomethyluridine modification loss caused by human disease gene MTO1.
a, Total ion chromatogram of 5-formylcytidine monophosphate in digested mitochondrial tRNAs upon loss of SHMT2. The same samples were analysed for 5-taurinomethyluridine monophosphate (p-τm5U) in Fig. 4b. The combined data demonstrate that SHMT2 deletion causes loss of τm5U but not 5-formylcytidine. b, Levels of τm5U, 5-taurinomethyl-2-thiouridine monophosphate (p-τm5s2U) and 2-thiouridine monophosphate (p-s2U) in wild-type HCT116 and SHMT2 deletion lines normalized to 5-formylcytidine monophosphate (p-f5C) (n = 3). c, Taurine levels in HCT116 wild-type and SHMT2-knockout cells (n = 3). d, τm5U levels in digested mitochondrial tRNAs upon re-expression of SHMT2 (n = 1). e, τm5U, τm5s2U and s2U levels normalized to f5C in HCT116 SHMT2/MTHFD2 knockout lines after sarcosine supplementation and HCT116 upon loss of MTO1 (n = 2). For all panels, data are mean ± s.e.m. or individual data points only. f, Labelling pattern of 5-taurinomethyluridine and 5-formylcytidine monophosphate extracted from mitochondrial tRNAs after growth in media containing either [3-13C]serine or [U-13C]methionine. g, Mean cumulative count of ribosome protected fragments (RPF) mapping to mitochondrial protein coding transcripts upon ribosome profiling in HCT116 MTO1-knockout cell lines. Data were normalized to RPM (n = 2); n indicates the number of biological replicates. *P < 0.01, two-tailed Student’s t-test (see Supplementary Table 7 for exact P values).
Extended Data Figure 8 Investigation of mRNA and protein secondary structure effects on mitochondrial ribosome stalling sites.
a, Identification of mitochondrial RNA secondary structure based on analysis of the mitochondrial transcript data from the dimethyl sulfate sequencing dataset published previously34. R values and Gini differences were calculated to detect changes in nucleotide reactivity between the in vivo and denatured condition for the complete mitochondrial transcriptome. Coloured points indicate structured regions given in Supplementary Table 4. b, Determination of ribosome stalling sites in SHMT2-knockout HCT116 cell lines. Data points represent individual codons of all 13 mitochondrial protein-coding transcripts. For each codon, the y axis indicates the ribosome counts normalized to the gene median in RPM. The x axis indicates the ratio of normalized counts in SHMT2-knockout to normalized counts in wild-type HCT116. Two and three s.d. above the mean of all codons in the genome are indicated by the grey and black dotted line, respectively. Highlighted in red are codons with greater than 2 s.d. c, Mapping of AAG and UUG codons from SHMT2 knockout-specific ribosome stalling sites (>3 s.d.) on protein structures. For b and c, analysis is based on ribosome profiling data in Fig. 3, with two technical replicates of two independent samples. A list of identified codons and mapped AAG and UUG sites is provided in Supplementary Table 5.
Extended Data Figure 9 Mitochondrial transcript codon occupancy from ribosome profiling of individual patient lines.
a, Codon-specific mitochondrial ribosome occupancy ratio (patient/control fibroblasts) in individual patient derived cell lines (n = 1 for each individual patient, normalized to mean of n = 2 control fibroblast lines). Patients either had nuclear MTO1 missense mutations (patient A c.[1261-5T>G];[1430G>A], patient B c.[1222T>A];[1222T>A]) or were diagnosed with MELAS and carry the recurrent point mutation m.3243A>G in the mitochondrial gene for tRNA Leu1 (MT-TL1). b, Next-generation sequencing of mtDNA mutation load m.3243A>G (MT-TL1) in control fibroblasts and MELAS patient cell lines. Each bar shows one biological replicate for control and patient cell lines. Integrative genomics viewer sequencing raw data are shown on the right.
a, Mitochondrial complex I and II levels after growth in the absence of folate for five passages or in the presence of the indicated methotrexate concentration for 96 h. Ethidium bromide (250 nM) was used as a positive control. b, Cellular mtDNA levels in HCT116 cells after folate depletion (with or without 100 μM hypoxanthine and 16 μM thymidine (HT) as rescue agents) or in the presence of methotrexate for 96 h (n = 3). c, To determine whether the decrease in respiration due to methotrexate arises from methotrexate depleting mitochondrial DNA, impairing mitochondrial translation, or a combination, in HCT116 cells we compared the effects of methotrexate (50 nM) to ethidium bromide (250 nM = 100 ng ml−1), which is classically used to deplete mitochondrial DNA, and to chloramphenicol (310 μM = 100 μg ml−1), which blocks mitochondrial translation. After 48 h of treatment, methotrexate and ethidium bromide both decreased oxygen consumption and DNA content. Importantly, despite ethidium bromide depleting mitochondrial DNA much more strongly, methotrexate had an equivalent effect on oxygen consumption, consistent with the effect of methotrexate on oxygen consumption being in part via mitochondrial translation inhibition. Data are normalized and compared to untreated control (all n = 3; except oxygen consumption methotrexate 96 h n = 6 and control n = 4). Data are mean ± s.e.m. n indicates the number of biological replicates. *P < 0.01, two-tailed Student’s t-test (see Supplementary Table 7 for exact P values).
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Morscher, R., Ducker, G., Li, S. et al. Mitochondrial translation requires folate-dependent tRNA methylation. Nature 554, 128–132 (2018) doi:10.1038/nature25460
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