Characterizing metabolism in cancer is crucial for understanding tumor biology and for developing potential therapies. Although most metabolic investigations analyze averaged metabolite levels from all cell compartments, subcellular metabolomics can provide more detailed insight into the biochemical processes associated with the disease. Methodological limitations have historically prevented the wider application of subcellular metabolomics in cancer research. Recently, however, ways to distinguish and identify metabolic pathways within organelles have been developed, including state-of-the-art methods to monitor metabolism in situ (such as mass spectrometry-based imaging, Raman spectroscopy and fluorescence microscopy), to isolate key organelles via new approaches and to use tailored isotope-tracing strategies. Herein, we examine the advantages and limitations of these developments and look to the future of this field of research.
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Wellen, K. E. & Snyder, N. W. Should we consider subcellular compartmentalization of metabolites, and if so, how do we measure them? Curr. Opin. Clin. Nutr. 22, 347–354 (2019).
Obel, L. F. et al. Brain glycogen-new perspectives on its metabolic function and regulation at the subcellular level. Front. Neuroenergetics 4, 3 (2012).
Pavlova, N. N. & Thompson, C. B. The emerging hallmarks of cancer metabolism. Cell Metab. 23, 27–47 (2016).
Perera, R. M. et al. Transcriptional control of autophagy-lysosome function drives pancreatic cancer metabolism. Nature 524, 361–365 (2015).
Bankaitis, V. A., Garcia-Mata, R. & Mousley, C. J. Golgi membrane dynamics and lipid metabolism. Curr. Biol. 22, R414–R424 (2012).
El Mjiyad, N., Caro-Maldonado, A., Ramirez-Peinado, S. & Munoz-Pinedo, C. Sugar-free approaches to cancer cell killing. Oncogene 30, 253–264 (2011).
Bui, S., Mejia, I., Díaz, B. & Wang, Y. Adaptation of the Golgi apparatus in cancer cell invasion and metastasis. Front. Cell Dev. Biol. https://doi.org/10.3389/fcell.2021.806482 (2021).
Puthalakath, H. et al. ER stress triggers apoptosis by activating BH3-only protein Bim. Cell 129, 1337–1349 (2007).
Garg, A. D., Maes, H., van Vliet, A. R. & Agostinis, P. Targeting the hallmarks of cancer with therapy-induced endoplasmic reticulum (ER) stress. Mol. Cell Oncol. 2, e975089 (2015).
Piao, S. & Amaravadi, R. K. Targeting the lysosome in cancer. Ann. NY Acad. Sci. 1371, 45–54 (2016).
Koivusalo, M. et al. Amiloride inhibits macropinocytosis by lowering submembranous pH and preventing Rac1 and Cdc42 signaling. J. Cell Biol. 188, 547–563 (2010).
de Araujo, M. E. & Huber, L. A. Subcellular fractionation. Methods Mol. Biol. 357, 73–85 (2007).
Michelsen, U. & von Hagen, J. Isolation of subcellular organelles and structures. Methods Enzymol. 463, 305–328 (2009).
Suzuki, K., Bose, P., Leong-Quong, R. Y., Fujita, D. J. & Riabowol, K. REAP: a two minute cell fractionation method. BMC Res. Notes 3, 294 (2010).
Zoncu, R. et al. mTORC1 senses lysosomal amino acids through an inside-out mechanism that requires the vacuolar H+-ATPase. Science 334, 678–683 (2011).
Franko, A. et al. Efficient isolation of pure and functional mitochondria from mouse tissues using automated tissue disruption and enrichment with anti-TOM22 magnetic beads. Plos ONE https://doi.org/10.1371/journal.pone.0082392 (2013).
Xiong, J. et al. Rapid affinity purification of intracellular organelles using a Twin-Strep-tag. J. Cell Sci. https://doi.org/10.1242/jcs.235390 (2019).
Fan, J. et al. Quantitative flux analysis reveals folate-dependent NADPH production. Nature 510, 298 (2014).
Lewis, C. A. et al. Tracing compartmentalized NADPH metabolism in the cytosol and mitochondria of mammalian cells. Mol. Cell 55, 253–263 (2014).
Lita, A. et al. Toward single-organelle lipidomics in live cells. Anal. Chem. 91, 11380–11387 (2019).
Duncan, K. D., Fyrestam, J. & Lanekoff, I. Advances in mass spectrometry based single-cell metabolomics. Analyst 144, 782–793 (2019).
Hogeboom, G. H., Schneider, W. C. & Pallade, G. E. Cytochemical studies of mammalian tissues: i. isolation of intact mitochondria from rat liver; some biochemical properties of mitochondria and submicroscopic particulate materiaL. J. Biol. Chem. 172, 619–635 (1948).
Michelsen, U. & von Hagen, J. in Methods in Enzymology Vol. 463 (eds Burgess, R. R. & Deutscher, M. P.) 305–328 (Academic Press, 2009).
Frezza, C., Cipolat, S. & Scorrano, L. Organelle isolation: functional mitochondria from mouse liver, muscle and cultured filroblasts. Nat. Protoc. 2, 287–295 (2007).
Graham, J. M. Isolation of lysosomes from tissues and cells by differential and density gradient centrifugation. Curr. Protoc. Cell Biol. Chapter 3, Unit 3.6 (2001).
Graham, J. M. Purification of a crude mitochondrial fraction by density-gradient centrifugation. Curr. Protoc. Cell Biol. Chapter 3, Unit 3.4 (2001).
Graham, J. M. Isolation of Golgi membranes from tissues and cells by differential and density gradient centrifugation. Curr. Protoc. Cell Biol. Chapter 3, Unit 3.9 (2001).
Ray, G. J. et al. A PEROXO-tag enables rapid isolation of peroxisomes from human cells. iScience 23, 101109 (2020).
Chen, W. W., Freinkman, E. & Sabatini, D. M. Rapid immunopurification of mitochondria for metabolite profiling and absolute quantification of matrix metabolites. Nat. Protoc. 12, 2215–2231 (2017).
Abu-Remaileh, M. et al. Lysosomal metabolomics reveals V-ATPase- and mTOR-dependent regulation of amino acid efflux from lysosomes. Science 358, 807 (2017).
Commisso, C. et al. Macropinocytosis of protein is an amino acid supply route in Ras-transformed cells. Nature 497, 633 (2013).
Bartel, K. et al. Connecting lysosomes and mitochondria - a novel role for lipid metabolism in cancer cell death. Cell Commun. Signal. https://doi.org/10.1186/s12964-019-0399-2 (2019).
Schieder, M., Rotzer, K., Bruggemann, A., Biel, M. & Wahl-Schott, C. Planar patch clamp approach to characterize ionic currents from intact lysosomes. Sci. Signal 3, pl3 (2010).
Kessler, S. W. Rapid isolation of antigens from cells with a staphylococcal protein A-antibody adsorbent: parameters of the interaction of antibody-antigen complexes with protein A. J. Immunol. 115, 1617–1624 (1975).
Schrader, M. & Fahimi, H. D. Peroxisomes and oxidative stress. Biochim.Biophys. Acta Mol. Cell Res. 1763, 1755–1766 (2006).
Gronemeyer, T. et al. The proteome of human liver peroxisomes: identification of five new peroxisomal constituents by a label-free quantitative proteomics survey. PLoS ONE 8, e57395 (2013).
Waterham, H. R. & Ebberink, M. S. Genetics and molecular basis of human peroxisome biogenesis disorders. Biochim. Biophys. Acta 1822, 1430–1441 (2012).
Fransen, M., Nordgren, M., Wang, B. & Apanasets, O. Role of peroxisomes in ROS/RNS-metabolism: implications for human disease. Biochim. Biophys. Acta 1822, 1363–1373 (2012).
Misra, P. & Reddy, J. K. Peroxisome proliferator-activated receptor-α activation and excess energy burning in hepatocarcinogenesis. Biochimie 98, 63–74 (2014).
Dahabieh, M. S. et al. Peroxisomes protect lymphoma cells from HDAC inhibitor-mediated apoptosis. Cell Death Differ. 24, 1912–1924 (2017).
Meyer, K. et al. Molecular profiling of hepatocellular carcinomas developing spontaneously in acyl-CoA oxidase deficient mice: comparison with liver tumors induced in wild-type mice by a peroxisome proliferator and a genotoxic carcinogen. Carcinogenesis 24, 975–984 (2003).
Box, A., Alshalalfa, M., Hegazy, S. A., Donnelly, B. & Bismar, T. A. High α-methylacyl-CoA racemase (AMACR) is associated with ERG expression and with adverse clinical outcome in patients with localized prostate cancer. Tumor Biol. 37, 12287–12299 (2016).
Benjamin, D. I. et al. Ether lipid generating enzyme AGPS alters the balance of structural and signaling lipids to fuel cancer pathogenicity. PNAS 110, 14912–14917 (2013).
Volkl, A. & Fahimi, H. D. Isolation and Characterization of Peroxisomes from the Liver of Normal Untreated Rats. Eur. J. Biochem. 149, 257–265 (1985).
Singh, I., Carillo, O. & Namboodiri, A. Isolation and biochemical characterization of peroxisomes from cultured rat glial cells. Neurochem. Res. 25, 197–203 (2000).
Chen, X. F. et al. SIRT5 inhibits peroxisomal ACOX1 to prevent oxidative damage and is downregulated in liver cancer. Embo Rep. https://doi.org/10.15252/embr.201745124 (2018).
Antonenkov, V. D., Sormunen, R. T. & Hiltunen, J. K. The rat liver peroxisomal membrane forms a permeability barrier for cofactors but not for small metabolites in vitro. J. Cell Sci. 117, 5633–5642 (2004).
DeBerardinis, R. J. & Chandel, N. S. Fundamentals of cancer metabolism. Sci. Adv. 2, e1600200 (2016).
Gaude, E. & Frezza, C. Defects in mitochondrial metabolism and cancer. Cancer Metab. https://doi.org/10.1186/2049-3002-2-10 (2014).
Roede, J. R., Park, Y., Li, S. Z., Strobel, F. H. & Jones, D. P. Detailed mitochondrial phenotyping by high resolution metabolomics. Plos ONE https://doi.org/10.1371/journal.pone.0033020 (2012).
Sims, N. R. Rapid isolation of metabolically active mitochondria from rat-brain and subregions using Percoll density gradient centrifugation. J. Neurochem. 55, 698–707 (1990).
Fernández-Vizarra, E. et al. Isolation of mitochondria for biogenetical studies: an update. Mitochondrion 10, 253–262 (2010).
Hornig-Do, H. T. et al. Isolation of functional pure mitochondria by superparamagnetic microbeads. Anal. Biochem. 389, 1–5 (2009).
Chen, W. W., Freinkman, E., Wang, T., Birsoy, K. & Sabatini, D. M. Absolute quantification of matrix metabolites reveals the dynamics of mitochondrial metabolism. Cell 166, 1324 (2016).
Bayraktar, E. C. et al. MITO-tag mice enable rapid isolation and multimodal profiling of mitochondria from specific cell types in vivo. PNAS 116, 303–312 (2019).
Lee, A. H. & Glimcher, L. H. Intersection of the unfolded protein response and hepatic lipid metabolism. Cell. Mol. Life Sci. 66, 2835–2850 (2009).
Chakravarthi, S., Jessop, C. E. & Bulleid, N. J. The role of glutathione in disulphide bond formation and endoplasmic-reticulum-generated oxidative stress. Embo Rep. 7, 271–275 (2006).
Zanotto-Filho, A. et al. Alkylating agent-induced NRF2 blocks endoplasmic reticulum stress-mediated apoptosis via control of glutathione pools and protein thiol homeostasis. Mol. Cancer Ther. 15, 3000–3014 (2016).
Staudacher, J. J. et al. Hypoxia-induced gene expression results from selective mRNA partitioning to the endoplasmic reticulum. Nucleic Acids Res. 43, 3219–3236 (2015).
Lita, A. et al. IDH1 mutations induce organelle defects via dysregulated phospholipids. Nat. Commun. 12, 614 (2021).
Cleves, A., McGee, T. & Bankaitis, V. Phospholipid transfer proteins: a biological debut. Trends Cell Biol. 1, 30–34 (1991).
Mizuno-Yamasaki, E., Medkova, M., Coleman, J. & Novick, P. Phosphatidylinositol 4-phosphate controls both membrane recruitment and a regulatory switch of the Rab GEF Sec2p. Dev. Cell 18, 828–840 (2010).
Morrow, A. A. et al. The lipid kinase PI4KIIIβ is highly expressed in breast tumors and activates Akt in cooperation with Rab11a. Mol. Cancer Res 12, 1492–1508 (2014).
Tokuda, E. et al. Phosphatidylinositol 4-phosphate in the Golgi apparatus regulates cell-cell adhesion and invasive cell migration in human breast cancer. Cancer Res. 74, 3054–3066 (2014).
Leelavathi, D. E., Estes, L. W., Feingold, D. S. & Lombardi, B. Isolation of a Golgi-rich fraction from rat liver. Biochim. Biophys. Acta Biomembranes 211, 124–138 (1970).
Fleischer, B. in Methods in Enzymology Vol. 98, 60–67 (Academic Press, 1983).
Wibo, M., Thinès-Sempoux, D., Amar-Costesec, A., Beaufay, H. & Godelaine, D. Analytical study of microsomes and isolated subcellular membranes from rat liver VIII. Subfractionation of preparations enriched with plasma membranes, outer mitochondrial membranes, or Golgi complex membranes. J. Cell Biol. 89, 456–474 (1981).
Cheetham, R. D., Mooré, D. J. & Yunghans, W. N. Isolation of a Golgi apparatus-rich fraction from rat liver. II. Enzymatic characterization and comparison with other cell fractions. J. Cell Biol. 44, 492–500 (1970).
Morré, D. J. & Mollenhauer, H. H. Isolation of the Golgi apparatus from plant cells. J. Cell Biol. 23, 295–305 (1964).
Morré, D. J. et al. Isolation of a Golgi apparatus-rich fraction from rat liver. I. Method and morphology. J. Cell Biol. 44, 484–491 (1970).
Sivanand, S. et al. Nuclear acetyl-CoA production by ACLY promotes homologous recombination. Mol. Cell 67, 252–265 (2017).
Li, X. et al. Nucleus-translocated ACSS2 promotes gene transcription for lysosomal biogenesis and autophagy. Mol. Cell 66, 684–697 (2017).
Boukouris, A. E., Zervopoulos, S. D. & Michelakis, E. D. Metabolic enzymes moonlighting in the nucleus: metabolic regulation of gene transcription. Trends Biochem. Sci. 41, 712–730 (2016).
Sutendra, G. et al. A nuclear pyruvate dehydrogenase complex is important for the generation of Acetyl-CoA and histone acetylation. Cell 158, 84–97 (2014).
Jang, C., Chen, L. & Rabinowitz, J. D. Metabolomics and isotope tracing. Cell 173, 822–837 (2018).
Le, A. et al. Glucose-independent glutamine metabolism via TCA cycling for proliferation and survival in B cells. Cell Metab. 15, 110–121 (2012).
Dong, W., Moon, S. J., Kelleher, J. K. & Stephanopoulos, G. Dissecting mammalian cell metabolism through 13C- and 2H-isotope tracing: interpretations at the molecular and systems levels. Ind. Eng. Chem. Res. 59, 2593–2610 (2020).
Zhang, G. F. et al. Reductive TCA cycle metabolism fuels glutamine- and glucose-stimulated insulin secretion. Cell Metab. 33, 804–817 (2021).
Pongratz, R. L., Kibbey, R. G., Shulman, G. I. & Cline, G. W. Cytosolic and mitochondrial malic enzyme isoforms differentially control insulin secretion. J. Biol. Chem. 282, 200–207 (2007).
Loeber, G., Dworkin, M. B., Infante, A. & Ahorn, H. Characterization of cytosolic malic enzyme in human tumor cells. FEBS Lett. 344, 181–186 (1994).
Warburg, O. On the origin of cancer cells. Science 123, 309–314 (1956).
Ruiz-Rodado, V. et al. Metabolic reprogramming associated with aggressiveness occurs in the G-CIMP-high molecular subtypes of IDH1mut lower grade gliomas. Neuro. Oncol. 22, 480–492 (2019).
Yang, C. et al. Glutamine oxidation maintains the TCA cycle and cell survival during impaired mitochondrial pyruvate transport. Mol. Cell 56, 414–424 (2014).
Murai, S. et al. Inhibition of malic enzyme 1 disrupts cellular metabolism and leads to vulnerability in cancer cells in glucose-restricted conditions. Oncogenesis 6, e329 (2017).
Zhang, Z., Chen, L., Liu, L., Su, X. & Rabinowitz, J. D. Chemical basis for deuterium labeling of fat and NADPH. JACS 139, 14368–14371 (2017).
Badur, M. G. et al. Oncogenic R132 IDH1 mutations limit NADPH for de novo lipogenesis through (D)2-hydroxyglutarate production in fibrosarcoma cells. Cell Rep. 25, 1680 (2018).
Lim, E. W., Parker, S. J. & Metallo, C. M. Deuterium tracing to interrogate compartment-specific NAD(P)H metabolism in cultured mammalian cells. Methods Mol. Biol. 2088, 51–71 (2020).
Tedeschi, P. M. et al. Contribution of serine, folate and glycine metabolism to the ATP, NADPH and purine requirements of cancer cells. Cell Death Dis. 4, e877 (2013).
Bradley, K. K. & Bradley, M. E. Purine nucleoside-dependent inhibition of cellular proliferation in 1321N1 human astrocytoma cells. J. Pharmacol. Exp. Ther. 299, 748–752 (2001).
Nonnenmacher, Y. et al. Analysis of mitochondrial metabolism in situ: Combining stable isotope labeling with selective permeabilization. Metab. Eng. 43, 147–155 (2017).
Davidson, S. M. et al. Direct evidence for cancer-cell-autonomous extracellular protein catabolism in pancreatic tumors. Nat. Med. 23, 235–241 (2017).
Trefely, S. et al. Subcellular metabolic pathway kinetics are revealed by correcting for artifactual post harvest metabolism. Mol. Metab. 30, 61–71 (2019).
Zenobi, R. Single-cell metabolomics: analytical and biological perspectives. Science 342, 1243259 (2013).
Caprioli, R. M., Farmer, T. B. & Gile, J. Molecular imaging of biological samples: localization of peptides and proteins using MALDI-TOF MS. Anal. Chem. 69, 4751–4760 (1997).
Takats, Z., Wiseman, J. M., Gologan, B. & Cooks, R. G. Mass spectrometry sampling under ambient conditions with desorption electrospray ionization. Science 306, 471–473 (2004).
Nemes, P. & Vertes, A. Laser ablation electrospray ionization for atmospheric pressure, in vivo, and imaging mass spectrometry. Anal. Chem. 79, 8098–8106 (2007).
Gillen, G., Simons, D. S. & Williams, P. Molecular ion imaging and dynamic secondary ion mass spectrometry of organic compounds. Anal. Chem. 62, 2122–2130 (1990).
Gilmore, I. S., Heiles, S. & Pieterse, C. L. Metabolic imaging at the single-cell scale: recent advances in mass spectrometry imaging. Annu Rev. Anal. Chem. 12, 201–224 (2019).
Hu, R., Li, Y., Yang, Y. & Liu, M. Mass spectrometry-based strategies for single-cell metabolomics. Mass Spectrom. Rev. https://doi.org/10.1002/mas.21704 (2021).
Alexandrov, T. Spatial metabolomics and imaging mass spectrometry in the age of artificial intelligence. Annu Rev. Biomed. Data Sci. 3, 61–87 (2020).
Porta Siegel, T. et al. Mass spectrometry imaging and integration with other imaging modalities for greater molecular understanding of biological tissues. Mol. Imaging Biol. 20, 888–901 (2018).
Amantonico, A., Urban, P. L., Fagerer, S. R., Balabin, R. M. & Zenobi, R. Single-cell MALDI-MS as an analytical tool for studying intrapopulation metabolic heterogeneity of unicellular organisms. Anal. Chem. 82, 7394–7400 (2010).
Li, L., Garden, R. W. & Sweedler, J. V. Single-cell MALDI: a new tool for direct peptide profiling. Trends Biotechnol. 18, 151–160 (2000).
Xiong, C. et al. Development of visible-wavelength MALDI cell mass spectrometry for high-efficiency single-cell analysis. Anal. Chem. 88, 11913–11918 (2016).
Ong, T. H. et al. Classification of large cellular populations and discovery of rare cells using single cell matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Anal. Chem. 87, 7036–7042 (2015).
Castro, D. C., Xie, Y. R., Rubakhin, S. S., Romanova, E. V. & Sweedler, J. V. Image-guided MALDI mass spectrometry for high-throughput single-organelle characterization. Nat. Methods 18, 1233–1238 (2021).
Chelgani, S. C. & Hart, B. TOF-SIMS studies of surface chemistry of minerals subjected to flotation separation: a review. Miner. Eng. 57, 1–11 (2014).
Denbigh, J. L. & Lockyer, N. P. ToF-SIMS as a tool for profiling lipids in cancer and other diseases. Mater. Sci. Tech. 31, 137–147 (2015).
Fearn, S. Characterisation of biological material with ToF-SIMS: a review. Mater. Sci. Tech. 31, 148–161 (2015).
Gilmore, I. S. SIMS of organics-Advances in 2D and 3D imaging and future outlook. J Vac. Sci. Technol. A https://doi.org/10.1116/1.4816935 (2013).
Fletcher, J. S., Lockyer, N. P., Vaidyanathan, S. & Vickerman, J. C. TOF-SIMS 3D biomolecular imaging of Xenopus laevis oocytes using buckminsterfullerene (C60) primary ions. Anal. Chem. 79, 2199–2206 (2007).
Piehowski, P. D. et al. MS/MS methodology to improve subcellular mapping of cholesterol Using TOF-SIMS. Anal. Chem. 80, 8662–8667 (2008).
Passarelli, M. K. et al. The 3D OrbiSIMS-label-free metabolic imaging with subcellular lateral resolution and high mass-resolving power. Nat. Methods 14, 1175–1183 (2017).
Mizuno, H., Tsuyama, N., Date, S., Harada, T. & Masujima, T. Live single-cell metabolomics of tryptophan and histidine metabolites in a rat basophil leukemia cell. Anal. Sci. 24, 1525–1527 (2008).
Mizuno, H., Tsuyama, N., Harada, T. & Masujima, T. Live single-cell video-mass spectrometry for cellular and subcellular molecular detection and cell classification. J. Mass Spectrom. 43, 1692–1700 (2008).
Ali, A. et al. Single-cell metabolomics by mass spectrometry: advances, challenges, and future applications. Trends Anal. Chem. https://doi.org/10.1016/j.trac.2019.02.033 (2019).
Zhu, H. et al. Metabolomic profiling of single enlarged lysosomes. Nat. Methods 18, 788–798 (2021).
Cheng, J. X. & Xie, X. S. Vibrational spectroscopic imaging of living systems: an emerging platform for biology and medicine. Science https://doi.org/10.1126/science.aaa8870 (2015).
Baker, M. J. et al. Using Fourier transform IR spectroscopy to analyze biological materials. Nat. Protoc. 9, 1771–1791 (2014).
Li, J. J. & Cheng, J. X. Direct visualization of de novo lipogenesis in single living cells. Sci. Rep. https://doi.org/10.1038/srep06807 (2014).
Quaroni, L. Characterization of intact eukaryotic cells with subcellular spatial resolution by photothermal-induced resonance infrared spectroscopy and imaging. Molecules https://doi.org/10.3390/molecules24244504 (2019).
Shipp, D. W., Sinjab, F. & Notingher, I. Raman spectroscopy: techniques and applications in the life sciences. Adv. Opt. Photonics 9, 315–428 (2017).
Kuzmin, A. N. et al. Resonance Raman probes for organelle-specific labeling in live cells. Sci. Rep. https://doi.org/10.1038/srep28483 (2016).
Xie, C. G., Goodman, C., Dinno, M. A. & Li, Y. Q. Real-time Raman spectroscopy of optically trapped living cells and organelles. Opt. Express. 12, 6208–6214 (2004).
Rahmelow, K. & Hubner, W. Infrared spectroscopy in aqueous solution: difficulties and accuracy of water subtraction. Appl. Spectrosc. 51, 160–170 (1997).
Kuzmin, A. N., Levchenko, S. M., Pliss, A., Qu, J. L. & Prasad, P. N. Molecular profiling of single organelles for quantitative analysis of cellular heterogeneity. Sci. Rep. https://doi.org/10.1038/s41598-017-06936-z (2017).
Kuzmin, A. N., Pliss, A., Rzhevskii, A., Lita, A. & Larion, M. BCAbox algorithm expands capabilities of Raman microscope for single organelles assessment. Biosensors https://doi.org/10.3390/bios8040106 (2018).
Levchenko, S. M., Kuzmin, A. N., Pliss, A., Qu, J. L. & Prasad, P. N. Macromolecular profiling of organelles in normal diploid and cancer cells. Anal. Chem. 89, 10985–10990 (2017).
Pliss, A., Kuzmin, A. N., Kachynski, A. V. & Prasad, P. N. Nonlinear optical imaging and raman microspectrometry of the cell nucleus throughout the cell cycle. Biophys. J. 99, 3483–3491 (2010).
Mourant, J. R. et al. Biochemical differences in tumorigenic and nontumorigenic cells measured by Raman and infrared spectroscopy. J. Biomed. Opt. https://doi.org/10.1117/1.1928050 (2005)
Short, K. W., Carpenter, S., Freyer, J. P. & Mourant, J. R. Raman spectroscopy detects biochemical changes due to proliferation in mammalian cell cultures. Biophys. J. 88, 4274–4288 (2005).
Hu, F., Shi, L. & Min, W. Biological imaging of chemical bonds by stimulated Raman scattering microscopy. Nat. Methods 16, 830–842 (2019).
Zhang, L. et al. Spectral tracing of deuterium for imaging glucose metabolism. Nat. Biomed. Eng. 3, 402–413 (2019).
Pezacki, J. P. et al. Chemical contrast for imaging living systems: molecular vibrations drive CARS microscopy. Nat. Chem. Biol. 7, 137–145 (2011).
Yu, Y., Ramachandran, P. V. & Wang, M. C. Shedding new light on lipid functions with CARS and SRS microscopy. Mol. Cell Biol. 1841, 1120–1129 (2014).
Le, T. T., Huff, T. B. & Cheng, J. X. Coherent anti-Stokes Raman scattering imaging of lipids in cancer metastasis. BMC Cancer https://doi.org/10.1186/1471-2407-9-42 (2009).
Wei, L. et al. Super-multiplex vibrational imaging. Nature 544, 465–470 (2017).
Goldbeck, O., Eck, A. W. & Seibold, G. M. Real-time monitoring of NADPH concentrations in Corynebacterium glutamicum and Escherichia coli via the genetically encoded sensor mBFP. Front. Microbiol. https://doi.org/10.3389/fmicb.2018.02564 (2018).
Shen, Y. et al. Organelle-targeting gold nanorods for macromolecular profiling of subcellular organelles and enhanced cancer cell killing. ACS Appl. Mater. Interfaces 10, 7910–7918 (2018).
Shen, Y. et al. Organelle-targeting surface-enhanced Raman scattering (SERS) nanosensors for subcellular pH sensing. Nanoscale 10, 1622–1630 (2018).
Sun, C. L., Gao, M. X. & Zhang, X. M. Surface-enhanced Raman scattering (SERS) imaging-guided real-time photothermal ablation of target cancer cells using polydopamine-encapsulated gold nanorods as multifunctional agents. Anal. Bioanal. Chem. 409, 4915–4926 (2017).
Sheppard, C. J. R. Multiphoton microscopy: a personal historical review, with some future predictions. J. Biomed. Opt. https://doi.org/10.1117/1.Jbo.25.1.014511 (2020).
Denk, W., Strickler, J. H. & Webb, W. W. Two-photon laser scanning fluorescence microscopy. Science 248, 73–76 (1990).
Roshanzadeh, A. et al. Real-time monitoring of NADPH levels in living mammalian cells using fluorescence-enhancing protein bound to NADPHs. Biosens. Bioelectron. https://doi.org/10.1016/j.bios.2019.111753 (2019).
Xu, A., Tang, Y. & Lin, W. Endoplasmic reticulum-targeted two-photon turn-on fluorescent probe for nitroreductase in tumor cells and tissues. Spectrochim. Acta A Mol. Biomol. Spectrosc. 204, 770–776 (2018).
Hong, S., Pawel, G. T., Pei, R. & Lu, Y. Recent progress in developing fluorescent probes for imaging cell metabolites. Biomed. Mater. 16, 044108 (2021).
Choi, N. E., Lee, J. Y., Park, E. C., Lee, J. H. & Lee, J. Recent Advances in Organelle-Targeted Fluorescent Probes. Molecules https://doi.org/10.3390/molecules26010217 (2021).
Gao, P., Pan, W., Li, N. & Tang, B. Fluorescent probes for organelle-targeted bioactive species imaging. Chem. Sci. 10, 6035–6071 (2019).
Xu, Z. & Xu, L. Fluorescent probes for the selective detection of chemical species inside mitochondria. Chem. Commun. 52, 1094–1119 (2016).
Blacker, T. S. et al. Separating NADH and NADPH fluorescence in live cells and tissues using FLIM. Nat. Commun. https://doi.org/10.1038/ncomms4936 (2014).
Skala, M. C. et al. In vivo multiphoton fluorescence lifetime imaging of protein-bound and free nicotinamide adenine dinucleotide in normal and precancerous epithelia. J. Biomed. Opt. https://doi.org/10.1117/1.2717503 (2007).
Tao, R. K. et al. Genetically encoded fluorescent sensors reveal dynamic regulation of NADPH metabolism. Nat. Methods 14, 720 (2017).
Penjweini, R. et al. Single cell-based fluorescence lifetime imaging of intracellular oxygenation and metabolism. Redox Biol. 34, 101549 (2020).
Dickinson, B. C. & Chang, C. J. A targetable fluorescent probe for imaging hydrogen peroxide in the mitochondria of living cells. JACS 130, 9638–9639 (2008).
Tan, K.-Y. et al. Real-time monitoring ATP in mitochondrion of living cells: a specific fluorescent probe for atp by dual recognition sites. Anal. Chem. 89, 1749–1756 (2017).
Johnson-Cadwell, L. I., Jekabsons, M. B., Wang, A., Polster, B. M. & Nicholls, D. G. ‘Mild uncoupling’ does not decrease mitochondrial superoxide levels in cultured cerebellar granule neurons but decreases spare respiratory capacity and increases toxicity to glutamate and oxidative stress. J. Neurochem. 101, 1619–1631 (2007).
Zielonka, J. et al. Global profiling of reactive oxygen and nitrogen species in biological systems high-throughput real-time analyses. J. Biol. Chem. 287, 2984–2995 (2012).
Polster, B. M., Nicholls, D. G., Ge, S. X. & Roelofs, B. A. Use of potentiometric fluorophores in the measurement of mitochondrial reactive oxygen species. Method. Enzymol. 547, 225–250 (2014).
Michalski, R., Michalowski, B., Sikora, A., Zielonka, J. & Kalyanaraman, B. On the use of fluorescence lifetime imaging and dihydroethidium to detect superoxide in intact animals and ex vivo tissues: a reassessment. Free Radic. Bio. Med. 67, 278–284 (2014).
Patterson, G. H., Knobel, S. M., Arkhammar, P., Thastrup, O. & Piston, D. W. Separation of the glucose-stimulated cytoplasmic and mitochondrial NAD(P)H responses in pancreatic islet β cells. Proc. Natl Acad. Sci. USA 97, 5203 (2000).
Vishnu, N. et al. ATP increases within the lumen of the endoplasmic reticulum upon intracellular Ca2+ release. Mol. Biol. Cell 25, 368–379 (2014).
Zhang, W. et al. Two-photon fluorescence imaging reveals a Golgi apparatus superoxide anion-mediated hepatic ischaemia-reperfusion signalling pathway. Chem. Sci. 10, 879–883 (2018).
Zhang, X. et al. A highly specific Golgi-targetable fluorescent probe for tracking cysteine generation during the Golgi stress response. Sens. Actuators B 310, 127820 (2020).
Circu, M. L. & Aw, T. Y. Reactive oxygen species, cellular redox systems, and apoptosis. Free Radic. Biol. Med. 48, 749–762 (2010).
Jun, Y. W. et al. A ratiometric two-photon fluorescent probe for tracking lysosomal ATP: direct in cellulo observation of lysosomal membrane fusion processes. Angew. Chem. Int. Ed. 57, 10142–10147 (2018).
Wen, Y. et al. A highly sensitive ratiometric fluorescent probe for the detection of cytoplasmic and nuclear hydrogen peroxide. Anal. Chem. 86, 9970–9976 (2014).
Kompauer, M., Heiles, S. & Spengler, B. Autofocusing MALDI mass spectrometry imaging of tissue sections and 3D chemical topography of nonflat surfaces. Nat. Methods 14, 1156–1158 (2017).
Qi, M., Philip, M. C., Yang, N. & Sweedler, J. V. Single cell neurometabolomics. ACS Chem. Neurosci. 9, 40–50 (2018).
Ali, A. et al. Single-cell screening of tamoxifen abundance and effect using mass spectrometry and Raman-spectroscopy. Anal. Chem. 91, 2710–2718 (2019).
We thank E. He, from Medical Arts of the National Institutes of Health for help with figures. This research was supported by the National Institutes of Health Intramural Research Program through an NCI FLEX award to A.L. and M.L. entitled ‘Live cell metabolism via Raman imaging microscopy.’
The authors declare no competing interests.
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Nature Methods thanks Monther Abu-Remaileh, Wei Xiong and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Primary Handling editor: Rita Strack, in collaboration with the Nature Methods team.
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Ruiz-Rodado, V., Lita, A. & Larion, M. Advances in measuring cancer cell metabolism with subcellular resolution. Nat Methods 19, 1048–1063 (2022). https://doi.org/10.1038/s41592-022-01572-6