AMPK as a mediator of tissue preservation: time for a shift in dogma?

Hui, S. et al. Quantitative fluxomics of circulating metabolites. Cell Metab. 32, 676–688 (2020).

pubmed: 32791100 pmcid: 7544659 doi: 10.1016/j.cmet.2020.07.013

Rolfe, D. F. & Brown, G. C. Cellular energy utilization and molecular origin of standard metabolic rate in mammals. Physiol. Rev. 77, 731–758 (1997).

pubmed: 9234964 doi: 10.1152/physrev.1997.77.3.731

Sartori, R., Romanello, V. & Sandri, M. Mechanisms of muscle atrophy and hypertrophy: implications in health and disease. Nat. Commun. 12, 330 (2021).

pubmed: 33436614 pmcid: 7803748 doi: 10.1038/s41467-020-20123-1

Mitch, W. E. & Goldberg, A. L. Mechanisms of muscle wasting – the role of the ubiquitin–proteasome pathway. N. Engl. J. Med. 335, 1897–1905 (1996).

pubmed: 8948566 doi: 10.1056/NEJM199612193352507

Herzig, S. & Shaw, R. J. AMPK: guardian of metabolism and mitochondrial homeostasis. Nat. Rev. Mol. Cell Biol. 19, 121–135 (2018).

pubmed: 28974774 doi: 10.1038/nrm.2017.95

Hardie, D. G., Ross, F. A. & Hawley, S. A. AMPK: a nutrient and energy sensor that maintains energy homeostasis. Nat. Rev. Mol. Cell Biol. 13, 251–262 (2012).

pubmed: 22436748 pmcid: 5726489 doi: 10.1038/nrm3311

Steinberg, G. R. & Hardie, D. G. New insights into activation and function of the AMPK. Nat. Rev. Mol. Cell Biol. 24, 255–272 (2023).

pubmed: 36316383 doi: 10.1038/s41580-022-00547-x

Hawley, S. A. et al. Complexes between the LKB1 tumor suppressor, STRADα/β and MO25α/β are upstream kinases in the AMP-activated protein kinase cascade. J. Biol. 2, 28 (2003).

pubmed: 14511394 pmcid: 333410 doi: 10.1186/1475-4924-2-28

Woods, A. et al. Ca

pubmed: 16054096 doi: 10.1016/j.cmet.2005.06.005

Steinberg, G. R. et al. Tumor necrosis factor ɑ-induced skeletal muscle insulin resistance involves suppression of AMP-kinase signaling. Cell Metab. 4, 465–474 (2006).

pubmed: 17141630 doi: 10.1016/j.cmet.2006.11.005

Joseph, B. K. et al. Inhibition of AMP kinase by the protein phosphatase 2A heterotrimer, PP2A

pubmed: 25694423 pmcid: 4409226 doi: 10.1074/jbc.M114.626259

Sanders, M. J., Grondin, P. O., Hegarty, B. D., Snowden, M. A. & Carling, D. Investigating the mechanism for AMP activation of the AMP-activated protein kinase cascade. Biochem. J. 403, 139–148 (2007).

pubmed: 17147517 pmcid: 1828883 doi: 10.1042/BJ20061520

Zhang, Y. L. et al. AMP as a low-energy charge signal autonomously initiates assembly of AXIN-AMPK-LKB1 complex for AMPK activation. Cell Metab. 18, 546–555 (2013).

pubmed: 24093678 doi: 10.1016/j.cmet.2013.09.005

Fearon, K. C. H., Glass, D. J. & Guttridge, D. C. Cancer cachexia: mediators, signaling, and metabolic pathways. Cell Metab. 16, 153–166 (2012).

pubmed: 22795476 doi: 10.1016/j.cmet.2012.06.011

White, J. P. et al. Muscle mTORC1 suppression by IL-6 during cancer cachexia: a role for AMPK. Am. J. Physiol. Endocrinol. Metab. 304, E1042–E1052 (2013).

pubmed: 23531613 pmcid: 3651620 doi: 10.1152/ajpendo.00410.2012

Aguilar-Recarte, D. et al. GDF15 mediates the metabolic effects of PPARβ/δ by activating AMPK. Cell Rep. 36, 109501 (2021).

pubmed: 34380027 doi: 10.1016/j.celrep.2021.109501

Grossberg, A. J., Scarlett, J. M. & Marks, D. L. Hypothalamic mechanisms in cachexia. Physiol. Behav. 100, 478–489 (2010).

pubmed: 20346963 pmcid: 2927357 doi: 10.1016/j.physbeh.2010.03.011

Bennani-Baiti, N. & Walsh, D. Animal models of the cancer anorexia-cachexia syndrome. Support. Care Cancer 19, 1451–1463 (2011).

pubmed: 20714754 doi: 10.1007/s00520-010-0972-0

Winter, A., MacAdams, J. & Chevalier, S. Normal protein anabolic response to hyperaminoacidemia in insulin-resistant patients with lung cancer cachexia. Clin. Nutr. 31, 765–773 (2012).

pubmed: 22647419 doi: 10.1016/j.clnu.2012.05.003

Yoshikawa, T., Noguchi, Y., Doi, C., Makino, T. & Nomura, K. Insulin resistance in patients with cancer: relationships with tumor site, tumor stage, body-weight loss, acute-phase response, and energy expenditure. Nutrition 17, 590–593 (2001).

pubmed: 11448578 doi: 10.1016/S0899-9007(01)00561-5

Yoshikawa, T. et al. Insulin resistance was connected with the alterations of substrate utilization in patients with cancer. Cancer Lett. 141, 93–98 (1999).

pubmed: 10454248 doi: 10.1016/S0304-3835(99)00086-5

Heber, D., Byerly, L. O. & Chlebowski, R. T. Metabolic abnormalities in the cancer patient. Cancer 55, 225–229 (1985).

pubmed: 3880655 doi: 10.1002/1097-0142(19850101)55:1+<225::AID-CNCR2820551304>3.0.CO;2-7

Màrmol, J. M. et al. Insulin resistance in patients with cancer: a systematic review and meta-analysis. Acta Oncol. 62, 364–371 (2023).

pubmed: 37042166 doi: 10.1080/0284186X.2023.2197124

Smith, K. L. & Tisdale, M. J. Increased protein degradation and decreased protein synthesis in skeletal muscle during cancer cachexia. Br. J. Cancer 67, 680–685 (1993).

pubmed: 8471425 pmcid: 1968351 doi: 10.1038/bjc.1993.126

Emery, P. W., Edwards, R. H., Rennie, M. J., Souhami, R. L. & Halliday, D. Protein synthesis in muscle measured in vivo in cachectic patients with cancer. Br. Med. J. 289, 584–586 (1984).

doi: 10.1136/bmj.289.6445.584

Jeevanandam, M., Lowry, S., Horowitz, G. & Brennan, M. Cancer cachexia and protein metabolism. Lancet 323, 1423–1426 (1984).

doi: 10.1016/S0140-6736(84)91929-9

Lundholm, K., Edström, S., Karlberg, I., Ekman, L. & Scherstén, T. Glucose turnover, gluconeogenesis from glycerol, and estimation of net glucose cycling in cancer patients. Cancer 50, 1142–1150 (1982).

pubmed: 7104955 doi: 10.1002/1097-0142(19820915)50:6<1142::AID-CNCR2820500618>3.0.CO;2-I

Han, X. et al. Cancer causes metabolic perturbations associated with reduced insulin-stimulated glucose uptake in peripheral tissues and impaired muscle microvascular perfusion. Metabolism 105, 154169 (2020).

pubmed: 31987858 doi: 10.1016/j.metabol.2020.154169

Goncalves, M. D. et al. Fenofibrate prevents skeletal muscle loss in mice with lung cancer. Pro. Natl Acad. Sci. USA 115, E743–E752 (2018).

doi: 10.1073/pnas.1714703115

Rohm, M. et al. An AMP-activated protein kinase-stabilizing peptide ameliorates adipose tissue wasting in cancer cachexia in mice. Nat. Med. 22, 1120–1130 (2016).

pubmed: 27571348 doi: 10.1038/nm.4171

Beck, S. A. & Tisdale, M. J. Effect of cancer cachexia on triacylglycerol/fatty acid substrate cycling in white adipose tissue. Lipids 39, 1187–1189 (2004).

pubmed: 15736914 doi: 10.1007/s11745-004-1346-8

Dilman, V. M., Berstein, L. M., Ostroumova, M. N., Tsyrlina, Y. V. & Golubev, A. G. Peculiarities of hyperlipidaemia in tumour patients. Br. J. Cancer 43, 637–643 (1981).

pubmed: 7248149 pmcid: 2010684 doi: 10.1038/bjc.1981.94

Kazemi-Bajestani, S. M., Mazurak, V. C. & Baracos, V. Computed tomography-defined muscle and fat wasting are associated with cancer clinical outcomes. Semin. Cell Dev. Biol. 54, 2–10 (2016).

pubmed: 26343952 doi: 10.1016/j.semcdb.2015.09.001

Stene, G. B. et al. Changes in skeletal muscle mass during palliative chemotherapy in patients with advanced lung cancer. Acta Oncol. 54, 340–348 (2015).

pubmed: 25225010 doi: 10.3109/0284186X.2014.953259

Antoun, S., Borget, I. & Lanoy, E. Impact of sarcopenia on the prognosis and treatment toxicities in patients diagnosed with cancer. Curr. Opin. Support. Palliat. Care 7, 383–389 (2013).

pubmed: 24189893 doi: 10.1097/SPC.0000000000000011

Couch, M. et al. Cancer cachexia syndrome in head and neck cancer patients: part I. Diagnosis, impact on quality of life and survival, and treatment. Head. Neck 29, 401–411 (2007).

pubmed: 17285641 doi: 10.1002/hed.20447

Argilés, J. M., Busquets, S., Stemmler, B. & López-Soriano, F. J. Cancer cachexia: understanding the molecular basis. Nat. Rev. Cancer 14, 754–762 (2014).

pubmed: 25291291 doi: 10.1038/nrc3829

Anker, M. S. et al. Orphan disease status of cancer cachexia in the USA and in the European Union: a systematic review. J. Cachexia Sarcopenia Muscle 10, 22–34 (2019).

pubmed: 30920776 pmcid: 6438416 doi: 10.1002/jcsm.12402

Roeland, E. J. et al. Management of cancer cachexia: ASCO guideline. J. Clin. Oncol. 38, 2438–2453 (2020).

pubmed: 32432946 doi: 10.1200/JCO.20.00611

Segatto, M. et al. Epigenetic targeting of bromodomain protein BRD4 counteracts cancer cachexia and prolongs survival. Nat. Commun. 8, 1707 (2017).

pubmed: 29167426 pmcid: 5700099 doi: 10.1038/s41467-017-01645-7

Raun, S. H., Knudsen, J. R., Han, X., Jensen, T. E. & Sylow, L. Cancer causes dysfunctional insulin signaling and glucose transport in a muscle-type-specific manner. FASEB J. 36, e22211 (2022).

pubmed: 35195922 doi: 10.1096/fj.202101759R

Bohnert, K. R. et al. Inhibition of ER stress and unfolding protein response pathways causes skeletal muscle wasting during cancer cachexia. FASEB J. 30, 3053–3068 (2016).

pubmed: 27206451 pmcid: 5001510 doi: 10.1096/fj.201600250RR

Raun, S. H. et al. Adenosine monophosphate-activated protein kinase is elevated in human cachectic muscle and prevents cancer-induced metabolic dysfunction in mice. J. Cachexia Sarcopenia Muscle 14, 1631–1647 (2023).

pubmed: 37194385 pmcid: 10401533 doi: 10.1002/jcsm.13238

Argilés, J. M., Fontes-Oliveira, C. C., Toledo, M., López-Soriano, F. J. & Busquets, S. Cachexia: a problem of energetic inefficiency. J. Cachexia Sarcopenia Muscle 5, 279–286 (2014).

pubmed: 25118829 pmcid: 4248416 doi: 10.1007/s13539-014-0154-x

Sanchís, D. et al. Skeletal muscle UCP2 and UCP3 gene expression in a rat cancer cachexia model. FEBS Lett. 436, 415–418 (1998).

pubmed: 9801160 doi: 10.1016/S0014-5793(98)01178-8

Kitaoka, Y., Miyazaki, M. & Kikuchi, S. Voluntary exercise prevents abnormal muscle mitochondrial morphology in cancer cachexia mice. Physiol. Rep. 9, e15016 (2021).

pubmed: 34427401 pmcid: 8383714 doi: 10.14814/phy2.15016

Shum, A. M. Y. et al. Proteomic profiling of skeletal and cardiac muscle in cancer cachexia: alterations in sarcomeric and mitochondrial protein expression. Oncotarget 9, 22001 (2018).

pubmed: 29774118 pmcid: 5955146 doi: 10.18632/oncotarget.25146

van der Ende, M. et al. Mitochondrial dynamics in cancer-induced cachexia. Biochim. Biophys. Acta Rev. Cancer 1870, 137–150 (2018).

pubmed: 30059724 doi: 10.1016/j.bbcan.2018.07.008

VanderVeen, B. N., Fix, D. K. & Carson, J. A. Disrupted skeletal muscle mitochondrial dynamics, mitophagy, and biogenesis during cancer cachexia: a role for inflammation. Oxid. Med. Cell. Longev. 2017, 3292087 (2017).

pubmed: 28785374 pmcid: 5530417 doi: 10.1155/2017/3292087

White, J. P. et al. IL-6 regulation on skeletal muscle mitochondrial remodeling during cancer cachexia in the Apc

pubmed: 22769563 pmcid: 3431229 doi: 10.1186/2044-5040-2-14

Brown, J. L. et al. Mitochondrial degeneration precedes the development of muscle atrophy in progression of cancer cachexia in tumour-bearing mice. J. Cachexia Sarcopenia Muscle 8, 926–938 (2017).

pubmed: 28845591 pmcid: 5700433 doi: 10.1002/jcsm.12232

Beltrà, M., Pin, F., Ballarò, R., Costelli, P. & Penna, F. Mitochondrial dysfunction in cancer cachexia: impact on muscle health and regeneration. Cells 10, 3150 (2021).

pubmed: 34831373 pmcid: 8621344 doi: 10.3390/cells10113150

de Castro, G. S. et al. Human cachexia induces changes in mitochondria, autophagy and apoptosis in the skeletal muscle. Cancers 11, 1264 (2019).

pubmed: 31466311 pmcid: 6770124 doi: 10.3390/cancers11091264

Op den Kamp, C. M. et al. Preserved muscle oxidative metabolic phenotype in newly diagnosed non-small cell lung cancer cachexia. J. Cachexia Sarcopenia Muscle 6, 164–173 (2015).

pubmed: 26136192 pmcid: 4458082 doi: 10.1002/jcsm.12007

Pin, F., Barreto, R., Couch, M. E., Bonetto, A. & O’Connell, T. M. Cachexia induced by cancer and chemotherapy yield distinct perturbations to energy metabolism. J. Cachexia Sarcopenia Muscle 10, 140–154 (2019).

pubmed: 30680954 pmcid: 6438345 doi: 10.1002/jcsm.12360

Atkinson, D. E. The energy charge of the adenylate pool as a regulatory parameter. Interaction with feedback modifiers. Biochemistry 7, 4030–4034 (1968).

pubmed: 4972613 doi: 10.1021/bi00851a033

Egan, B. & Zierath, J. R. Exercise metabolism and the molecular regulation of skeletal muscle adaptation. Cell Metab. 17, 162–184 (2013).

pubmed: 23395166 doi: 10.1016/j.cmet.2012.12.012

Fix, D. K. et al. Wheel running improves fasting-induced AMPK signaling in skeletal muscle from tumor-bearing mice. Physiol. Rep. 9, e14924 (2021).

pubmed: 34270178 pmcid: 8284248 doi: 10.14814/phy2.14924

Ballarò, R. et al. Moderate exercise improves experimental cancer cachexia by modulating the redox homeostasis. Cancers 11, 285 (2019).

pubmed: 30823492 pmcid: 6468783 doi: 10.3390/cancers11030285

Ballarò, R. et al. Moderate exercise in mice improves cancer plus chemotherapy-induced muscle wasting and mitochondrial alterations. FASEB J. 33, 5482–5494 (2019).

pubmed: 30653354 doi: 10.1096/fj.201801862R

Wyart, E. et al. Iron supplementation is sufficient to rescue skeletal muscle mass and function in cancer cachexia. EMBO Rep. 23, e53746 (2022).

pubmed: 35199910 pmcid: 8982578 doi: 10.15252/embr.202153746

Velázquez, K. T. et al. Quercetin supplementation attenuates the progression of cancer cachexia in Apc

pubmed: 24759931 pmcid: 4018949 doi: 10.3945/jn.113.188367

Beltrà, M. et al. NAD

pubmed: 37012289 pmcid: 10070388 doi: 10.1038/s41467-023-37595-6

Bujak, A. L. et al. AMPK activation of muscle autophagy prevents fasting-induced hypoglycemia and myopathy during aging. Cell Metab. 21, 883–890 (2015).

pubmed: 26039451 pmcid: 5233441 doi: 10.1016/j.cmet.2015.05.016

Mikhail, A. I., Ng, S. Y., Mattina, S. R. & Ljubicic, V. AMPK is mitochondrial medicine for neuromuscular disorders. Trends Mol. Med. 29, 512–529 (2023).

pubmed: 37080889 doi: 10.1016/j.molmed.2023.03.008

Ozaki, Y. et al. Myonectin protects against skeletal muscle dysfunction in male mice through activation of AMPK/PGC1α pathway. Nat. Commun. 14, 4675 (2023).

pubmed: 37542026 pmcid: 10403505 doi: 10.1038/s41467-023-40435-2

Thomas, M. M. et al. Muscle-specific AMPK β1β2-null mice display a myopathy due to loss of capillary density in nonpostural muscles. FASEB J. 28, 2098–2107 (2014).

pubmed: 24522207 pmcid: 3986832 doi: 10.1096/fj.13-238972

O’Neill, H. M. et al. AMP-activated protein kinase (AMPK) β1β2 muscle null mice reveal an essential role for AMPK in maintaining mitochondrial content and glucose uptake during exercise. Proc. Natl Acad. Sci. USA 108, 16092–16097 (2011).

pubmed: 21896769 pmcid: 3179037 doi: 10.1073/pnas.1105062108

Hall, D. T. et al. The AMPK agonist 5‐aminoimidazole‐4‐carboxamide ribonucleotide (AICAR), but not metformin, prevents inflammation‐associated cachectic muscle wasting. EMBO Mol. Med. 10, e8307 (2018).

pubmed: 29844217 pmcid: 6034131 doi: 10.15252/emmm.201708307

Rohdenburg, G. L., Bernhard, A. & Krehbiel, O. Sugar tolerance in cancer. J. Am. Med. Assoc. 72, 1528–1530 (1919).

doi: 10.1001/jama.1919.02610210024007

Hwangbo, Y. et al. Incidence of diabetes after cancer development: a Korean National Cohort Study. JAMA Oncol. 4, 1099–1105 (2018).

pubmed: 29879271 pmcid: 6143049 doi: 10.1001/jamaoncol.2018.1684

Sylow, L. et al. Incidence of new-onset type 2 diabetes after cancer: a Danish Cohort Study. Diabetes Care 45, e105–e106 (2022).

pubmed: 35653597 doi: 10.2337/dc22-0232

Kjøbsted, R. et al. Enhanced muscle insulin sensitivity after contraction/exercise is mediated by AMPK. Diabetes 66, 598–612 (2017).

pubmed: 27797909 doi: 10.2337/db16-0530

Kjøbsted, R. et al. Prior AICAR stimulation increases insulin sensitivity in mouse skeletal muscle in an AMPK-dependent manner. Diabetes 64, 2042–2055 (2015).

pubmed: 25552597 doi: 10.2337/db14-1402

Pehmøller, C. et al. Genetic disruption of AMPK signaling abolishes both contraction- and insulin-stimulated TBC1D1 phosphorylation and 14-3-3 binding in mouse skeletal muscle. Am. J. Physiol. Endocrinol. Metab. 297, E665–E675 (2009).

pubmed: 19531644 pmcid: 2739697 doi: 10.1152/ajpendo.00115.2009

Kjøbsted, R. et al. TBC1D4 is necessary for enhancing muscle insulin sensitivity in response to AICAR and contraction. Diabetes 68, 1756–1766 (2019).

pubmed: 31175100 doi: 10.2337/db18-0769

Sylow, L. et al. Rac1 signaling is required for insulin-stimulated glucose uptake and is dysregulated in insulin-resistant murine and human skeletal muscle. Diabetes 62, 1865–1875 (2013).

pubmed: 23423567 pmcid: 3661612 doi: 10.2337/db12-1148

Rudich, A. & Klip, A. Putting Rac1 on the path to glucose uptake. Diabetes 62, 1831–1832 (2013).

pubmed: 23704529 pmcid: 3661647 doi: 10.2337/db13-0381

Sylow, L. et al. Akt and Rac1 signaling are jointly required for insulin-stimulated glucose uptake in skeletal muscle and downregulated in insulin resistance. Cell. Signal. 26, 323–331 (2014).

pubmed: 24216610 doi: 10.1016/j.cellsig.2013.11.007

Sylow, L. et al. Rac1 in muscle is dispensable for improved insulin action after exercise in mice. Endocrinology 157, 3009–3015 (2016).

pubmed: 27285860 doi: 10.1210/en.2016-1220

Small, L. et al. Acute activation of pyruvate dehydrogenase increases glucose oxidation in muscle without changing glucose uptake. Am. J. Physiol. Endocrinol. Metab. 315, E258–E266 (2018).

pubmed: 29406780 doi: 10.1152/ajpendo.00386.2017

Langer, H. T. et al. Restoring adiponectin via rosiglitazone ameliorates tissue wasting in mice with lung cancer. Preprint at BioRxiv https://doi.org/10.1101/2023.07.31.551241 (2023).

Frøsig, C. et al. AMPK and insulin action – responses to ageing and high fat diet. PLoS ONE 8, e62338 (2013).

pubmed: 23671593 pmcid: 3645997 doi: 10.1371/journal.pone.0062338

Fujii, N. et al. Ablation of AMP-activated protein kinase ɑ2 activity exacerbates insulin resistance induced by high-fat feeding of mice. Diabetes 57, 2958–2966 (2008).

pubmed: 18728234 pmcid: 2570392 doi: 10.2337/db07-1187

Lantier, L. et al. Reciprocity between skeletal muscle AMPK deletion and insulin action in diet-induced obese mice. Diabetes 69, 1636–1649 (2020).

pubmed: 32439824 pmcid: 7372072 doi: 10.2337/db19-1074

Johanns, M. et al. Direct and indirect activation of eukaryotic elongation factor 2 kinase by AMP-activated protein kinase. Cell. Signal. 36, 212–221 (2017).

pubmed: 28502587 doi: 10.1016/j.cellsig.2017.05.010

Gwinn, D. M. et al. AMPK phosphorylation of raptor mediates a metabolic checkpoint. Mol. Cell 30, 214–226 (2008).

pubmed: 18439900 pmcid: 2674027 doi: 10.1016/j.molcel.2008.03.003

Inoki, K., Zhu, T. & Guan, K.-L. TSC2 mediates cellular energy response to control cell growth and survival. Cell 115, 577–590 (2003).

pubmed: 14651849 doi: 10.1016/S0092-8674(03)00929-2

Bolster, D. R., Crozier, S. J., Kimball, S. R. & Jefferson, L. S. AMP-activated protein kinase suppresses protein synthesis in rat skeletal muscle through down-regulated mammalian target of rapamycin (mTOR) signaling. J. Biol. Chem. 277, 23977–23980 (2002).

pubmed: 11997383 doi: 10.1074/jbc.C200171200

Lantier, L. et al. Coordinated maintenance of muscle cell size control by AMP-activated protein kinase. FASEB J. 24, 3555–3561 (2010).

pubmed: 20460585 doi: 10.1096/fj.10-155994

Kido, K. et al. AMPK is indispensable for overload-induced muscle glucose uptake and glycogenesis but dispensable for inducing hypertrophy in mice. FASEB J. 35, e21459 (2021).

pubmed: 33710687 doi: 10.1096/fj.202002164R

Jang, T. et al. 5′‐AMP‐activated protein kinase activity is elevated early during primary brain tumor development in the rat. Int. J. Cancer 128, 2230–2239 (2011).

pubmed: 20635388 pmcid: 2992079 doi: 10.1002/ijc.25558

Godlewski, J. et al. MicroRNA-451 regulates LKB1/AMPK signaling and allows adaptation to metabolic stress in glioma cells. Mol. Cell 37, 620–632 (2010).

pubmed: 20227367 pmcid: 3125113 doi: 10.1016/j.molcel.2010.02.018

Kim, S. M. et al. PTEN deficiency and AMPK activation promote nutrient scavenging and anabolism in prostate cancer cells. Cancer Discov. 8, 866–883 (2018).

pubmed: 29572236 pmcid: 6030497 doi: 10.1158/2159-8290.CD-17-1215

Hart, P. C. et al. MnSOD upregulation sustains the Warburg effect via mitochondrial ROS and AMPK-dependent signalling in cancer. Nat. Commun. 6, 6053 (2015).

pubmed: 25651975 doi: 10.1038/ncomms7053

Eichner, L. J. et al. Genetic analysis reveals AMPK is required to support tumor growth in murine Kras-dependent lung cancer models. Cell Metab. 29, 285–302 (2019).

pubmed: 30415923 doi: 10.1016/j.cmet.2018.10.005

La Montagna, M. et al. AMPKα loss promotes KRAS-mediated lung tumorigenesis. Cell Death Differ. 28, 2673–2689 (2021).

pubmed: 34040167 pmcid: 8408205 doi: 10.1038/s41418-021-00777-0

Murray, C. W. et al. An LKB1-SIK axis suppresses lung tumor growth and controls differentiation. Cancer Discov. 9, 1590–1605 (2019).

pubmed: 31350327 pmcid: 6825558 doi: 10.1158/2159-8290.CD-18-1237

Hardie, D. G. Molecular pathways: is AMPK a friend or a foe in cancer? Clin. Cancer Res. 21, 3836–3840 (2015).

pubmed: 26152739 pmcid: 4558946 doi: 10.1158/1078-0432.CCR-14-3300

Li, W., Saud, S. M., Young, M. R., Chen, G. & Hua, B. Targeting AMPK for cancer prevention and treatment. Oncotarget 6, 7365 (2015).

pubmed: 25812084 pmcid: 4480686 doi: 10.18632/oncotarget.3629

Faubert, B., Vincent, E. E., Poffenberger, M. C. & Jones, R. G. The AMP-activated protein kinase (AMPK) and cancer: many faces of a metabolic regulator. Cancer Lett. 356, 165–170 (2015).

pubmed: 24486219 doi: 10.1016/j.canlet.2014.01.018

Luo, Z., Saha, A. K., Xiang, X. & Ruderman, N. B. AMPK, the metabolic syndrome and cancer. Trends Pharmacol. Sci. 26, 69–76 (2005).

pubmed: 15681023 doi: 10.1016/j.tips.2004.12.011

Zadra, G., Batista, J. L. & Loda, M. Dissecting the dual role of AMPK in cancer: from experimental to human studies. Mol. Cancer Res. 13, 1059–1072 (2015).

pubmed: 25956158 pmcid: 4504770 doi: 10.1158/1541-7786.MCR-15-0068

Pigna, E. et al. Aerobic exercise and pharmacological treatments counteract cachexia by modulating autophagy in colon cancer. Sci. Rep. 6, 26991 (2016).

pubmed: 27244599 pmcid: 4886631 doi: 10.1038/srep26991

Asp, M. L., Tian, M., Wendel, A. A. & Belury, M. A. Evidence for the contribution of insulin resistance to the development of cachexia in tumor-bearing mice. Int. J. Cancer 126, 756–763 (2010).

pubmed: 19634137 doi: 10.1002/ijc.24784

Chen, S.-Z. & Xiao, J.-D. Rosiglitazone and imidapril alone or in combination alleviate muscle and adipose depletion in a murine cancer cachexia model. Tumor Biol. 35, 323–332 (2014).

doi: 10.1007/s13277-013-1043-1

Trobec, K. et al. Rosiglitazone reduces body wasting and improves survival in a rat model of cancer cachexia. Nutrition 30, 1069–1075 (2014).

pubmed: 24976415 doi: 10.1016/j.nut.2013.12.005

Beluzi, M. et al. Pioglitazone treatment increases survival and prevents body weight loss in tumor-bearing animals: possible anti-cachectic effect. PLoS ONE 10, e0122660 (2015).

pubmed: 25807446 pmcid: 4373945 doi: 10.1371/journal.pone.0122660

Morinaga, M. et al. Aerobic exercise ameliorates cancer cachexia-induced muscle wasting through adiponectin signaling. Int. J. Mol. Sci. 22, 3110 (2021).

pubmed: 33803685 pmcid: 8002946 doi: 10.3390/ijms22063110

Wang, X., Pickrell, A. M., Zimmers, T. A. & Moraes, C. T. Increase in muscle mitochondrial biogenesis does not prevent muscle loss but increased tumor size in a mouse model of acute cancer-induced cachexia. PLoS ONE 7, e33426 (2012).

pubmed: 22428048 pmcid: 3299794 doi: 10.1371/journal.pone.0033426

Morena da Silva, F. et al. PGC-1α overexpression is not sufficient to mitigate cancer cachexia in either male or female mice. Appl. Physiol. Nutr. Metab. 47, 933–948 (2022).

pubmed: 35700525 pmcid: 10201462 doi: 10.1139/apnm-2022-0086

Bodine, S. C. et al. Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can prevent muscle atrophy in vivo. Nat. Cell Biol. 3, 1014–1019 (2001).

pubmed: 11715023 doi: 10.1038/ncb1101-1014

Baar, K. & Esser, K. Phosphorylation of p70

doi: 10.1152/ajpcell.1999.276.1.C120

Castets, P. et al. Sustained activation of mTORC1 in skeletal muscle inhibits constitutive and starvation-induced autophagy and causes a severe, late-onset myopathy. Cell Metab. 17, 731–744 (2013).

pubmed: 23602450 doi: 10.1016/j.cmet.2013.03.015

Joseph, G. A. et al. Partial inhibition of mTORC1 in aged rats counteracts the decline in muscle mass and reverses molecular signaling associated with sarcopenia. Mol. Cell. Biol. 39, e00141-19 (2019).

pubmed: 31308131 pmcid: 6751631 doi: 10.1128/MCB.00141-19

Ham, D. J. et al. The neuromuscular junction is a focal point of mTORC1 signaling in sarcopenia. Nat. Commun. 11, 4510 (2020).

pubmed: 32908143 pmcid: 7481251 doi: 10.1038/s41467-020-18140-1

Kim, D. H. et al. mTOR interacts with raptor to form a nutrient-sensitive complex that signals to the cell growth machinery. Cell 110, 163–175 (2002).

pubmed: 12150925 doi: 10.1016/S0092-8674(02)00808-5

Geremia, A. et al. Activation of Akt-mTORC1 signalling reverts cancer-dependent muscle wasting. J. Cachexia Sarcopenia Muscle 13, 648–661 (2022).

pubmed: 34741441 doi: 10.1002/jcsm.12854

Hawley, S. A. et al. Phosphorylation by Akt within the ST loop of AMPK-α1 down-regulates its activation in tumour cells. Biochem. J. 459, 275–287 (2014).

pubmed: 24467442 doi: 10.1042/BJ20131344

Chen, M. et al. AKT2 regulates development and metabolic homeostasis via AMPK-depedent pathway in skeletal muscle. Clin. Sci. 134, 2381–2398 (2020).

doi: 10.1042/CS20191320

Han, F. et al. The critical role of AMPK in driving Akt activation under stress, tumorigenesis and drug resistance. Nat. Commun. 9, 4728 (2018).

pubmed: 30413706 pmcid: 6226490 doi: 10.1038/s41467-018-07188-9

Kazyken, D. et al. AMPK directly activates mTORC2 to promote cell survival during acute energetic stress. Sci. Signal. 12, eaav3249 (2019).

pubmed: 31186373 pmcid: 6935248 doi: 10.1126/scisignal.aav3249

Röhrig, F. & Schulze, A. The multifaceted roles of fatty acid synthesis in cancer. Nat. Rev. Cancer 16, 732–749 (2016).

pubmed: 27658529 doi: 10.1038/nrc.2016.89

Schönke, M., Massart, J. & Zierath, J. R. Effects of high-fat diet and AMP-activated protein kinase modulation on the regulation of whole-body lipid metabolism. J. Lipid Res. 59, 1276–1282 (2018).

pubmed: 29739863 pmcid: 6027908 doi: 10.1194/jlr.D082370

Gauthier, M. S. et al. AMP-activated protein kinase is activated as a consequence of lipolysis in the adipocyte: potential mechanism and physiological relevance. J. Biol. Chem. 283, 16514–16524 (2008).

pubmed: 18390901 pmcid: 2423258 doi: 10.1074/jbc.M708177200

Mottillo, E. P. et al. Lack of adipocyte AMPK exacerbates insulin resistance and hepatic steatosis through brown and beige adipose tissue function. Cell Metab. 24, 118–129 (2016).

pubmed: 27411013 pmcid: 5239668 doi: 10.1016/j.cmet.2016.06.006

Koh, H. J. et al. Adrenaline is a critical mediator of acute exercise-induced AMP-activated protein kinase activation in adipocytes. Biochem. J. 403, 473–481 (2007).

pubmed: 17253964 pmcid: 1876380 doi: 10.1042/BJ20061479

Pulinilkunnil, T. et al. Adrenergic regulation of AMP-activated protein kinase in brown adipose tissue in vivo. J. Biol. Chem. 286, 8798–8809 (2011).

pubmed: 21209093 pmcid: 3059037 doi: 10.1074/jbc.M111.218719

Moule, S. K. & Denton, R. M. The activation of p38 MAPK by the β-adrenergic agonist isoproterenol in rat epididymal fat cells. FEBS Lett. 439, 287–290 (1998).

pubmed: 9845339 doi: 10.1016/S0014-5793(98)01392-1

Hepp, D., Challoner, D. R. & Williams, R. H. Respiration in isolated fat cells and the effects of epinephrine. J. Biol. Chem. 243, 2321–2327 (1968).

pubmed: 5648434 doi: 10.1016/S0021-9258(18)93478-2

Bihler, I. & Jeanrenaud, B. ATP content of isolated fat cells. Effects of insulin, ouabain, and lipolytic agents. Biochim. Biophys. Acta 202, 496–506 (1970).

pubmed: 4315138 doi: 10.1016/0005-2760(70)90120-7

Ahmad, B., Serpell, C. J., Fong, I. L. & Wong, E. H. Molecular mechanisms of adipogenesis: the anti-adipogenic role of AMP-activated protein kinase. Front. Mol. Biosci. 7, 76 (2020).

pubmed: 32457917 pmcid: 7226927 doi: 10.3389/fmolb.2020.00076

Vila-Bedmar, R., Lorenzo, M. & Fernández-Veledo, S. Adenosine 5’-monophosphate-activated protein kinase-mammalian target of rapamycin cross talk regulates brown adipocyte differentiation. Endocrinology 151, 980–992 (2010).

pubmed: 20133456 doi: 10.1210/en.2009-0810

Balaban, S. et al. Adipocyte lipolysis links obesity to breast cancer growth: adipocyte-derived fatty acids drive breast cancer cell proliferation and migration. Cancer Metab. 5, 1 (2017).

pubmed: 28101337 pmcid: 5237166 doi: 10.1186/s40170-016-0163-7

Das, S. K. et al. Adipose triglyceride lipase contributes to cancer-associated cachexia. Science 333, 233–238 (2011).

pubmed: 21680814 doi: 10.1126/science.1198973

Queiroz, A. L. et al. Blocking ActRIIB and restoring appetite reverses cachexia and improves survival in mice with lung cancer. Nat. Commun. 13, 4633 (2022).

pubmed: 35941104 pmcid: 9360437 doi: 10.1038/s41467-022-32135-0

Fukawa, T. et al. Excessive fatty acid oxidation induces muscle atrophy in cancer cachexia. Nat. Med. 22, 666–671 (2016).

pubmed: 27135739 doi: 10.1038/nm.4093

Kim, S. J. et al. AMPK phosphorylates desnutrin/ATGL and hormone-sensitive lipase to regulate lipolysis and fatty acid oxidation within adipose tissue. Mol. Cell. Biol. 36, 1961–1976 (2016).

pubmed: 27185873 pmcid: 4936063 doi: 10.1128/MCB.00244-16

Daval, M. et al. Anti-lipolytic action of AMP-activated protein kinase in rodent adipocytes. J. Biol. Chem. 280, 25250–25257 (2005).

pubmed: 15878856 doi: 10.1074/jbc.M414222200

Vaughan, M. The production and release of glycerol by adipose tissue incubated in vitro. J. Biol. Chem. 237, 3354–3358 (1962).

pubmed: 13996476 doi: 10.1016/S0021-9258(19)70821-7

Zhang, T., Liu, J., Tong, Q. & Lin, L. SIRT3 acts as a positive autophagy regulator to promote lipid mobilization in adipocytes via activating AMPK. Int. J. Mol. Sci. 21, 372 (2020).

pubmed: 31936019 pmcid: 7013837 doi: 10.3390/ijms21020372

Zhang, Z. et al. Berberine activates thermogenesis in white and brown adipose tissue. Nat. Commun. 5, 5493 (2014).

pubmed: 25423280 doi: 10.1038/ncomms6493

Fischer, A. W. et al. UCP1 inhibition in Cidea-overexpressing mice is physiologically counteracted by brown adipose tissue hyperrecruitment. Am. J. Physiol. Endocrinol. Metab. 312, E72–E87 (2017).

pubmed: 27923808 doi: 10.1152/ajpendo.00284.2016

Qi, J. et al. Downregulation of AMP-activated protein kinase by Cidea-mediated ubiquitination and degradation in brown adipose tissue. EMBO J. 27, 1537–1548 (2008).

pubmed: 18480843 pmcid: 2426729 doi: 10.1038/emboj.2008.92

Ji, H. et al. Development of a peptide drug restoring AMPK and adipose tissue functionality in cancer cachexia. Mol. Ther. 31, 2408–2421 (2023).

pubmed: 37408309 doi: 10.1016/j.ymthe.2023.06.020

Bing, C. et al. Adipose atrophy in cancer cachexia: morphologic and molecular analysis of adipose tissue in tumour-bearing mice. Br. J. Cancer 95, 1028–1037 (2006).

pubmed: 17047651 pmcid: 2360696 doi: 10.1038/sj.bjc.6603360

Sudo, Y. et al. Differential metabolic responses to adipose atrophy associated with cancer cachexia and caloric restriction in rats and the effect of rikkunshito in cancer cachexia. Int. J. Mol. Sci. 19, 3852 (2018).

pubmed: 30513935 pmcid: 6321026 doi: 10.3390/ijms19123852

Jeon, S. M. Regulation and function of AMPK in physiology and diseases. Exp. Mol. Med. 48, e245 (2016).

pubmed: 27416781 pmcid: 4973318 doi: 10.1038/emm.2016.81

Wojtaszewski, J. F., Nielsen, P., Hansen, B. F., Richter, E. A. & Kiens, B. Isoform-specific and exercise intensity-dependent activation of 5’-AMP-activated protein kinase in human skeletal muscle. J. Physiol. 528, 221–226 (2000).

pubmed: 11018120 pmcid: 2270117 doi: 10.1111/j.1469-7793.2000.t01-1-00221.x

Fearon, K. et al. Definition and classification of cancer cachexia: an international consensus. Lancet Oncol. 12, 489–495 (2011).

pubmed: 21296615 doi: 10.1016/S1470-2045(10)70218-7

Keys, A., Brožek, J., Henschel, A., Mickelsen, O. & Taylor, H. L. The Biology of Human Starvation Vol. 2 (University of Minnesota Press, 1950).

Sponarova, J. et al. Involvement of AMP-activated protein kinase in fat depot-specific metabolic changes during starvation. FEBS Lett. 579, 6105–6110 (2005).

pubmed: 16229840 doi: 10.1016/j.febslet.2005.09.078

Langer, H. T. et al. The proteasome regulates body weight and systemic nutrient metabolism during fasting. Am. J. Physiol. Endocrinol. Metab. 325, E500–E512 (2023).

pubmed: 37672249 pmcid: 10864006 doi: 10.1152/ajpendo.00069.2023

Storoschuk, K. L. et al. Impact of fasting on the AMPK and PGC-1α axis in rodent and human skeletal muscle: a systematic review. Metabolism 152, 155768 (2023).

pubmed: 38154612 doi: 10.1016/j.metabol.2023.155768

Nelson, M. E. et al. Phosphoproteomics reveals conserved exercise-stimulated signaling and AMPK regulation of store-operated calcium entry. EMBO J. 38, e102578 (2019).

pubmed: 31381180 pmcid: 6912027 doi: 10.15252/embj.2019102578

Dreyer, H. C. et al. Resistance exercise increases AMPK activity and reduces 4E-BP1 phosphorylation and protein synthesis in human skeletal muscle. J. Physiol. 576, 613–624 (2006).

pubmed: 16873412 pmcid: 1890364 doi: 10.1113/jphysiol.2006.113175

Wilkinson, S. B. et al. Differential effects of resistance and endurance exercise in the fed state on signalling molecule phosphorylation and protein synthesis in human muscle. J. Physiol. 586, 3701–3717 (2008).

pubmed: 18556367 pmcid: 2538832 doi: 10.1113/jphysiol.2008.153916

Langer, H. T. et al. Myofibrillar protein synthesis rates are increased in chronically exercised skeletal muscle despite decreased anabolic signaling. Sci. Rep. 12, 7553 (2022).

pubmed: 35534615 pmcid: 9085756 doi: 10.1038/s41598-022-11621-x

Wojtaszewski, J. F. et al. 5’AMP activated protein kinase expression in human skeletal muscle: effects of strength training and type 2 diabetes. J. Physiol. 564, 563–573 (2005).

pubmed: 15718261 pmcid: 1464439 doi: 10.1113/jphysiol.2005.082669

Li, M. et al. Reduced AMPK-ACC and mTOR signaling in muscle from older men, and effect of resistance exercise. Mech. Ageing Dev. 133, 655–664 (2012).

pubmed: 23000302 pmcid: 3631591 doi: 10.1016/j.mad.2012.09.001

Apró, W. et al. Resistance exercise-induced S6K1 kinase activity is not inhibited in human skeletal muscle despite prior activation of AMPK by high-intensity interval cycling. Am. J. Physiol. Endocrinol. Metab. 308, E470–E481 (2015).

pubmed: 25605643 doi: 10.1152/ajpendo.00486.2014

Apró, W., Wang, L., Pontén, M., Blomstrand, E. & Sahlin, K. Resistance exercise induced mTORC1 signaling is not impaired by subsequent endurance exercise in human skeletal muscle. Am. J. Physiol. Endocrinol. Metab. 305, E22–E32 (2013).

pubmed: 23632629 doi: 10.1152/ajpendo.00091.2013

Needham, E. J. et al. Personalized phosphoproteomics identifies functional signaling. Nat. Biotechnol. 40, 576–584 (2022).

pubmed: 34857927 doi: 10.1038/s41587-021-01099-9

Lantier, L. et al. AMPK controls exercise endurance, mitochondrial oxidative capacity, and skeletal muscle integrity. FASEB J. 28, 3211–3224 (2014).

pubmed: 24652947 doi: 10.1096/fj.14-250449

Narkar, V. A. et al. AMPK and PPARδ agonists are exercise mimetics. Cell 134, 405–415 (2008).

pubmed: 18674809 pmcid: 2706130 doi: 10.1016/j.cell.2008.06.051

Winder, W. W. et al. Activation of AMP-activated protein kinase increases mitochondrial enzymes in skeletal muscle. J. Appl. Physiol. 88, 2219–2226 (2000).

pubmed: 10846039 doi: 10.1152/jappl.2000.88.6.2219

Pedersen, L. et al. Voluntary running suppresses tumor growth through epinephrine- and IL-6-dependent NK cell mobilization and redistribution. Cell Metab. 23, 554–562 (2016).

pubmed: 26895752 doi: 10.1016/j.cmet.2016.01.011

Sweegers, M. G. et al. Effects and moderators of exercise on muscle strength, muscle function and aerobic fitness in patients with cancer: a meta-analysis of individual patient data. Br. J. Sports Med. 53, 812 (2019).

pubmed: 30181323 doi: 10.1136/bjsports-2018-099191

Bourke, L. et al. Exercise for men with prostate cancer: a systematic review and meta-analysis. Eur. Urol. 69, 693–703 (2016).

pubmed: 26632144 doi: 10.1016/j.eururo.2015.10.047

Singh, B. et al. Exercise and colorectal cancer: a systematic review and meta-analysis of exercise safety, feasibility and effectiveness. Int. J. Behav. Nutr. Phys. Act. 17, 122 (2020).

pubmed: 32972439 pmcid: 7513291 doi: 10.1186/s12966-020-01021-7

Lopez, P., Taaffe, D. R., Newton, R. U. & Galvão, D. A. Resistance exercise dosage in men with prostate cancer: systematic review, meta-analysis, and meta-regression. Med. Sci. Sports Exerc. 53, 459–469 (2021).

pubmed: 32890199 doi: 10.1249/MSS.0000000000002503

Grande, A. J. et al. Exercise for cancer cachexia in adults. Cochrane Database Syst. Rev. 3, Cd010804 (2021).

pubmed: 33735441

Hain, B. A., Xu, H. & Waning, D. L. Loss of REDD1 prevents chemotherapy‐induced muscle atrophy and weakness in mice. J. Cachexia Sarcopenia Muscle 12, 1597–1612 (2021).

pubmed: 34664403 pmcid: 8718092 doi: 10.1002/jcsm.12795

Hain, B. A. et al. REDD1 deletion attenuates cancer cachexia in mice. J. Appl. Physiol. 131, 1718–1730 (2021).

pubmed: 34672766 pmcid: 10392697 doi: 10.1152/japplphysiol.00536.2021

Hingst, J. R. et al. Inducible deletion of skeletal muscle AMPKα reveals that AMPK is required for nucleotide balance but dispensable for muscle glucose uptake and fat oxidation during exercise. Mol. Metab. 40, 101028 (2020).

pubmed: 32504885 pmcid: 7356270 doi: 10.1016/j.molmet.2020.101028

Hardie, D. G. Targeting an energy sensor to treat diabetes. Science 357, 455–456 (2017).

pubmed: 28774917 doi: 10.1126/science.aao1913

Zhou, G. et al. Role of AMP-activated protein kinase in mechanism of metformin action. J. Clin. Invest. 108, 1167–1174 (2001).

pubmed: 11602624 pmcid: 209533 doi: 10.1172/JCI13505

Bowker, S. L., Majumdar, S. R., Veugelers, P. & Johnson, J. A. Increased cancer-related mortality for patients with type 2 diabetes who use sulfonylureas or insulin. Diabetes Care 29, 254–258 (2006).

pubmed: 16443869 doi: 10.2337/diacare.29.02.06.dc05-1558

Vihervuori, V. J. et al. Antidiabetic drugs and prostate cancer prognosis in a Finnish population-based cohort. Cancer Epidemiol. Biomark. Prev. 30, 982–989 (2021).

doi: 10.1158/1055-9965.EPI-19-0580

Lee, Y. et al. Randomized phase II study of platinum-based chemotherapy plus controlled diet with or without metformin in patients with advanced non-small cell lung cancer. Lung Cancer 151, 8–15 (2021).

pubmed: 33278671 doi: 10.1016/j.lungcan.2020.11.011

Hunter, R. W. et al. Metformin reduces liver glucose production by inhibition of fructose-1-6-bisphosphatase. Nat. Med. 24, 1395–1406 (2018).

pubmed: 30150719 pmcid: 6207338 doi: 10.1038/s41591-018-0159-7

Kjøbsted, R. et al. Metformin improves glycemia independently of skeletal muscle AMPK via enhanced intestinal glucose clearance. Preprint at BioRxiv https://doi.org/10.1101/2022.05.22.492936 (2022).

Auger, C. et al. Metformin prevents the pathological browning of subcutaneous white adipose tissue. Mol. Metab. 29, 12–23 (2019).

pubmed: 31668383 pmcid: 6728757 doi: 10.1016/j.molmet.2019.08.011

Konopka, A. R. et al. Metformin inhibits mitochondrial adaptations to aerobic exercise training in older adults. Aging Cell 18, e12880 (2019).

pubmed: 30548390 doi: 10.1111/acel.12880

Oliveira, A. G. & Gomes-Marcondes, M. C. Metformin treatment modulates the tumour-induced wasting effects in muscle protein metabolism minimising the cachexia in tumour-bearing rats. BMC Cancer 16, 418 (2016).

pubmed: 27388367 pmcid: 4936094 doi: 10.1186/s12885-016-2424-9

Hawley, S. A. et al. The Na

pubmed: 27381369 doi: 10.2337/db16-0058

Brunmair, B. et al. Thiazolidinediones, like metformin, inhibit respiratory complex I: a common mechanism contributing to their antidiabetic actions? Diabetes 53, 1052–1059 (2004).

pubmed: 15047621 doi: 10.2337/diabetes.53.4.1052

LeBrasseur, N. K. et al. Thiazolidinediones can rapidly activate AMP-activated protein kinase in mammalian tissues. Am. J. Physiol. Endocrinol. Metab. 291, E175–E181 (2006).

pubmed: 16464908 doi: 10.1152/ajpendo.00453.2005

Hawley, S. A. et al. The ancient drug salicylate directly activates AMP-activated protein kinase. Science 336, 918–922 (2012).

pubmed: 22517326 pmcid: 3399766 doi: 10.1126/science.1215327

Cusi, K. et al. Efficacy and safety of PXL770, a direct AMP kinase activator, for the treatment of non-alcoholic fatty liver disease (STAMP-NAFLD): a randomised, double-blind, placebo-controlled, phase 2a study. Lancet Gastroenterol. Hepatol. 6, 889–902 (2021).

pubmed: 34560015 doi: 10.1016/S2468-1253(21)00300-9

Steneberg, P. et al. PAN-AMPK activator O304 improves glucose homeostasis and microvascular perfusion in mice and type 2 diabetes patients. JCI Insight 3, e99114 (2018).

pubmed: 29925691 pmcid: 6124394 doi: 10.1172/jci.insight.99114

Myers, R. W. et al. Systemic pan-AMPK activator MK-8722 improves glucose homeostasis but induces cardiac hypertrophy. Science 357, 507–511 (2017).

pubmed: 28705990 doi: 10.1126/science.aah5582

Cokorinos, E. C. et al. Activation of skeletal muscle AMPK promotes glucose disposal and glucose lowering in non-human primates and mice. Cell Metab. 25, 1147–1159 (2017).

pubmed: 28467931 doi: 10.1016/j.cmet.2017.04.010

Ng, S. Y. et al. Acute, next-generation AMPK activation initiates a disease-resistant gene expression program in dystrophic skeletal muscle. FASEB J. 37, e22863 (2023).

pubmed: 37016990 doi: 10.1096/fj.202201846RR

Saha, A. K. et al. Pioglitazone treatment activates AMP-activated protein kinase in rat liver and adipose tissue in vivo. Biochem. Biophys. Res. Commun. 314, 580–585 (2004).

pubmed: 14733947 doi: 10.1016/j.bbrc.2003.12.120

Maruyama, S. et al. Adiponectin ameliorates doxorubicin-induced cardiotoxicity through Akt protein-dependent mechanism. J. Biol. Chem. 286, 32790–32800 (2011).

pubmed: 21784858 pmcid: 3173230 doi: 10.1074/jbc.M111.245985

Konishi, M. et al. Adiponectin protects against doxorubicin-induced cardiomyopathy by anti-apoptotic effects through AMPK up-regulation. Cardiovasc. Res. 89, 309–319 (2011).

pubmed: 20978005 doi: 10.1093/cvr/cvq335

Lee, C. G. et al. Insulin sensitizers may attenuate lean mass loss in older men with diabetes. Diabetes Care 34, 2381–2386 (2011).

pubmed: 21926282 pmcid: 3198278 doi: 10.2337/dc11-1032

Bray, G. A. et al. Effect of pioglitazone on body composition and bone density in subjects with prediabetes in the ACT NOW trial. Diabetes Obes. Metab. 15, 931–937 (2013).

pubmed: 23551856 doi: 10.1111/dom.12099

Arad, M. et al. Constitutively active AMP kinase mutations cause glycogen storage disease mimicking hypertrophic cardiomyopathy. J. Clin. Invest. 109, 357–362 (2002).

pubmed: 11827995 pmcid: 150860 doi: 10.1172/JCI0214571

Dagorn, P. G. et al. A novel direct adenosine monophosphate kinase activator ameliorates disease progression in preclinical models of autosomal dominant polycystic kidney disease. Kidney Int. 103, 917–929 (2023).

pubmed: 36804411 doi: 10.1016/j.kint.2023.01.026

Gluais-Dagorn, P. et al. Direct AMPK activation corrects NASH in rodents through metabolic effects and direct action on inflammation and fibrogenesis. Hepatol. Commun. 6, 101–119 (2022).

pubmed: 34494384 doi: 10.1002/hep4.1799

Wu, J. et al. Chemoproteomic analysis of intertissue and interspecies isoform diversity of AMP-activated protein kinase (AMPK). J. Biol. Chem. 288, 35904–35912 (2013).

pubmed: 24187138 pmcid: 3861640 doi: 10.1074/jbc.M113.508747

Kopietz, F., Degerman, E. & Göransson, O. AMPKβ isoform expression patterns in various adipocyte models and in relation to body mass index. Front. Physiol. 13, 928964 (2022).

pubmed: 35991175 pmcid: 9386264 doi: 10.3389/fphys.2022.928964

Kopietz, F. et al. AMPK activation by A-769662 and 991 does not affect catecholamine-induced lipolysis in human adipocytes. Am. J. physiol. Endocrinol. Metab. 315, E1075–E1085 (2018).

pubmed: 30253109 doi: 10.1152/ajpendo.00110.2018

Thornton, C., Snowden, M. A. & Carling, D. Identification of a novel AMP-activated protein kinase β subunit isoform that is highly expressed in skeletal muscle. J. Biol. Chem. 273, 12443–12450 (1998).

pubmed: 9575201 doi: 10.1074/jbc.273.20.12443

Ericsson, M., Steneberg, P., Nyrén, R. & Edlund, H. AMPK activator O304 improves metabolic and cardiac function, and exercise capacity in aged mice. Commun. Biol. 4, 1306 (2021).

pubmed: 34795407 pmcid: 8602430 doi: 10.1038/s42003-021-02837-0

Yavari, A. et al. Chronic activation of γ2 AMPK induces obesity and reduces β cell function. Cell Metab. 23, 821–836 (2016).

pubmed: 27133129 pmcid: 4873618 doi: 10.1016/j.cmet.2016.04.003

Pushpakom, S. et al. Drug repurposing: progress, challenges and recommendations. Nat. Rev. Drug. Discov. 18, 41–58 (2019).

pubmed: 30310233 doi: 10.1038/nrd.2018.168

Merrill, G. F., Kurth, E. J., Hardie, D. G. & Winder, W. W. AICA riboside increases AMP-activated protein kinase, fatty acid oxidation, and glucose uptake in rat muscle. Am. J. Physiol. 273, E1107–E1112 (1997).

pubmed: 9435525

Cluzeau, T. et al. Acadesine circumvents azacitidine resistance in myelodysplastic syndrome and acute myeloid leukemia. Int. J. Mol. Sci. 21, 164 (2019).

pubmed: 31881723 pmcid: 6981810 doi: 10.3390/ijms21010164

Dzamko, N. et al. AMPK-independent pathways regulate skeletal muscle fatty acid oxidation. J. Physiol. 586, 5819–5831 (2008).

pubmed: 18845612 pmcid: 2655404 doi: 10.1113/jphysiol.2008.159814

Zhou, L. et al. Adiponectin activates AMP-activated protein kinase in muscle cells via APPL1/LKB1-dependent and phospholipase C/Ca

pubmed: 19520843 pmcid: 2755964 doi: 10.1074/jbc.M109.028357

Mao, X. et al. APPL1 binds to adiponectin receptors and mediates adiponectin signalling and function. Nat. Cell Biol. 8, 516–523 (2006).

pubmed: 16622416 doi: 10.1038/ncb1404

Balasubramanian, P. et al. Adiponectin receptor agonist AdipoRon improves skeletal muscle function in aged mice. eLife 11, e71282 (2022).

pubmed: 35297761 pmcid: 8963882 doi: 10.7554/eLife.71282

Selvais, C. M. et al. AdipoRon enhances healthspan in middle-aged obese mice: striking alleviation of myosteatosis and muscle degenerative markers. J. Cachexia Sarcopenia Muscle 14, 464–478 (2023).

pubmed: 36513619 doi: 10.1002/jcsm.13148

Abou-Samra, M. et al. AdipoRon, a new therapeutic prospect for Duchenne muscular dystrophy. J. Cachexia Sarcopenia Muscle 11, 518–533 (2020).

pubmed: 31965757 pmcid: 7113498 doi: 10.1002/jcsm.12531

Feng, D. et al. Discovery of MK-8722: a systemic, direct pan-activator of AMP-activated protein kinase. ACS Med. Chem. Lett. 9, 39–44 (2018).

pubmed: 29348809 doi: 10.1021/acsmedchemlett.7b00417

Calabrese, M. F. et al. Structural basis for AMPK activation: natural and synthetic ligands regulate kinase activity from opposite poles by different molecular mechanisms. Structure 22, 1161–1172 (2014).

pubmed: 25066137 doi: 10.1016/j.str.2014.06.009

Quinn, B. J., Kitagawa, H., Memmott, R. M., Gills, J. J. & Dennis, P. A. Repositioning metformin for cancer prevention and treatment. Trends Endocrinol. Metab. 24, 469–480 (2013).

pubmed: 23773243 doi: 10.1016/j.tem.2013.05.004

Hawley, S. A. et al. Characterization of the AMP-activated protein kinase kinase from rat liver and identification of threonine 172 as the major site at which it phosphorylates AMP-activated protein kinase. J. Biol. Chem. 271, 27879–27887 (1996).

pubmed: 8910387 doi: 10.1074/jbc.271.44.27879

Drake, J. C. et al. Mitochondria-localized AMPK responds to local energetics and contributes to exercise and energetic stress-induced mitophagy. Proc. Natl Acad. Sci. USA 118, e2025932118 (2021).

pubmed: 34493662 pmcid: 8449344 doi: 10.1073/pnas.2025932118

Hardie, D. G. & Hawley, S. A. AMP-activated protein kinase: the energy charge hypothesis revisited. BioEssays 23, 1112–1119 (2001).

pubmed: 11746230 doi: 10.1002/bies.10009

Lizcano, J. M. et al. LKB1 is a master kinase that activates 13 kinases of the AMPK subfamily, including MARK/PAR-1. EMBO J. 23, 833–843 (2004).

pubmed: 14976552 pmcid: 381014 doi: 10.1038/sj.emboj.7600110

Green, M. F., Anderson, K. A. & Means, A. R. Characterization of the CaMKKβ–AMPK signaling complex. Cell. Signal. 23, 2005–2012 (2011).

pubmed: 21807092 pmcid: 3184326 doi: 10.1016/j.cellsig.2011.07.014

Vara-Ciruelos, D. et al. Genotoxic damage activates the AMPK-α1 isoform in the nucleus via Ca

pubmed: 29133590 doi: 10.1158/1541-7786.MCR-17-0323

Negoita, F. et al. CaMKK2 is not involved in contraction-stimulated AMPK activation and glucose uptake in skeletal muscle. Mol. Metab. 75, 101761 (2023).

pubmed: 37380024 pmcid: 10362367 doi: 10.1016/j.molmet.2023.101761

Yang, Z., Kahn, B. B., Shi, H. & Xue, B. Z. Macrophage ɑ1 AMP-activated protein kinase (ɑ1AMPK) antagonizes fatty acid-induced inflammation through SIRT1. J. Biol. Chem. 285, 19051–19059 (2010).

pubmed: 20421294 pmcid: 2885183 doi: 10.1074/jbc.M110.123620

Iwabu, M. et al. Adiponectin and AdipoR1 regulate PGC-1ɑ and mitochondria by Ca

pubmed: 20357764 doi: 10.1038/nature08991

Minokoshi, Y. et al. Leptin stimulates fatty-acid oxidation by activating AMP-activated protein kinase. Nature 415, 339–343 (2002).

pubmed: 11797013 doi: 10.1038/415339a

White, J. P. et al. The regulation of skeletal muscle protein turnover during the progression of cancer cachexia in the Apc

pubmed: 21949739 pmcid: 3176277 doi: 10.1371/journal.pone.0024650

Sylow, L., Kleinert, M., Richter, E. A. & Jensen, T. E. Exercise-stimulated glucose uptake – regulation and implications for glycaemic control. Nat. Rev. Endocrinol. 13, 133–148 (2017).

pubmed: 27739515 doi: 10.1038/nrendo.2016.162

Warburg, O. P., Posener, K. & Negelein, E. Über den Stoffwechsel der Carcinomzelle [German]. Biochem. Z. 152, 309–344 (1924).

Vander Heiden, M. G., Cantley, L. C. & Thompson, C. B. Understanding the Warburg effect: the metabolic requirements of cell proliferation. Science 324, 1029–1033 (2009).

doi: 10.1126/science.1160809

Calle, E. E., Rodriguez, C., Walker-Thurmond, K. & Thun, M. J. Overweight, obesity, and mortality from cancer in a prospectively studied cohort of U.S. adults. N. Engl. J. Med. 348, 1625–1638 (2003).

pubmed: 12711737 doi: 10.1056/NEJMoa021423

Barone, B. B. et al. Long-term all-cause mortality in cancer patients with preexisting diabetes mellitus: a systematic review and meta-analysis. JAMA 300, 2754–2764 (2008).

pubmed: 19088353 pmcid: 3093051 doi: 10.1001/jama.2008.824

Renehan, A. G., Zwahlen, M. & Egger, M. Adiposity and cancer risk: new mechanistic insights from epidemiology. Nat. Rev. Cancer 15, 484–498 (2015).

pubmed: 26205341 doi: 10.1038/nrc3967

Park, J., Morley, T. S., Kim, M., Clegg, D. J. & Scherer, P. E. Obesity and cancer – mechanisms underlying tumour progression and recurrence. Nat. Rev. Endocrinol. 10, 455–465 (2014).

pubmed: 24935119 pmcid: 4374431 doi: 10.1038/nrendo.2014.94

Zhong, W. & Mao, Y. Daily insulin dose and cancer risk among patients with type 1 diabetes. JAMA Oncol. 8, 1356–1358 (2022).

pubmed: 35900757 pmcid: 9335241 doi: 10.1001/jamaoncol.2022.2960

Goodwin, P. J. et al. Fasting insulin and outcome in early-stage breast cancer: results of a prospective cohort study. J. Clin. Oncol. 20, 42–51 (2002).

pubmed: 11773152 doi: 10.1200/JCO.2002.20.1.42

Zhang, A. M. Y. et al. Endogenous hyperinsulinemia contributes to pancreatic cancer development. Cell Metab. 30, 403–404 (2019).

pubmed: 31378465 doi: 10.1016/j.cmet.2019.07.003

Chovsepian, A. et al. Diabetes increases mortality in patients with pancreatic and colorectal cancer by promoting cachexia and its associated inflammatory status. Mol. Metab. 73, 101729 (2023).

pubmed: 37094629 pmcid: 10192649 doi: 10.1016/j.molmet.2023.101729

Seki, T. et al. Brown-fat-mediated tumour suppression by cold-altered global metabolism. Nature 608, 421–428 (2022).

pubmed: 35922508 pmcid: 9365697 doi: 10.1038/s41586-022-05030-3

Orava, J. et al. Different metabolic responses of human brown adipose tissue to activation by cold and insulin. Cell Metab. 14, 272–279 (2011).

pubmed: 21803297 doi: 10.1016/j.cmet.2011.06.012

Jung, S. M. et al. In vivo isotope tracing reveals the versatility of glucose as a brown adipose tissue substrate. Cell Rep. 36, 109459 (2021).

pubmed: 34320357 pmcid: 8369932 doi: 10.1016/j.celrep.2021.109459

Blondin, D. P. et al. Inhibition of intracellular triglyceride lipolysis suppresses cold-induced brown adipose tissue metabolism and increases shivering in humans. Cell Metab. 25, 438–447 (2017).

pubmed: 28089568 doi: 10.1016/j.cmet.2016.12.005

Kennedy, J. W. et al. Acute exercise induces GLUT4 translocation in skeletal muscle of normal human subjects and subjects with type 2 diabetes. Diabetes 48, 1192–1197 (1999).

pubmed: 10331428 doi: 10.2337/diabetes.48.5.1192

Mikines, K. J., Sonne, B., Farrell, P. A., Tronier, B. & Galbo, H. Effect of physical exercise on sensitivity and responsiveness to insulin in humans. Am. J. Physiol. 254, E248–E259 (1988).

pubmed: 3126668

AMPK as a mediator of tissue preservation: time for a shift in dogma?

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Henning Tim Langer (HT)

Maria Rohm (M)

Marcus DaSilva Goncalves (MD)

Lykke Sylow (L)

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