Does Amyotrophic Lateral Sclerosis (ALS) Have Metabolic Causes from Human Evolution?
Abstract
1. Introduction
2. Evolution and Glycosphingolipids, Viruses and Metabolism
3. Glycosphingolipids and Gangliosides in ALS
4. Cholera-Toxin Binding as a Tool to Study GM1, and Fucosylated Structures, and to Cause Denervation
5. Is GM1 Directly Implicated in ALS?
6. Links with BDNF
7. Ceramides and ALS
8. Key Role for Lipid and Lactate Metabolism in Human Evolution and ALS
9. Breakthroughs in Lactate Metabolism in Sports Science Which May Be Relevant to ALS
10. Lactate Shuttles and Energy Control
11. Lactate in Human Performance, Ageing and in ALS
12. Previous Exercise as a Risk for ALS: Subjects with C9orf72 Expansions May Be Outliers?
13. Future Directions for Research and Therapies
- The critical path for glycosphingolipid and ganglioside pathways is under-researched, especially as these pathways provide useful biomarkers. Much of the research of the effects of GM1 have involved TrkA, yet effects on TrkB in lipid rafts may also be important. Furthermore, which the effects of Neu5Ac-GM1 and its oligosaccharide have been well studied, direct comparisons with Neu5Gc-GM1 are lacking.While GM1 is a critical component of the inner nuclear membrane, associated with the Na+/Ca2+ exchanger [136], there are no studies on whether it has an impact on TDP-43 accumulation in the cytoplasm.
- The pivotal role of GCS, GBA1 and GBA2 has indicated ways forward which are under scrutiny. While inhibitors of the super-enhancer gene UGCG (GCS) have proven deleterious in ALS models [65,67], this has increased focus on GBA2 inhibitors, several of which are in development. The use of CTB, to evaluate GM1 and related structures, will be important in the future, as will the use of CTB-S to cause very specific denervation of motor neurons. The effects of GBA2 inhibitors as glucosyltransferase inhibitors, thereby glycosylating cholesterol in lipid rafts, may impact lipid raft stability with major implications. The status of the GBA2 inhibitor/GBA1 chaperone, ambroxol, has been reviewed above. However, most GBA2 inhibitors are in development for Parkinson’s disease: ALU1811 is in preclinical development by Biogen and Alectos for Parkinson’s disease, and the GCS and GBA2 inhibitor nizubaglustat is in development for PD [326]. While miglustat is predominantly an inhibitor of GCS, its inhibition of GBA2 has been speculated to be responsible for lack of deleterious effects [327]. Endogenous surfactants, such as saposin-C, may cause profound changes in GBA1 and GBA2 activity and drugs such as ambroxol increase saposin-C levels [328]. The very recent discovery that GCS is a super-enhancer [70] means that much research will be necessary to redefine the effects of GCS inhibitors.
- The breakthroughs in knowledge about the critical role of lactate in human performance should be applied to ALS research. The studies reviewed above do not provide definitive answers as to whether Ra or Rd are specifically affected in ALS and whether peripheral lactate may be neuroprotective. Sports scientists have perfected dried blood-spot collection, which can be easily transported, to assess lactate, carboxylic acids, fatty acids and acyl carnitines regularly in racing cyclists [181], and this relatively simple technology could be used to assess the metabolic position and progression of ALS patients. Specific low-impact training, tailored to each patient, to down-regulate lipid metabolism, and increase glucose metabolism and mitochondrial function has been recently reviewed and proposed [315], as is the case with athletes (Figure 4). However, as lactate can be increased by mitochondrial dysfunction, but reduced by denervation, conclusions can only be reached with protocols defining MLSS or lactate at exhaustion. So, can a programme like phase 2 training (low intensity training in cycling), to increase mitochondrial efficiency and lactate metabolism (Figure 4), restore glycolytic activity, and have an effect in ALS on lactate use and energy restitution, with perhaps protection against further spreading denervation? Patients with C9orf72 mutations (and probably patients with other highly penetrative mutations for which exercise is a ‘stressor’) may not benefit in comparison with sporadic patients. Intensive cycling has a low risk of ALS [298], despite requiring more hours of exercise than other high-intensity sports. Light cycling may produce the metabolic resilience and improved lactate metabolism associated with stage 2 training. But how can exercise be performed at the beginning of denervation? There are now available motorized exercise bikes, such as Motomed (https://www.motomed.com/en/, accessed on 1 October 2025.) allowing muscles to be stretched, doing minimal work without applying force, with spasm movement protectors. Lactate is increased even with passive cycling [329] and it seems important to perform trials of prolonged passive cycling (or powered arm exercises)/day with measures of lactate use [330,331]. This may allow the muscles to maintain measures of mitochondrial efficiency and produce lactate, even when denervated or denervating. Recently, dynamic cycling sessions have been shown to ameliorate subthalamic function in patients with Parkinson’s disease [332]. Spectacular results have been found using physiotherapy in dogs with degenerative myopathy (diagnosis at ~9 years and a mean survival time of 55 days) [333]: moderate physiotherapy increased survival to 130 days, and intensive physiotherapy to 255 days. Remote ischaemic conditioning has been used to increase lactate in a variety of disorders, by inflating blood pressure cuffs to 20 mm Hg above systolic pressure for four 5-min occlusions in affected limbs.
- Viral infections prior to the development of ALS. While viral infections have not been associated with ALS, the entry and exit of enveloped viruses are dependent on the same GSLs and GM1, and the associated enzymes. GCS, as a super-enhancer, is up-regulated following viral infection [70]. Viral infections also remodel lipid metabolism in similar ways to ALS, but via remodelling endoplasmic reticulum for replication and creation of viral envelopes [334]. It is only recently that multiple sclerosis has been linked to prior Epstein-Barr virus infection, despite an odds-ratio of ~30 [335]. As ALS may develop late after injury, prior viral infection would be missed.
- Combination with therapeutic agents may also be beneficial in order to promote reinnervation or protect from denervation. Therapeutic approaches targeting skeletal muscle, with efficacy in preclinical models, have been recently reviewed [336]. Drugs changing GSL metabolism are an obvious target [337]. However, some agents which had been developed for changing cardiac metabolism are being examined for ALS in the clinic. Thirty five years ago, switching cardiac lipid oxidation to glucose fuels was a major drug discovery topic, as ischaemia prevented full oxidation of lipids causing build-up of acylcarnitines and lysophospholipids [338,339,340]. Ranolazine is cardioprotective by effects on pyruvate dehydrogenase and reducing excess lipid oxidation [341,342]. The drug is being developed for cramps in ALS (NCT06527222). Trimetazidine [90,343] is a forty-year old antianginal drug, which modulates mitochondrial metabolism and inhibits long-chain mitochondrial 3-ketoacyl coenzyme A thiolase, a key enzyme in lipid oxidation [344,345]: the drug has been banned from use in sports because of extensive doping allegations linked to its effects on cardiac energy metabolism [346]. Trimetazidine extended survival of SOD1G93A mice and protected NMJs reducing motor neuron loss [343]. The drug is being studied for effects on metabolic flexibility in ALS (NCT04788745), although it is contra-indicated in Parkinson’s disease [347]. These metabolic approaches may be particularly effective coupled to personalized exercise programs. However, as these drugs are being repurposed, development is not easy [348] and a coordinated approach by the ALS community and sponsors may be necessary for registration if clinical trials are positive.
14. Conclusions
Funding
Data Availability Statement
Acknowledgments
Conflicts of Interest
Abbreviations
| LDH | lactate dehydrogenase |
| LacCer | lactosylceramide |
| ERK | extracellular signal-regulated kinase |
| CPT1 | carnitine-palmitoyl transferase1 |
| CMAH | cytidine monophospho-N-acetylneuraminic acid hydroxylase |
| CTB-S | cholera toxin β-subunit bound to saporin |
| COPT | chronic obstructive pulmonary disease |
| CTB | cholera toxin β-subunit |
| BDNF | brain-derived neurotrophic factor |
| Bcl-2 | B-cell lymphoma-2 |
| ALSFRS-R | Amyotrophic lateral sclerosis functiona rating scale-revised |
| ALS | Amyotrophic lateral sclerosis |
| iPSC | induced pluripotent stem cells |
| HILIC–ESI–MS/MS | hydrophilic interaction, electrospray ionization tandem mass spectrometry |
| HCA1 | hydroxycarboxylic acid-1 receptor |
| GSL | glycosphingolipid |
| GM1 | monosialotetrahexosylganglioside |
| GlcNAc | N-acetylglucosamine |
| GCS (or UGCG) | Ceramide glucosyltransferase |
| GBD | Ganglioside-binding domain |
| GBA2 | non-lysosomal β-glucosylceramidase |
| GBA1 | lysosomal β-glucosylceramidase |
| GalNAc | N-acetylgalactosamine |
| FUS | fused in sarcoma RNA-binding protein |
| GlcCer | glucosyl ceramide |
| GalCer | galactosyl ceramide |
| FGF | fibroblast growth factor |
| FAPP2 | phosphatidylinositol-four-phosphate adapter protein 2 |
| MAPK | mitogen-activated protein kinases |
| MLSS | maximal lactate steady-state |
| NGF | nerve growth factor |
| NMJ | neuromuscular junction |
| MAPK | mitogen-activated protein kinase |
| MCT | monocarboxylate transporter isoforms |
| MEP | motor evoked potential |
| MND | motor neuron disease |
| Mya | million years ago |
| Neu5Ac | N-acetylneuraminic acid |
| NfL | neurofilament light chain |
| NGF | nerve growth factor, NSC: neural stem cell |
| ORMDL | sphingolipid biosynthesis regulator |
| pAMPK | phosphorylated AMP-activated protein kinase |
| PGC1a | peroxisome proliferator-activated receptor gamma coactivator 1-alpha |
| PPAR | peroxisome proliferator-activated receptor coactivator |
| Ra | lactate production rate |
| Rd | lactate metabolism rate |
| RCI | mitochondrial respiratory coupling index |
| SICI | short interval intracortical inhibition |
| Siglecs | sialic acid-binding immunoglobulin-type lectins |
| SOD1 | superoxide dismutase 1 |
| SPT | serine palmitoyl transferase |
| SSEA | stage-specific embryonic antigen |
| TDP-43 | transactive response DNA binding protein of 43 kDa |
| TrkA | tropomyosin receptor kinase A |
| TrkB | tropomyosin receptor kinase B |
| UGT8 | ceramide galactosyltransferase |
| VDAC1 | voltage-dependent anion channel 1 |
References
- Wolfson, C.; Gauvin, D.E.; Ishola, F.; Oskoui, M. Global Prevalence and Incidence of Amyotrophic Lateral Sclerosis: A Systematic Review. Neurology 2023, 101, e613–e623. [Google Scholar] [CrossRef]
- Pattle, S.B.; O’Shaughnessy, J.; Kantelberg, O.; Rifai, O.M.; Pate, J.; Nellany, K.; Hays, N.; Arends, M.J.; Horrocks, M.H.; Waldron, F.M.; et al. pTDP-43 aggregates accumulate in non-central nervous system tissues prior to symptom onset in amyotrophic lateral sclerosis: A case series linking archival surgical biopsies with clinical phenotypic data. J. Pathol. Clin. Res. 2023, 9, 44–55. [Google Scholar] [CrossRef]
- Smith, S.E.; McCoy-Gross, K.; Malcolm, A.; Oranski, J.; Markway, J.W.; Miller, T.M.; Bucelli, R.C. Tofersen treatment leads to sustained stabilization of disease in SOD1 ALS in a “real-world” setting. Ann. Clin. Transl. Neurol. 2025, 12, 311–319. [Google Scholar] [CrossRef]
- Miller, T.M.; Cudkowicz, M.E.; Genge, A.; Shaw, P.J.; Sobue, G.; Bucelli, R.C.; Chiò, A.; Damme, P.V.; Ludolph, A.C.; Glass, J.D.; et al. Trial of Antisense Oligonucleotide Tofersen for SOD1 ALS. N. Engl. J. Med. 2022, 387, 1099–1110. [Google Scholar] [CrossRef]
- Korobeynikov, V.A.; Lyashchenko, A.K.; Blanco-Redondo, B.; Jafar-Nejad, P.; Shneider, N.A. Antisense oligonucleotide silencing of FUS expression as a therapeutic approach in amyotrophic lateral sclerosis. Nat. Med. 2022, 28, 104–116. [Google Scholar] [CrossRef]
- Wong, C.; Stavrou, M.; Elliott, E.; Gregory, J.M.; Leigh, N.; Pinto, A.A.; Williams, T.L.; Chataway, J.; Swingler, R.; Parmar, M.K.B.; et al. Clinical trials in amyotrophic lateral sclerosis: A systematic review and perspective. Brain Commun. 2021, 3, fcab242. [Google Scholar] [CrossRef] [PubMed]
- Brooks, B.R.; Miller, R.G.; Swash, M.; Munsat, T.L. World Federation of Neurology Research Group on Motor Neuron Diseases El Escorial revisited: Revised criteria for the diagnosis of amyotrophic lateral sclerosis. Amyotroph. Lateral Scler. Other Motor Neuron Disord. 2000, 1, 293–299. [Google Scholar] [CrossRef] [PubMed]
- Vucic, S.; Ferguson, T.A.; Cummings, C.; Hotchkin, M.T.; Genge, A.; Glanzman, R.; Roet, K.C.D.; Cudkowicz, M.; Kiernan, M.C. Gold Coast diagnostic criteria: Implications for ALS diagnosis and clinical trial enrollment. Muscle Nerve 2021, 64, 532–537. [Google Scholar] [CrossRef]
- Benatar, M.; Wuu, J.; Huey, E.D.; McMillan, C.T.; Petersen, R.C.; Postuma, R.; McHutchison, C.; Dratch, L.; Arias, J.J.; Crawley, A.; et al. The Miami Framework for ALS and related neurodegenerative disorders: An integrated view of phenotype and biology. Nat. Rev. Neurol. 2024, 20, 364–376. [Google Scholar] [CrossRef]
- Al-Chalabi, A.; Calvo, A.; Chio, A.; Colville, S.; Ellis, C.M.; Hardiman, O.; Heverin, M.; Howard, R.S.; Huisman, M.H.B.; Keren, N.; et al. Analysis of amyotrophic lateral sclerosis as a multistep process: A population-based modelling study. Lancet Neurol. 2014, 13, 1108–1113. [Google Scholar] [CrossRef] [PubMed]
- Chiò, A.; Mazzini, L.; D’Alfonso, S.; Corrado, L.; Canosa, A.; Moglia, C.; Manera, U.; Bersano, E.; Brunetti, M.; Barberis, M.; et al. The multistep hypothesis of ALS revisited: The role of genetic mutations. Neurology 2018, 91, e635–e642. [Google Scholar] [CrossRef]
- Scott, S.; Kranz, J.E.; Cole, J.; Lincecum, J.M.; Thompson, K.; Kelly, N.; Bostrom, A.; Theodoss, J.; Al-Nakhala, B.M.; Vieira, F.G.; et al. Design, power, and interpretation of studies in the standard murine model of ALS. Amyotroph. Lateral Scler. 2008, 9, 4–15. [Google Scholar] [CrossRef]
- Ghasemi, M.; Brown, R.H. Genetics of Amyotrophic Lateral Sclerosis. Cold Spring Harb. Perspect. Med. 2018, 8, a024125. [Google Scholar] [CrossRef] [PubMed]
- Zhou, L.; Xie, M.; Wang, X.; Xu, R. The usage and advantages of several common amyotrophic lateral sclerosis animal models. Front. Neurosci. 2024, 18, 1341109. [Google Scholar] [CrossRef]
- Awano, T.; Johnson, G.S.; Wade, C.M.; Katz, M.L.; Johnson, G.C.; Taylor, J.F.; Perloski, M.; Biagi, T.; Baranowska, I.; Long, S.; et al. Genome-wide association analysis reveals a SOD1 mutation in canine degenerative myelopathy that resembles amyotrophic lateral sclerosis. Proc. Natl. Acad. Sci. USA 2009, 106, 2794–2799. [Google Scholar] [CrossRef] [PubMed]
- Varki, A. Colloquium paper: Uniquely human evolution of sialic acid genetics and biology. Proc. Natl. Acad. Sci. USA 2010, 107 (Suppl. S2), 8939–8946. [Google Scholar] [CrossRef]
- Sasmal, A.; Khan, N.; Khedri, Z.; Kellman, B.P.; Srivastava, S.; Verhagen, A.; Yu, H.; Bruntse, A.B.; Diaz, S.; Varki, N.; et al. Simple and practical sialoglycan encoding system reveals vast diversity in nature and identifies a universal sialoglycan-recognizing probe derived from AB5 toxin B subunits. Glycobiology 2022, 32, 1101–1115. [Google Scholar] [CrossRef] [PubMed]
- Cohen, M.; Varki, A. The Sialome—Far More Than the Sum of Its Parts. OMICS J. Integr. Biol. 2010, 14, 455–464. [Google Scholar] [CrossRef]
- Laine, R.A. A calculation of all possible oligosaccharide isomers both branched and linear yields 1.05 × 10(12) structures for a reducing hexasaccharide: The Isomer Barrier to development of single-method saccharide sequencing or synthesis systems. Glycobiology 1994, 4, 759–767. [Google Scholar] [CrossRef]
- Guo, Z. Ganglioside GM1 and the Central Nervous System. Int. J. Mol. Sci. 2023, 24, 9558. [Google Scholar] [CrossRef]
- Schengrund, C.-L. Sphingolipids: Less Enigmatic but Still Many Questions about the Role(s) of Ceramide in the Synthesis/Function of the Ganglioside Class of Glycosphingolipids. Int. J. Mol. Sci. 2024, 25, 6312. [Google Scholar] [CrossRef] [PubMed]
- Guo, Z. The Structural Diversity of Natural Glycosphingolipids (GSLs). J. Carbohydr. Chem. 2022, 41, 63–154. [Google Scholar] [CrossRef]
- Crocker, P.R.; Paulson, J.C.; Varki, A. Siglecs and their roles in the immune system. Nat. Rev. Immunol. 2007, 7, 255–266. [Google Scholar] [CrossRef] [PubMed]
- Schwarz, F.; Fong, J.J.; Varki, A. Human-specific evolutionary changes in the biology of siglecs. Adv. Exp. Med. Biol. 2015, 842, 1–16. [Google Scholar] [CrossRef]
- Irie, A.; Koyama, S.; Kozutsumi, Y.; Kawasaki, T.; Suzuki, A. The Molecular Basis for the Absence ofN-Glycolylneuraminic Acid in Humans. J. Biol. Chem. 1998, 273, 15866–15871. [Google Scholar] [CrossRef]
- Peri, S.; Kulkarni, A.; Feyertag, F.; Berninsone, P.M.; Alvarez-Ponce, D. Phylogenetic Distribution of CMP-Neu5Ac Hydroxylase (CMAH), the Enzyme Synthetizing the Proinflammatory Human Xenoantigen Neu5Gc. Genome Biol. Evol. 2017, 10, 207–219. [Google Scholar] [CrossRef]
- Chou, H.H.; Takematsu, H.; Diaz, S.; Iber, J.; Nickerson, E.; Wright, K.L.; Muchmore, E.A.; Nelson, D.L.; Warren, S.T.; Varki, A. A mutation in human CMP-sialic acid hydroxylase occurred after the Homo-Pan divergence. Proc. Natl. Acad. Sci. USA 1998, 95, 11751–11756. [Google Scholar] [CrossRef]
- Dunker, K.; Pedersen, K.M.; Toraskar, S.; Diaz, S.; Varki, A.; Sletmoen, M.; Kikkeri, R. Human-specific evolutionary genetic loss of addition of a single oxygen atom from sialic acids increases hydrophobicity of cells and proteins. Carbohydr. Res. 2025, 552, 109469. [Google Scholar] [CrossRef]
- Li, Y.; Li, Y.; Gu, X.; Liu, Y.; Dong, D.; Kang, J.H.; Wang, M.; Eliassen, H.; Willett, W.C.; Stampfer, M.J.; et al. Long-Term Intake of Red Meat in Relation to Dementia Risk and Cognitive Function in US Adults. Neurology 2025, 104, e210286. [Google Scholar] [CrossRef]
- Naito-Matsui, Y.; Davies, L.R.L.; Takematsu, H.; Chou, H.-H.; Tangvoranuntakul, P.; Carlin, A.F.; Verhagen, A.; Heyser, C.J.; Yoo, S.-W.; Choudhury, B.; et al. Physiological Exploration of the Long Term Evolutionary Selection against Expression of N-Glycolylneuraminic Acid in the Brain. J. Biol. Chem. 2017, 292, 2557–2570. [Google Scholar] [CrossRef] [PubMed]
- Chou, H.-H.; Hayakawa, T.; Diaz, S.; Krings, M.; Indriati, E.; Leakey, M.; Paabo, S.; Satta, Y.; Takahata, N.; Varki, A. Inactivation of CMP-N-acetylneuraminic acid hydroxylase occurred prior to brain expansion during human evolution. Proc. Natl. Acad. Sci. USA 2002, 99, 11736–11741. [Google Scholar] [CrossRef]
- Srivastava, S.; Verhagen, A.; Sasmal, A.; Wasik, B.R.; Diaz, S.; Yu, H.; Bensing, B.A.; Khan, N.; Khedri, Z.; Secrest, P.; et al. Development and applications of sialoglycan-recognizing probes (SGRPs) with defined specificities: Exploring the dynamic mammalian sialoglycome. Glycobiology 2022, 32, 1116–1136. [Google Scholar] [CrossRef]
- Khan, N.; de Manuel, M.; Peyregne, S.; Do, R.; Prufer, K.; Marques-Bonet, T.; Varki, N.; Gagneux, P.; Varki, A. Multiple Genomic Events Altering Hominin SIGLEC Biology and Innate Immunity Predated the Common Ancestor of Humans and Archaic Hominins. Genome Biol. Evol. 2020, 12, 1040–1050. [Google Scholar] [CrossRef]
- Varki, A. Multiple changes in sialic acid biology during human evolution. Glycoconj. J. 2009, 26, 231–245. [Google Scholar] [CrossRef] [PubMed]
- Chen, Z.; Reynolds, R.H.; Pardiñas, A.F.; Gagliano Taliun, S.A.; van Rheenen, W.; Lin, K.; Shatunov, A.; Gustavsson, E.K.; Fogh, I.; Jones, A.R.; et al. The contribution of Neanderthal introgression and natural selection to neurodegenerative diseases. Neurobiol. Dis. 2023, 180, 106082. [Google Scholar] [CrossRef]
- Miyagi, T.; Yamaguchi, K. Mammalian sialidases: Physiological and pathological roles in cellular functions. Glycobiology 2012, 22, 880–896. [Google Scholar] [CrossRef]
- Pan, X.; De Aragão, C.D.B.P.; Velasco-Martin, J.P.; Priestman, D.A.; Wu, H.Y.; Takahashi, K.; Yamaguchi, K.; Sturiale, L.; Garozzo, D.; Platt, F.M.; et al. Neuraminidases 3 and 4 regulate neuronal function by catabolizing brain gangliosides. FASEB J. 2017, 31, 3467–3483. [Google Scholar] [CrossRef]
- Pronker, M.F.; Lemstra, S.; Snijder, J.; Heck, A.J.R.; Thies-Weesie, D.M.E.; Pasterkamp, R.J.; Janssen, B.J.C. Structural basis of myelin-associated glycoprotein adhesion and signalling. Nat. Commun. 2016, 7, 13584. [Google Scholar] [CrossRef] [PubMed]
- Suzuki, K.G.N.; Ando, H.; Komura, N.; Fujiwara, T.K.; Kiso, M.; Kusumi, A. Development of new ganglioside probes and unraveling of raft domain structure by single-molecule imaging. Biochim. Biophys. Acta (BBA)—General Subj. 2017, 1861, 2494–2506. [Google Scholar] [CrossRef] [PubMed]
- Nguyen, L.; McCord, K.A.; Bui, D.T.; Bouwman, K.M.; Kitova, E.N.; Elaish, M.; Kumawat, D.; Daskhan, G.C.; Tomris, I.; Han, L.; et al. Sialic acid-containing glycolipids mediate binding and viral entry of SARS-CoV-2. Nat. Chem. Biol. 2022, 18, 81–90. [Google Scholar] [CrossRef]
- Dey, M.; Sharma, A.; Dhanawat, G.; Gupta, D.; Harshan, K.H.; Parveen, N. Synergistic Binding of SARS-CoV-2 to ACE2 and Gangliosides in Native Lipid Membranes. ACS Infect. Dis. 2024, 10, 907–916. [Google Scholar] [CrossRef]
- Negi, G.; Sharma, A.; Chaudhary, M.; Gupta, D.; Harshan, K.H.; Parveen, N. SARS-CoV-2 Binding to Terminal Sialic Acid of Gangliosides Embedded in Lipid Membranes. ACS Infect. Dis. 2023, 9, 1346–1361. [Google Scholar] [CrossRef]
- Van Blerkom, L.M. Role of viruses in human evolution. Am. J. Phys. Anthropol. 2003, 122 (Suppl. S37), 14–46. [Google Scholar] [CrossRef]
- Schneider-Schaulies, J.; Schneider-Schaulies, S. Sphingolipids in viral infection. Biol. Chem. 2015, 396, 585–595. [Google Scholar] [CrossRef]
- Schneider-Schaulies, S.; Schumacher, F.; Wigger, D.; Schöl, M.; Waghmare, T.; Schlegel, J.; Seibel, J.; Kleuser, B. Sphingolipids: Effectors and Achilles Heals in Viral Infections? Cells 2021, 10, 2175. [Google Scholar] [CrossRef] [PubMed]
- Drews, K.; Calgi, M.P.; Harrison, W.C.; Drews, C.M.; Costa-Pinheiro, P.; Shaw, J.J.P.; Jobe, K.A.; Nelson, E.A.; Han, J.D.; Fox, T.; et al. Glucosylceramidase Maintains Influenza Virus Infection by Regulating Endocytosis. J. Virol. 2019, 93, e00017-19. [Google Scholar] [CrossRef] [PubMed]
- Drews, K.; Calgi, M.P.; Harrison, W.C.; Drews, C.M.; Costa-Pinheiro, P.; Shaw, J.J.P.; Jobe, K.A.; Han, J.D.; Fox, T.E.; White, J.M.; et al. Glucosylceramide synthase maintains influenza virus entry and infection. PLoS ONE 2020, 15, e0228735. [Google Scholar] [CrossRef] [PubMed]
- Chotiwan, N.; Andre, B.G.; Sanchez-Vargas, I.; Islam, M.N.; Grabowski, J.M.; Hopf-Jannasch, A.; Gough, E.; Nakayasu, E.; Blair, C.D.; Belisle, J.T.; et al. Dynamic remodeling of lipids coincides with dengue virus replication in the midgut of Aedes aegypti mosquitoes. PLoS Pathog. 2018, 14, e1006853. [Google Scholar] [CrossRef]
- Moll, T.; Marshall, J.N.G.; Soni, N.; Zhang, S.; Cooper-Knock, J.; Shaw, P.J. Membrane lipid raft homeostasis is directly linked to neurodegeneration. Essays Biochem. 2021, 65, 999–1011. [Google Scholar] [CrossRef]
- Rossi, L.; Santos, K.B.S.; Mota, B.I.S.; Pimenta, J.; Oliveira, B.; Machado, C.A.; Fernandes, H.B.; Barbosa, L.A.; Rodrigues, H.A.; Teixeira, G.H.M.; et al. Neuromuscular defects after infection with a beta coronavirus in mice. Neurochem. Int. 2023, 169, 105567. [Google Scholar] [CrossRef]
- Raymond, J.; Berry, J.D.; Larson, T.; Horton, D.K.; Mehta, P. Effects of COVID-19 on motor neuron disease mortality in the United States: A population-based cross-sectional study. Amyotroph. Lateral Scler. Front. Degener. 2025, 26, 149–156. [Google Scholar] [CrossRef]
- Gay, L.; Desquiret-Dumas, V.; Nagot, N.; Rapenne, C.; Van de Perre, P.; Reynier, P.; Molès, J.-P. Long-term persistence of mitochondrial dysfunctions after viral infections and antiviral therapies: A review of mechanisms involved. J. Med. Virol. 2024, 96, e29886. [Google Scholar] [CrossRef]
- Bramble, D.M.; Lieberman, D.E. Endurance running and the evolution of Homo. Nature 2004, 432, 345–352. [Google Scholar] [CrossRef]
- Noakes, T.; Spedding, M. Olympics: Run for your life. Nature 2012, 487, 295–296. [Google Scholar] [CrossRef]
- Lieberman, D.E. The Story of the Human Body: Evolution, Health and Disease. Fam. Med. 2016, 48, 822–823. [Google Scholar]
- Okerblom, J.; Fletes, W.; Patel, H.H.; Schenk, S.; Varki, A.; Breen, E.C. Human-like Cmah inactivation in mice increases running endurance and decreases muscle fatigability: Implications for human evolution. Proc. Biol. Sci. 2018, 285, 20181656. [Google Scholar] [CrossRef] [PubMed]
- Martin, P.T.; Camboni, M.; Xu, R.; Golden, B.; Chandrasekharan, K.; Wang, C.-M.; Varki, A.; Janssen, P.M.L. N-Glycolylneuraminic acid deficiency worsens cardiac and skeletal muscle pathophysiology in α-sarcoglycan-deficient mice. Glycobiology 2013, 23, 833–843. [Google Scholar] [CrossRef]
- Mattson, M.P. Evolutionary aspects of human exercise—Born to run purposefully. Ageing Res. Rev. 2012, 11, 347–352. [Google Scholar] [CrossRef] [PubMed]
- Mattson, M.P. Lifelong brain health is a lifelong challenge: From evolutionary principles to empirical evidence. Ageing Res. Rev. 2015, 20, 37–45. [Google Scholar] [CrossRef] [PubMed]
- Bozek, K.; Wei, Y.; Yan, Z.; Liu, X.; Xiong, J.; Sugimoto, M.; Tomita, M.; Pääbo, S.; Sherwood, C.C.; Hof, P.R.; et al. Organization and evolution of brain lipidome revealed by large-scale analysis of human, chimpanzee, macaque, and mouse tissues. Neuron 2015, 85, 695–702. [Google Scholar] [CrossRef]
- Keeney, J.G.; Astling, D.; Andries, V.; Vandepoele, K.; Anderson, N.; Davis, J.M.; Lopert, P.; Vandenbussche, J.; Gevaert, K.; Staes, A.; et al. Olduvai domain expression downregulates mitochondrial pathways: Implications for human brain evolution and neoteny. bioRxiv 2024. bioRxiv:2024.10.21.619278. [Google Scholar] [CrossRef]
- Xing, L.; Gkini, V.; Nieminen, A.I.; Zhou, H.-C.; Aquilino, M.; Naumann, R.; Reppe, K.; Tanaka, K.; Carmeliet, P.; Heikinheimo, O.; et al. Functional synergy of a human-specific and an ape-specific metabolic regulator in human neocortex development. Nat. Commun. 2024, 15, 3468. [Google Scholar] [CrossRef]
- Bouscary, A.; Quessada, C.; Mosbach, A.; Callizot, N.; Spedding, M.; Loeffler, J.-P.; Henriques, A. Ambroxol Hydrochloride Improves Motor Functions and Extends Survival in a Mouse Model of Familial Amyotrophic Lateral Sclerosis. Front. Pharmacol. 2019, 10, 883. [Google Scholar] [CrossRef]
- Cutler, R.G.; Pedersen, W.A.; Camandola, S.; Rothstein, J.D.; Mattson, M.P. Evidence that accumulation of ceramides and cholesterol esters mediates oxidative stress-induced death of motor neurons in amyotrophic lateral sclerosis. Ann. Neurol. 2002, 52, 448–457. [Google Scholar] [CrossRef]
- Henriques, A.; Croixmarie, V.; Priestman, D.A.; Rosenbohm, A.; Dirrig-Grosch, S.; D’Ambra, E.; Huebecker, M.; Hussain, G.; Boursier-Neyret, C.; Echaniz-Laguna, A.; et al. Amyotrophic lateral sclerosis and denervation alter sphingolipids and up-regulate glucosylceramide synthase. Hum. Mol. Genet. 2015, 24, 7390–7405. [Google Scholar] [CrossRef]
- Henriques, A.; Croixmarie, V.; Bouscary, A.; Mosbach, A.; Keime, C.; Boursier-Neyret, C.; Walter, B.; Spedding, M.; Loeffler, J.-P. Sphingolipid Metabolism Is Dysregulated at Transcriptomic and Metabolic Levels in the Spinal Cord of an Animal Model of Amyotrophic Lateral Sclerosis. Front. Mol. Neurosci. 2017, 10, 433. [Google Scholar] [CrossRef]
- Dodge, J.C.; Treleaven, C.M.; Pacheco, J.; Cooper, S.; Bao, C.; Abraham, M.; Cromwell, M.; Sardi, S.P.; Chuang, W.-L.; Sidman, R.L.; et al. Glycosphingolipids are modulators of disease pathogenesis in amyotrophic lateral sclerosis. Proc. Natl. Acad. Sci. USA 2015, 112, 8100–8105. [Google Scholar] [CrossRef] [PubMed]
- Dodge, J.C. Lipid Involvement in Neurodegenerative Diseases of the Motor System: Insights from Lysosomal Storage Diseases. Front. Mol. Neurosci. 2017, 10, 356. [Google Scholar] [CrossRef] [PubMed]
- Blasco, H.; Corcia, P.; Pradat, P.-F.; Bocca, C.; Gordon, P.H.; Veyrat-Durebex, C.; Mavel, S.; Nadal-Desbarats, L.; Moreau, C.; Devos, D.; et al. Metabolomics in cerebrospinal fluid of patients with amyotrophic lateral sclerosis: An untargeted approach via high-resolution mass spectrometry. J. Proteome Res. 2013, 12, 3746–3754. [Google Scholar] [CrossRef] [PubMed]
- Morrison, T.A.; Vigee, J.; Tovar, K.A.; Talley, T.A.; Mujal, A.M.; Kono, M.; Philips, R.; Nagashima, H.; Brooks, S.R.; Dada, H.; et al. Selective requirement of glycosphingolipid synthesis for natural killer and cytotoxic T cells. Cell 2025, 188, 3497–3512.e16. [Google Scholar] [CrossRef]
- Chen, S.; Wang, X.; Jounaidi, Y. UGCG and the glycosphingolipid rheostat: A metabolic checkpoint governing immune activation and tumor immune evasion. Signal Transduct. Target. Ther. 2025, 10, 298. [Google Scholar] [CrossRef]
- Longo, J.; DeCamp, L.M.; Oswald, B.M.; Teis, R.; Reyes-Oliveras, A.; Dahabieh, M.S.; Ellis, A.E.; Vincent, M.P.; Damico, H.; Gallik, K.L.; et al. Glucose-dependent glycosphingolipid biosynthesis fuels CD8+ T cell function and tumor control. Cell Metab. 2025, 37, 1890–1906.e11. [Google Scholar] [CrossRef]
- Henriques, A.; Huebecker, M.; Blasco, H.; Keime, C.; Andres, C.R.; Corcia, P.; Priestman, D.A.; Platt, F.M.; Spedding, M.; Loeffler, J.P. Inhibition of β-Glucocerebrosidase Activity Preserves Motor Unit Integrity in a Mouse Model of Amyotrophic Lateral Sclerosis. Sci. Rep. 2017, 7, 5235. [Google Scholar] [CrossRef]
- Akiyama, H.; Ide, M.; Nagatsuka, Y.; Sayano, T.; Nakanishi, E.; Uemura, N.; Yuyama, K.; Yamaguchi, Y.; Kamiguchi, H.; Takahashi, R.; et al. Glucocerebrosidases catalyze a transgalactosylation reaction that yields a newly-identified brain sterol metabolite, galactosylated cholesterol. J. Biol. Chem. 2020, 295, 5257–5277. [Google Scholar] [CrossRef]
- Boggs, J.M. Role of Galactosylceramide and Sulfatide in Oligodendrocytes and CNS Myelin: Formation of a Glycosynapse. In Glycobiology of the Nervous System; Yu, R.K., Schengrund, C.-L., Eds.; Springer: New York, NY, USA, 2014; pp. 263–291. ISBN 978-1-4939-1154-7. [Google Scholar]
- Woeste, M.A.; Stern, S.; Raju, D.N.; Grahn, E.; Dittmann, D.; Gutbrod, K.; Dörmann, P.; Hansen, J.N.; Schonauer, S.; Marx, C.E.; et al. Species-specific differences in nonlysosomal glucosylceramidase GBA2 function underlie locomotor dysfunction arising from loss-of-function mutations. J. Biol. Chem. 2019, 294, 3853–3871. [Google Scholar] [CrossRef]
- Martin, E.; Schüle, R.; Smets, K.; Rastetter, A.; Boukhris, A.; Loureiro, J.L.; Gonzalez, M.A.; Mundwiller, E.; Deconinck, T.; Wessner, M.; et al. Loss of function of glucocerebrosidase GBA2 is responsible for motor neuron defects in hereditary spastic paraplegia. Am. J. Hum. Genet. 2013, 92, 238–244. [Google Scholar] [CrossRef] [PubMed]
- Rapport, M.M.; Donnenfeld, H.; Brunner, W.; Hungund, B.; Bartfeld, H. Ganglioside patterns in amyotrophic lateral sclerosis brain regions. Ann. Neurol. 1985, 18, 60–67. [Google Scholar] [CrossRef]
- Alisson-Silva, F.; Liu, J.Z.; Diaz, S.L.; Deng, L.; Gareau, M.G.; Marchelletta, R.; Chen, X.; Nizet, V.; Varki, N.; Barrett, K.E.; et al. Human evolutionary loss of epithelial Neu5Gc expression and species-specific susceptibility to cholera. PLoS Pathog. 2018, 14, e1007133. [Google Scholar] [CrossRef] [PubMed]
- Koch, R. An Address on Cholera and its Bacillus. Br. Med. J. 1884, 2, 403–407. [Google Scholar] [CrossRef] [PubMed]
- Turnbull, W.B.; Precious, B.L.; Homans, S.W. Dissecting the Cholera Toxin−Ganglioside GM1 Interaction by Isothermal Titration Calorimetry. J. Am. Chem. Soc. 2004, 126, 1047–1054. [Google Scholar] [CrossRef]
- Holmgren, J.; Lönnroth, I.; Månsson, J.; Svennerholm, L. Interaction of cholera toxin and membrane GM1 ganglioside of small intestine. Proc. Natl. Acad. Sci. USA 1975, 72, 2520–2524. [Google Scholar] [CrossRef]
- Chiricozzi, E.; Mauri, L.; Ciampa, M.G.; Prinetti, A.; Sonnino, S. On the use of cholera toxin. Glycoconj. J. 2018, 35, 161–163. [Google Scholar] [CrossRef]
- Cervin, J.; Wands, A.M.; Casselbrant, A.; Wu, H.; Krishnamurthy, S.; Cvjetkovic, A.; Estelius, J.; Dedic, B.; Sethi, A.; Wallom, K.-L.; et al. GM1 ganglioside-independent intoxication by Cholera toxin. PLoS Pathog. 2018, 14, e1006862. [Google Scholar] [CrossRef] [PubMed]
- Youn, G.; Cervin, J.; Yu, X.; Bhatia, S.R.; Yrlid, U.; Sampson, N.S. Targeting Multiple Binding Sites on Cholera Toxin B with Glycomimetic Polymers Promotes Formation of Protein–Polymer Aggregates. Biomacromolecules 2020, 21, 4878–4887. [Google Scholar] [CrossRef] [PubMed]
- Singla, A.; Boucher, A.; Wallom, K.-L.; Lebens, M.; Kohler, J.J.; Platt, F.M.; Yrlid, U. Cholera intoxication of human enteroids reveals interplay between decoy and functional glycoconjugate ligands. Glycobiology 2023, 33, 801–816. [Google Scholar] [CrossRef]
- Dave, F.; Vaghela, P.; Heath, B.; Dunster, Z.; Dubinina, E.; Thakker, D.; Mann, K.; Chadwick, J.; Cane, G.; Kaira, B.G.; et al. SC134-TCB Targeting Fucosyl-GM1, a T Cell-Engaging Antibody with Potent Antitumor Activity in Preclinical Small Cell Lung Cancer Models. Mol. Cancer Ther. 2024, 23, 1626–1638. [Google Scholar] [CrossRef]
- Berois, N.; Pittini, A.; Osinaga, E. Targeting Tumor Glycans for Cancer Therapy: Successes, Limitations, and Perspectives. Cancers 2022, 14, 645. [Google Scholar] [CrossRef]
- Llewellyn-Smith, I.J.; Martin, C.L.; Arnolda, L.F.; Minson, J.B. Tracer-toxins: Cholera toxin B-saporin as a model. J. Neurosci. Methods 2000, 103, 83–90. [Google Scholar] [CrossRef]
- Ciuro, M.; Sangiorgio, M.; Cacciato, V.; Cantone, G.; Fichera, C.; Salvatorelli, L.; Magro, G.; Leanza, G.; Vecchio, M.; Valle, M.S.; et al. Mitigating the Functional Deficit after Neurotoxic Motoneuronal Loss by an Inhibitor of Mitochondrial Fission. Int. J. Mol. Sci. 2024, 25, 7059. [Google Scholar] [CrossRef] [PubMed]
- Gulino, R.; Vicario, N.; Giunta, M.A.S.; Spoto, G.; Calabrese, G.; Vecchio, M.; Gulisano, M.; Leanza, G.; Parenti, R. Neuromuscular Plasticity in a Mouse Neurotoxic Model of Spinal Motoneuronal Loss. Int. J. Mol. Sci. 2019, 20, 1500. [Google Scholar] [CrossRef]
- Lind, L.A.; Murphy, E.R.; Lever, T.E.; Nichols, N.L. Hypoglossal Motor Neuron Death Via Intralingual CTB-saporin (CTB-SAP) Injections Mimic Aspects of Amyotrophic Lateral Sclerosis (ALS) Related to Dysphagia. Neuroscience 2018, 390, 303–316. [Google Scholar] [CrossRef] [PubMed]
- Lewis, R.D.; Keilholz, A.N.; Smith, C.L.; Burd, E.A.; Nichols, N.L. Spinal TNF-α receptor 1 is differentially required for phrenic long-term facilitation (pLTF) over the course of motor neuron death in adult rats. Front. Physiol. 2024, 15, 1488951. [Google Scholar] [CrossRef]
- Chew, C.; Sengelaub, D.R. Exercise is neuroprotective on the morphology of somatic motoneurons following the death of neighboring motoneurons via androgen action at the target muscle. Dev. Neurobiol. 2021, 81, 22–35. [Google Scholar] [CrossRef]
- Lind, L.A.; Lever, T.E.; Nichols, N.L. Tongue and hypoglossal morphology after intralingual cholera toxin B-saporin injection. Muscle Nerve 2021, 63, 413–420. [Google Scholar] [CrossRef]
- Keilholz, A.N.; Pathak, I.; Smith, C.L.; Osman, K.L.; Smith, L.; Oti, G.; Golzy, M.; Ma, L.; Lever, T.E.; Nichols, N.L. Tongue exercise ameliorates structural and functional upper airway deficits in a rodent model of hypoglossal motor neuron loss. Front. Neurol. 2024, 15, 1441529. [Google Scholar] [CrossRef]
- Pestronk, A.; Choksi, R. Multifocal motor neuropathy. Serum IgM anti-GM1 ganglioside antibodies in most patients detected using covalent linkage of GM1 to ELISA plates. Neurology 1997, 49, 1289–1292. [Google Scholar] [CrossRef] [PubMed]
- Stikvoort García, D.J.L.; Kovalchuk, M.O.; Goedee, H.S.; van Schelven, L.J.; van den Berg, L.H.; Franssen, H.; Sleutjes, B.T.H.M. Motor unit integrity in multifocal motor neuropathy: A systematic evaluation with CMAP scans. Muscle Nerve 2022, 65, 317–325. [Google Scholar] [CrossRef] [PubMed]
- Zhu, W.; Li, K.; Cui, T.; Yan, Y. Detection of anti-ganglioside antibodies in Guillain-Barré syndrome. Ann. Transl. Med. 2023, 11, 289. [Google Scholar] [CrossRef]
- Okubo, S.; Maeda, M.; Katsuse, K.; Ishiura, H.; Shirota, Y.; Hamada, M.; Satake, W.; Toda, T. Subacute Upper Motor Neuron Dysfunction Possibly Associated with the Anti-GM1 Autoantibody: A Case Report. Intern. Med. 2025, 64, 1900–1905. [Google Scholar] [CrossRef]
- Niebroj-Dobosz, I.; Janik, P.; Kwieciński, H. Serum IgM anti-GM1 ganglioside antibodies in lower motor neuron syndromes. Eur. J. Neurol. 2004, 11, 13–16. [Google Scholar] [CrossRef]
- Pestronk, A.; Adams, R.N.; Clawson, L.; Cornblath, D.; Kuncl, R.W.; Griffin, D.; Drachman, D.B. Serum antibodies to GM1 ganglioside in amyotrophic lateral sclerosis. Neurology 1988, 38, 1457–1461. [Google Scholar] [CrossRef]
- Salazar-Grueso, E.F.; Routbort, M.J.; Martin, J.; Dawson, G.; Roos, R.P. Polyclonal IgM anti-GM1 ganglioside antibody in patients with motor neuron disease and variants. Ann. Neurol. 1990, 27, 558–563. [Google Scholar] [CrossRef] [PubMed]
- Voumvourakis, C.; Rombos, A.; Konstantoulakis, M.M.; Segditsa, I.; Papageorgiou, C. Serum anti-GM1 and anti-GD1a antibodies in patients with motor neuron disease. Acta Neurol. Scand. 1992, 86, 599–602. [Google Scholar] [CrossRef]
- Nobile-Orazio, E.; Carpo, M.; Scarlato, G. Clinical relevance of anti-GM1 IgM antibodies. Acta Neurol. 1991, 13, 514–519. [Google Scholar]
- Li, D.; Usuki, S.; Quarles, B.; Rivner, M.H.; Ariga, T.; Yu, R.K. Anti-Sulfoglucuronosyl Paragloboside Antibody: A Potential Serologic Marker of Amyotrophic Lateral Sclerosis. ASN Neuro 2016, 8, 1759091416669619. [Google Scholar] [CrossRef]
- Niebroj-Dobosz, I.; Jamrozik, Z.; Janik, P.; Hausmanowa-Petrusewicz, I.; Kwieciński, H. Anti-neural antibodies in serum and cerebrospinal fluid of amyotrophic lateral sclerosis (ALS) patients. Acta Neurol. Scand. 1999, 100, 238–243. [Google Scholar] [CrossRef]
- Kollewe, K.; Wurster, U.; Sinzenich, T.; Mohammadi, B.; Körner, S.; Dengler, R.; Petri, S. Ganglioside Antibodies in Amyotrophic Lateral Sclerosis. Klin. Neurophysiol. 2013, 44, P93. [Google Scholar] [CrossRef]
- Svennerholm, L.; Boström, K.; Jungbjer, B.; Olsson, L. Membrane lipids of adult human brain: Lipid composition of frontal and temporal lobe in subjects of age 20 to 100 years. J. Neurochem. 1994, 63, 1802–1811. [Google Scholar] [CrossRef]
- Chowdhury, S.; Wu, G.; Lu, Z.-H.; Kumar, R.; Ledeen, R. Age-Related Decline in Gangliosides GM1 and GD1a in Non-CNS Tissues of Normal Mice: Implications for Peripheral Symptoms of Parkinson’s Disease. Biomedicines 2023, 11, 209. [Google Scholar] [CrossRef]
- Chiricozzi, E.; Pomè, D.Y.; Maggioni, M.; Di Biase, E.; Parravicini, C.; Palazzolo, L.; Loberto, N.; Eberini, I.; Sonnino, S. Role of the GM1 ganglioside oligosaccharide portion in the TrkA-dependent neurite sprouting in neuroblastoma cells. J. Neurochem. 2017, 143, 645–659. [Google Scholar] [CrossRef] [PubMed]
- Chowdhury, S.; Ledeen, R. The Key Role of GM1 Ganglioside in Parkinson’s Disease. Biomolecules 2022, 12, 173. [Google Scholar] [CrossRef]
- Wu, G.; Lu, Z.-H.; Kulkarni, N.; Amin, R.; Ledeen, R.W. Mice lacking major brain gangliosides develop parkinsonism. Neurochem. Res. 2011, 36, 1706–1714. [Google Scholar] [CrossRef] [PubMed]
- Ledeen, R.W.; Wu, G. The multi-tasked life of GM1 ganglioside, a true factotum of nature. Trends Biochem. Sci. 2015, 40, 407–418. [Google Scholar] [CrossRef] [PubMed]
- Martinez, Z.; Zhu, M.; Han, S.; Fink, A.L. GM1 specifically interacts with alpha-synuclein and inhibits fibrillation. Biochemistry 2007, 46, 1868–1877. [Google Scholar] [CrossRef] [PubMed]
- La Vitola, P.; Szegö, E.M.; Pinto-Costa, R.; Rollar, A.; Harbachova, E.; Schapira, A.H.; Ulusoy, A.; Di Monte, D.A. Mitochondrial oxidant stress promotes α-synuclein aggregation and spreading in mice with mutated glucocerebrosidase. npj Park. Dis. 2024, 10, 233. [Google Scholar] [CrossRef]
- Blandini, F.; Cilia, R.; Cerri, S.; Pezzoli, G.; Schapira, A.H.V.; Mullin, S.; Lanciego, J.L. Glucocerebrosidase mutations and synucleinopathies: Toward a model of precision medicine. Mov. Disord. 2019, 34, 9–21. [Google Scholar] [CrossRef]
- Gegg, M.E.; Menozzi, E.; Schapira, A.H.V. Glucocerebrosidase-associated Parkinson disease: Pathogenic mechanisms and potential drug treatments. Neurobiol. Dis. 2022, 166, 105663. [Google Scholar] [CrossRef]
- Wang, R.; Tong, S.; Wang, M.; Zou, J.; Wang, N.; Sun, F.; Zhou, X.; Chen, J.; Wang, H. CREB5 hypermethylation involved in the ganglioside GM1 therapy of Parkinson’s disease. Front. Aging Neurosci. 2023, 15, 1122647. [Google Scholar] [CrossRef]
- Fazzari, M.; Lunghi, G.; Chiricozzi, E.; Mauri, L.; Sonnino, S. Gangliosides and the Treatment of Neurodegenerative Diseases: A Long Italian Tradition. Biomedicines 2022, 10, 363. [Google Scholar] [CrossRef]
- DeVries, G.H.; Campbell, B.; Saunders, R. Isolation and characterization of unmyelinated axolemma from bovine splenic nerve. J. Neurosci. Res. 1999, 57, 670–679. [Google Scholar] [CrossRef]
- Iacoangeli, A.; Lin, T.; Al Khleifat, A.; Jones, A.R.; Opie-Martin, S.; Coleman, J.R.I.; Shatunov, A.; Sproviero, W.; Williams, K.L.; Garton, F.; et al. Genome-wide Meta-analysis Finds the ACSL5-ZDHHC6 Locus Is Associated with ALS and Links Weight Loss to the Disease Genetics. Cell Rep. 2020, 33, 108323. [Google Scholar] [CrossRef]
- Lunghi, G.; Di Biase, E.; Carsana, E.V.; Henriques, A.; Callizot, N.; Mauri, L.; Ciampa, M.G.; Mari, L.; Loberto, N.; Aureli, M.; et al. GM1 ganglioside exerts protective effects against glutamate-excitotoxicity via its oligosaccharide in wild-type and amyotrophic lateral sclerosis motor neurons. FEBS Open Bio 2023, 13, 2324–2341. [Google Scholar] [CrossRef]
- Chiricozzi, E.; Di Biase, E.; Lunghi, G.; Fazzari, M.; Loberto, N.; Aureli, M.; Mauri, L.; Sonnino, S. Turning the spotlight on the oligosaccharide chain of GM1 ganglioside. Glycoconj. J. 2021, 38, 101–117. [Google Scholar] [CrossRef]
- Mutoh, T.; Hamano, T.; Yano, S.; Koga, H.; Yamamoto, H.; Furukawa, K.; Ledeen, R.W. Stable transfection of GM1 synthase gene into GM1-deficient NG108-15 cells, CR-72 cells, rescues the responsiveness of Trk-neurotrophin receptor to its ligand, NGF. Neurochem. Res. 2002, 27, 801–806. [Google Scholar] [CrossRef] [PubMed]
- Mutoh, T.; Tokuda, A.; Miyadai, T.; Hamaguchi, M.; Fujiki, N. Ganglioside GM1 binds to the Trk protein and regulates receptor function. Proc. Natl. Acad. Sci. USA 1995, 92, 5087–5091. [Google Scholar] [CrossRef]
- Chiricozzi, E.; Lunghi, G.; Di Biase, E.; Fazzari, M.; Sonnino, S.; Mauri, L. GM1 Ganglioside Is A Key Factor in Maintaining the Mammalian Neuronal Functions Avoiding Neurodegeneration. Int. J. Mol. Sci. 2020, 21, 868. [Google Scholar] [CrossRef]
- Wallom, K.-L.; Fernández-Suárez, M.E.; Priestman, D.A.; Te Vruchte, D.; Huebecker, M.; Hallett, P.J.; Isacson, O.; Platt, F.M. Glycosphingolipid metabolism and its role in ageing and Parkinson’s disease. Glycoconj. J. 2022, 39, 39–53. [Google Scholar] [CrossRef]
- Fernández-Beltrán, L.C.; Godoy-Corchuelo, J.M.; Losa-Fontangordo, M.; Williams, D.; Matias-Guiu, J.; Corrochano, S. A Transcriptomic Meta-Analysis Shows Lipid Metabolism Dysregulation as an Early Pathological Mechanism in the Spinal Cord of SOD1 Mice. Int. J. Mol. Sci. 2021, 22, 9553. [Google Scholar] [CrossRef]
- Fantini, J. Lipid rafts and human diseases: Why we need to target gangliosides. FEBS Open Bio 2023, 13, 1636. [Google Scholar] [CrossRef] [PubMed]
- Blasco, H.; Veyrat-Durebex, C.; Bocca, C.; Patin, F.; Vourc’h, P.; Kouassi Nzoughet, J.; Lenaers, G.; Andres, C.R.; Simard, G.; Corcia, P.; et al. Lipidomics Reveals Cerebrospinal-Fluid Signatures of ALS. Sci Rep. 2017, 7, 17652. [Google Scholar] [CrossRef] [PubMed]
- Thomma, R.C.; Fokke, C.; Walgaard, C.; Vermeulen-de Jongh, D.M.; Tio-Gillen, A.; van Rijs, W.; van Doorn, P.A.; Huizinga, R.; Jacobs, B.C. High and Persistent Anti-GM1 Antibody Titers Are Associated with Poor Clinical Recovery in Guillain-Barré Syndrome. Neurol. Neuroimmunol. Neuroinflamm. 2023, 10, e200107. [Google Scholar] [CrossRef]
- Bachis, A.; Rabin, S.J.; Del Fiacco, M.; Mocchetti, I. Gangliosides prevent excitotoxicity through activation of TrkB receptor. Neurotox. Res. 2002, 4, 225–234. [Google Scholar] [CrossRef]
- Ledeen, R.; Wu, G. GM1 in the nuclear envelope regulates nuclear calcium through association with a nuclear sodium-calcium exchanger. J. Neurochem. 2007, 103 (Suppl. S1), 126–134. [Google Scholar] [CrossRef]
- Ledeen, R.W.; Wu, G. Nuclear sphingolipids: Metabolism and signaling. J. Lipid Res. 2008, 49, 1176–1186. [Google Scholar] [CrossRef] [PubMed]
- Ledeen, R.; Wu, G. New findings on nuclear gangliosides: Overview on metabolism and function. J. Neurochem. 2011, 116, 714–720. [Google Scholar] [CrossRef] [PubMed]
- Finsterwald, C.; Dias, S.; Magistretti, P.J.; Lengacher, S. Ganglioside GM1 Targets Astrocytes to Stimulate Cerebral Energy Metabolism. Front. Pharmacol. 2021, 12, 653842. [Google Scholar] [CrossRef] [PubMed]
- Fazzari, M.; Audano, M.; Lunghi, G.; Di Biase, E.; Loberto, N.; Mauri, L.; Mitro, N.; Sonnino, S.; Chiricozzi, E. The oligosaccharide portion of ganglioside GM1 regulates mitochondrial function in neuroblastoma cells. Glycoconj. J. 2020, 37, 293–306. [Google Scholar] [CrossRef]
- Luikinga, S.; Henriques, A.; Ngo, S.T.; Rapasinghe, T.; Loeffler, J.-P.; Spedding, M.; Turner, B.J. Profound lipid dysregulation in mutant TDP-43 mice is ameliorated by the glucocerebrosidase 2 inhibitor ambroxol. bioRxiv 2022. bioRxiv:2022.08.30.505901. [Google Scholar] [CrossRef]
- Pitto, M.; Mutoh, T.; Kuriyama, M.; Ferraretto, A.; Palestini, P.; Masserini, M. Influence of endogenous GM1 ganglioside on TrkB activity, in cultured neurons. FEBS Lett. 1998, 439, 93–96. [Google Scholar] [CrossRef]
- Jiang, B.; Song, L.; Wang, C.-N.; Zhang, W.; Huang, C.; Tong, L.-J. Antidepressant-Like Effects of GM1 Ganglioside Involving the BDNF Signaling Cascade in Mice. Int. J. Neuropsychopharmacol. 2016, 19, pyw046. [Google Scholar] [CrossRef]
- Markham, A.; Cameron, I.; Bains, R.; Franklin, P.; Kiss, J.P.; Schwendimann, L.; Gressens, P.; Spedding, M. Brain-derived neurotrophic factor-mediated effects on mitochondrial respiratory coupling and neuroprotection share the same molecular signalling pathways. Eur. J. Neurosci. 2012, 35, 366–374. [Google Scholar] [CrossRef]
- Markham, A.; Bains, R.; Franklin, P.; Spedding, M. Changes in mitochondrial function are pivotal in neurodegenerative and psychiatric disorders: How important is BDNF? Br. J. Pharmacol. 2014, 171, 2206–2229. [Google Scholar] [CrossRef]
- Tan, W.; Naniche, N.; Bogush, A.; Pedrini, S.; Trotti, D.; Pasinelli, P. Small peptides against the mutant SOD1/Bcl-2 toxic mitochondrial complex restore mitochondrial function and cell viability in mutant SOD1-mediated ALS. J. Neurosci. 2013, 33, 11588–11598. [Google Scholar] [CrossRef]
- Stansberry, W.M.; Pierchala, B.A. Neurotrophic factors in the physiology of motor neurons and their role in the pathobiology and therapeutic approach to amyotrophic lateral sclerosis. Front. Mol. Neurosci. 2023, 16, 1238453. [Google Scholar] [CrossRef] [PubMed]
- Casimir, P.; Iwata, R.; Vanderhaeghen, P. Linking mitochondria metabolism, developmental timing, and human brain evolution. Curr. Opin. Genet. Dev. 2024, 86, 102182. [Google Scholar] [CrossRef]
- Assendorp, N.; Fossati, M.; Libé-Philippot, B.; Christopoulou, E.; Depp, M.; Rapone, R.; Dingli, F.; Loew, D.; Vanderhaeghen, P.; Charrier, C. CTNND2 moderates the pace of synaptic maturation and links human evolution to synaptic neoteny. Cell Rep. 2024, 43, 114797. [Google Scholar] [CrossRef] [PubMed]
- Iwata, R.; Vanderhaeghen, P. Metabolic mechanisms of species-specific developmental tempo. Dev. Cell 2024, 59, 1628–1639. [Google Scholar] [CrossRef]
- Zemke, N.R.; Armand, E.J.; Wang, W.; Lee, S.; Zhou, J.; Li, Y.E.; Liu, H.; Tian, W.; Nery, J.R.; Castanon, R.G.; et al. Conserved and divergent gene regulatory programs of the mammalian neocortex. Nature 2023, 624, 390–402. [Google Scholar] [CrossRef]
- Mosharov, E.V.; Rosenberg, A.M.; Monzel, A.S.; Osto, C.A.; Stiles, L.; Rosoklija, G.B.; Dwork, A.J.; Bindra, S.; Junker, A.; Zhang, Y.; et al. A human brain map of mitochondrial respiratory capacity and diversity. Nature 2025, 641, 749–758. [Google Scholar] [CrossRef] [PubMed]
- Vaill, M.; Kawanishi, K.; Varki, N.; Gagneux, P.; Varki, A. Comparative physiological anthropogeny: Exploring molecular underpinnings of distinctly human phenotypes. Physiol. Rev. 2023, 103, 2171–2229. [Google Scholar] [CrossRef]
- Blasco, H.; Bessy, C.; Plantier, L.; Lefevre, A.; Piver, E.; Bernard, L.; Marlet, J.; Stefic, K.; Benz-de Bretagne, I.; Cannet, P.; et al. The specific metabolome profiling of patients infected by SARS-CoV-2 supports the key role of tryptophan-nicotinamide pathway and cytosine metabolism. Sci. Rep. 2020, 10, 16824. [Google Scholar] [CrossRef]
- Area-Gomez, E.; Larrea, D.; Yun, T.; Xu, Y.; Hupf, J.; Zandkarimi, F.; Chan, R.B.; Mitsumoto, H. Lipidomics study of plasma from patients suggest that ALS and PLS are part of a continuum of motor neuron disorders. Sci. Rep. 2021, 11, 13562. [Google Scholar] [CrossRef]
- Holland, W.L.; Brozinick, J.T.; Wang, L.-P.; Hawkins, E.D.; Sargent, K.M.; Liu, Y.; Narra, K.; Hoehn, K.L.; Knotts, T.A.; Siesky, A.; et al. Inhibition of Ceramide Synthesis Ameliorates Glucocorticoid-, Saturated-Fat-, and Obesity-Induced Insulin Resistance. Cell Metab. 2007, 5, 167–179. [Google Scholar] [CrossRef]
- Summers, S.A.; Garza, L.A.; Zhou, H.; Birnbaum, M.J. Regulation of insulin-stimulated glucose transporter GLUT4 translocation and Akt kinase activity by ceramide. Mol. Cell Biol. 1998, 18, 5457–5464. [Google Scholar] [CrossRef] [PubMed]
- Zhou, H.; Summers, S.A.; Birnbaum, M.J.; Pittman, R.N. Inhibition of Akt kinase by cell-permeable ceramide and its implications for ceramide-induced apoptosis. J. Biol. Chem. 1998, 273, 16568–16575. [Google Scholar] [CrossRef]
- Guenther, G.G.; Edinger, A.L. A new take on ceramide: Starving cells by cutting off the nutrient supply. Cell Cycle 2009, 8, 1122–1126. [Google Scholar] [CrossRef]
- Guenther, G.G.; Peralta, E.R.; Rosales, K.R.; Wong, S.Y.; Siskind, L.J.; Edinger, A.L. Ceramide starves cells to death by downregulating nutrient transporter proteins. Proc. Natl. Acad. Sci. USA 2008, 105, 17402–17407. [Google Scholar] [CrossRef] [PubMed]
- Laurila, P.-P.; Luan, P.; Wohlwend, M.; Zanou, N.; Crisol, B.; Imamura de Lima, T.; Goeminne, L.J.E.; Gallart-Ayala, H.; Shong, M.; Ivanisevic, J.; et al. Inhibition of sphingolipid de novo synthesis counteracts muscular dystrophy. Sci. Adv. 2022, 8, eabh4423. [Google Scholar] [CrossRef]
- Trayssac, M.; Hannun, Y.A.; Obeid, L.M. Role of sphingolipids in senescence: Implication in aging and age-related diseases. J. Clin. Investig. 2018, 128, 2702–2712. [Google Scholar] [CrossRef] [PubMed]
- Sun, S.; Li, J.; Wang, S.; Li, J.; Ren, J.; Bao, Z.; Sun, L.; Ma, X.; Zheng, F.; Ma, S.; et al. CHIT1-positive microglia drive motor neuron ageing in the primate spinal cord. Nature 2023, 624, 611–620. [Google Scholar] [CrossRef]
- Poisson, J.; Daskalaki, I.; Potluri, V.; Morel, J.-D.; Rodriguez-Lopez, S.; De Masi, A.; Benegiamo, G.; Jain, S.; Lima, T.; Auwerx, J. Safe and Orally Bioavailable Inhibitor of Serine Palmitoyltransferase Improves Age-Related Sarcopenia. ACS Pharmacol. Transl. Sci. 2025, 8, 203–215. [Google Scholar] [CrossRef]
- Mohassel, P.; Donkervoort, S.; Lone, M.A.; Nalls, M.; Gable, K.; Gupta, S.D.; Foley, A.R.; Hu, Y.; Saute, J.A.M.; Moreira, A.L.; et al. Childhood amyotrophic lateral sclerosis caused by excess sphingolipid synthesis. Nat. Med. 2021, 27, 1197–1204. [Google Scholar] [CrossRef] [PubMed]
- Syeda, S.B.; Lone, M.A.; Mohassel, P.; Donkervoort, S.; Munot, P.; França, M.C.; Galarza-Brito, J.E.; Eckenweiler, M.; Asamoah, A.; Gable, K.; et al. Recurrent de novo SPTLC2 variant causes childhood-onset amyotrophic lateral sclerosis (ALS) by excess sphingolipid synthesis. J. Neurol. Neurosurg. Psychiatry 2024, 95, 103–113. [Google Scholar] [CrossRef]
- Mahawar, U.; Davis, D.L.; Kannan, M.; Suemitsu, J.; Oltorik, C.D.; Farooq, F.; Fulani, R.; Weintraub, C.; Allegood, J.; Wattenberg, B. The individual isoforms of ORMDL, the regulatory subunit of serine palmitoyltransferase, have distinctive sensitivities to ceramide. bioRxiv 2025. bioRxiv:2025.03.20.643044. [Google Scholar] [CrossRef]
- Mohassel, P.; Abdullah, M.; Eichler, F.S.; Dunn, T.M. Serine Palmitoyltransferase (SPT)-related Neurodegenerative and Neurodevelopmental Disorders. J. Neuromuscul. Dis. 2024, 11, 735–747. [Google Scholar] [CrossRef]
- Lunghi, G.; Fazzari, M.; Di Biase, E.; Mauri, L.; Chiricozzi, E.; Sonnino, S. The structure of gangliosides hides a code for determining neuronal functions. FEBS Open Bio 2021, 11, 3193–3200. [Google Scholar] [CrossRef]
- Wohlwend, M.; Laurila, P.-P.; Goeminne, L.J.; Lima, T.; Daskalaki, I.; Li, X.; von Alvensleben, G.; Crisol, B.; Mangione, R.; Gallart-Ayala, H.; et al. Inhibition of CERS1 in skeletal muscle exacerbates age-related muscle dysfunction. eLife 2024, 12, RP90522. [Google Scholar] [CrossRef] [PubMed]
- Gutner, U.A.; Shupik, M.A.; Maloshitskaya, O.A.; Sokolov, S.A.; Rezvykh, A.P.; Funikov, S.Y.; Lebedev, A.T.; Ustyugov, A.A.; Alessenko, A.V. Changes in the Metabolism of Sphingoid Bases in the Brain and Spinal Cord of Transgenic FUS(1-359) Mice, a Model of Amyotrophic Lateral Sclerosis. Biochemistry 2019, 84, 1166–1176. [Google Scholar] [CrossRef] [PubMed]
- McInnis, J.J.; Sood, D.; Guo, L.; Dufault, M.R.; Garcia, M.; Passaro, R.; Gao, G.; Zhang, B.; Dodge, J.C. Unravelling neuronal and glial differences in ceramide composition, synthesis, and sensitivity to toxicity. Commun. Biol. 2024, 7, 1597. [Google Scholar] [CrossRef]
- Choi, S.W.; Gu, Y.; Peters, R.S.; Salgame, P.; Ellner, J.J.; Timmins, G.S.; Deretic, V. Ambroxol Induces Autophagy and Potentiates Rifampin Antimycobacterial Activity. Antimicrob. Agents Chemother. 2018, 62, e01019-18. [Google Scholar] [CrossRef]
- Higashi, K.; Sonoda, Y.; Kaku, N.; Fujii, F.; Yamashita, F.; Lee, S.; Tocan, V.; Ebihara, G.; Matsuoka, W.; Tetsuhara, K.; et al. Rapid and long-lasting efficacy of high-dose ambroxol therapy for neuronopathic Gaucher disease: A case report and literature review. Mol. Genet. Genom. Med. 2024, 12, e2427. [Google Scholar] [CrossRef]
- Skrahin, A.; Horowitz, M.; Istaiti, M.; Skrahina, V.; Lukas, J.; Yahalom, G.; Cohen, M.E.; Revel-Vilk, S.; Goker-Alpan, O.; Becker-Cohen, M.; et al. GBA1-Associated Parkinson’s Disease Is a Distinct Entity. Int. J. Mol. Sci. 2024, 25, 7102. [Google Scholar] [CrossRef]
- Gustavsson, E.K.; Sethi, S.; Gao, Y.; Brenton, J.W.; García-Ruiz, S.; Zhang, D.; Garza, R.; Reynolds, R.H.; Evans, J.R.; Chen, Z.; et al. The annotation of GBA1 has been concealed by its protein-coding pseudogene GBAP1. Sci. Adv. 2024, 10, eadk1296. [Google Scholar] [CrossRef] [PubMed]
- Eisen, A.; Vucic, S.; Kiernan, M.C. Amyotrophic lateral sclerosis represents corticomotoneuronal system failure. Muscle Nerve 2024, 71, 499–511. [Google Scholar] [CrossRef]
- Bozek, K.; Wei, Y.; Yan, Z.; Liu, X.; Xiong, J.; Sugimoto, M.; Tomita, M.; Pääbo, S.; Pieszek, R.; Sherwood, C.C.; et al. Exceptional evolutionary divergence of human muscle and brain metabolomes parallels human cognitive and physical uniqueness. PLoS Biol. 2014, 12, e1001871. [Google Scholar] [CrossRef] [PubMed]
- Li, Q.; Bozek, K.; Xu, C.; Guo, Y.; Sun, J.; Pääbo, S.; Sherwood, C.C.; Hof, P.R.; Ely, J.J.; Li, Y.; et al. Changes in Lipidome Composition during Brain Development in Humans, Chimpanzees, and Macaque Monkeys. Mol. Biol. Evol. 2017, 34, 1155–1166. [Google Scholar] [CrossRef]
- O’Neill, M.C.; Umberger, B.R.; Holowka, N.B.; Larson, S.G.; Reiser, P.J. Chimpanzee super strength and human skeletal muscle evolution. Proc. Natl. Acad. Sci. USA 2017, 114, 7343–7348. [Google Scholar] [CrossRef]
- Shefner, J.M.; Musaro, A.; Ngo, S.T.; Lunetta, C.; Steyn, F.J.; Robitaille, R.; De Carvalho, M.; Rutkove, S.; Ludolph, A.C.; Dupuis, L. Skeletal muscle in amyotrophic lateral sclerosis. Brain 2023, 46, 4425–4436. [Google Scholar] [CrossRef]
- San-Millán, I.; Brooks, G.A. Assessment of Metabolic Flexibility by Means of Measuring Blood Lactate, Fat, and Carbohydrate Oxidation Responses to Exercise in Professional Endurance Athletes and Less-Fit Individuals. Sports Med. 2018, 48, 467–479. [Google Scholar] [CrossRef] [PubMed]
- Nemkov, T.; Cendali, F.; Stefanoni, D.; Martinez, J.L.; Hansen, K.C.; San-Millán, I.; D’Alessandro, A. Metabolic Signatures of Performance in Elite World Tour Professional Male Cyclists. Sports Med. 2023, 53, 1651–1665. [Google Scholar] [CrossRef]
- Edman, S.; Horwath, O.; Van der Stede, T.; Blackwood, S.J.; Moberg, I.; Strömlind, H.; Nordström, F.; Ekblom, M.; Katz, A.; Apró, W.; et al. Pro-Brain-Derived Neurotrophic Factor (BDNF), but Not Mature BDNF, Is Expressed in Human Skeletal Muscle: Implications for Exercise-Induced Neuroplasticity. Function 2024, 5, zqae005. [Google Scholar] [CrossRef]
- Lei, Z.; Mozaffaritabar, S.; Kawamura, T.; Koike, A.; Kolonics, A.; Kéringer, J.; Pinho, R.A.; Sun, J.; Shangguan, R.; Radák, Z. The effects of long-term lactate and high-intensity interval training (HIIT) on brain neuroplasticity of aged mice. Heliyon 2024, 10, e24421. [Google Scholar] [CrossRef]
- Zintel, T.M.; Pizzollo, J.; Claypool, C.G.; Babbitt, C.C. Astrocytes Drive Divergent Metabolic Gene Expression in Humans and Chimpanzees. Genome Biol. Evol. 2024, 16, evad239. [Google Scholar] [CrossRef]
- Lee, I.; Kazamel, M.; McPherson, T.; McAdam, J.; Bamman, M.; Amara, A.; Smith, D.L.; King, P.H. Fat mass loss correlates with faster disease progression in amyotrophic lateral sclerosis patients: Exploring the utility of dual-energy x-ray absorptiometry in a prospective study. PLoS ONE 2021, 16, e0251087. [Google Scholar] [CrossRef]
- Pontzer, H.; Brown, M.H.; Raichlen, D.A.; Dunsworth, H.; Hare, B.; Walker, K.; Luke, A.; Dugas, L.R.; Durazo-Arvizu, R.; Schoeller, D.; et al. Metabolic acceleration and the evolution of human brain size and life history. Nature 2016, 533, 390–392. [Google Scholar] [CrossRef]
- Herrmann, C.; Uzelac, Z.; Michels, S.; Weber, A.; Richter, L.; Elmas, Z.; Jagodzinski, L.; Wurster, C.; Schuster, J.; Dreyhaupt, J.; et al. Alterations of Fat and Ketone Body Metabolism in ALS and SMA-A Prospective Observational Study. Eur. J. Neurol. 2025, 32, e70132. [Google Scholar] [CrossRef]
- Palamiuc, L.; Schlagowski, A.; Ngo, S.T.; Vernay, A.; Dirrig-Grosch, S.; Henriques, A.; Boutillier, A.-L.; Zoll, J.; Echaniz-Laguna, A.; Loeffler, J.-P.; et al. A metabolic switch toward lipid use in glycolytic muscle is an early pathologic event in a mouse model of amyotrophic lateral sclerosis. EMBO Mol. Med. 2015, 7, 526–546. [Google Scholar] [CrossRef] [PubMed]
- Scaricamazza, S.; Salvatori, I.; Giacovazzo, G.; Loeffler, J.P.; Renè, F.; Rosina, M.; Quessada, C.; Proietti, D.; Heil, C.; Rossi, S.; et al. Skeletal-Muscle Metabolic Reprogramming in ALS-SOD1G93A Mice Predates Disease Onset and Is A Promising Therapeutic Target. iScience 2020, 23, 101087. [Google Scholar] [CrossRef] [PubMed]
- Bennett, E.J.; Mead, R.J.; Azzouz, M.; Shaw, P.J.; Grierson, A.J. Early detection of motor dysfunction in the SOD1G93A mouse model of Amyotrophic Lateral Sclerosis (ALS) using home cage running wheels. PLoS ONE 2014, 9, e107918. [Google Scholar] [CrossRef] [PubMed]
- San-Millán, I. The Key Role of Mitochondrial Function in Health and Disease. Antioxidants 2023, 12, 782. [Google Scholar] [CrossRef]
- San-Millan, I.; Sparagna, G.C.; Chapman, H.L.; Warkins, V.L.; Chatfield, K.C.; Shuff, S.R.; Martinez, J.L.; Brooks, G.A. Chronic Lactate Exposure Decreases Mitochondrial Function by Inhibition of Fatty Acid Uptake and Cardiolipin Alterations in Neonatal Rat Cardiomyocytes. Front. Nutr. 2022, 9, 809485. [Google Scholar] [CrossRef]
- Vadakkadath Meethal, S.; Atwood, C.S. Lactate dyscrasia: A novel explanation for amyotrophic lateral sclerosis. Neurobiol. Aging 2012, 33, 569–581. [Google Scholar] [CrossRef]
- Spedding, M.; Marvaud, R.; Marck, A.; Delarochelambert, Q.; Toussaint, J.F. Aging, VO2 max, entropy, and COVID-19. Indian J. Pharmacol. 2022, 54, 58–62. [Google Scholar] [CrossRef] [PubMed]
- Mattern, C.O.; Gutilla, M.J.; Bright, D.L.; Kirby, T.E.; Hinchcliff, K.W.; Devor, S.T. Maximal lactate steady state declines during the aging process. J. Appl. Physiol. (1985) 2003, 95, 2576–2582. [Google Scholar] [CrossRef] [PubMed]
- Lanfranconi, F.; Ferri, A.; Corna, G.; Bonazzi, R.; Lunetta, C.; Silani, V.; Riva, N.; Rigamonti, A.; Maggiani, A.; Ferrarese, C.; et al. Inefficient skeletal muscle oxidative function flanks impaired motor neuron recruitment in Amyotrophic Lateral Sclerosis during exercise. Sci. Rep. 2017, 7, 2951. [Google Scholar] [CrossRef] [PubMed]
- Finsterer, J. Lactate stress testing in sporadic amyotrophic lateral sclerosis. Int. J. Neurosci. 2005, 115, 583–591. [Google Scholar] [CrossRef]
- Zhang, Y.-J.; Fan, D.-S. Elimination Rate of Serum Lactate is Correlated with Amyotrophic Lateral Sclerosis Progression. Chin. Med. J. 2016, 129, 28–32. [Google Scholar] [CrossRef]
- Rabinowitz, J.D.; Enerbäck, S. Lactate: The ugly duckling of energy metabolism. Nat. Metab. 2020, 2, 566–571. [Google Scholar] [CrossRef]
- Brooks, S.J.; Parks, S.M.; Stamoulis, C. Widespread Positive Direct and Indirect Effects of Regular Physical Activity on the Developing Functional Connectome in Early Adolescence. Cereb. Cortex 2021, 31, bhab126. [Google Scholar] [CrossRef]
- Brooks, G.A.; Osmond, A.D.; Arevalo, J.A.; Duong, J.J.; Curl, C.C.; Moreno-Santillan, D.D.; Leija, R.G. Lactate as a myokine and exerkine: Drivers and signals of physiology and metabolism. J. Appl. Physiol. 2023, 134, 529–548. [Google Scholar] [CrossRef]
- Allen, P.J.; Brooks, G.A. Partial purification and reconstitution of the sarcolemmal L-lactate carrier from rat skeletal muscle. Biochem. J. 1994, 303, 207–212. [Google Scholar] [CrossRef]
- Magistretti, P.J.; Sorg, O.; Naichen, Y.; Pellerin, L.; de Rham, S.; Martin, J.L. Regulation of astrocyte energy metabolism by neurotransmitters. Ren. Physiol. Biochem. 1994, 17, 168–171. [Google Scholar] [CrossRef]
- Pellerin, L.; Pellegri, G.; Bittar, P.G.; Charnay, Y.; Bouras, C.; Martin, J.L.; Stella, N.; Magistretti, P.J. Evidence supporting the existence of an activity-dependent astrocyte-neuron lactate shuttle. Dev. Neurosci. 1998, 20, 291–299. [Google Scholar] [CrossRef] [PubMed]
- Magistretti, P.J.; Sorg, O.; Yu, N.; Martin, J.L.; Pellerin, L. Neurotransmitters regulate energy metabolism in astrocytes: Implications for the metabolic trafficking between neural cells. Dev. Neurosci. 1993, 15, 306–312. [Google Scholar] [CrossRef]
- Brooks, G.A. What the Lactate Shuttle Means for Sports Nutrition. Nutrients 2023, 15, 2178. [Google Scholar] [CrossRef]
- van Hall, G.; Strømstad, M.; Rasmussen, P.; Jans, O.; Zaar, M.; Gam, C.; Quistorff, B.; Secher, N.H.; Nielsen, H.B. Blood lactate is an important energy source for the human brain. J. Cereb. Blood Flow Metab. 2009, 29, 1121–1129. [Google Scholar] [CrossRef] [PubMed]
- Brooks, G.A.; Curl, C.C.; Leija, R.G.; Osmond, A.D.; Duong, J.J.; Arevalo, J.A. Tracing the lactate shuttle to the mitochondrial reticulum. Exp. Mol. Med. 2022, 54, 1332–1347. [Google Scholar] [CrossRef]
- Cori, C.F.; Cori, G.T. Carbohydrate metabolism. Annu. Rev. Biochem. 1946, 15, 193–218. [Google Scholar] [CrossRef] [PubMed]
- Brooks, G.A. The “Anaerobic Threshold” Concept Is Not Valid in Physiology and Medicine. Med. Sci. Sports Exerc. 2021, 53, 1093–1096. [Google Scholar] [CrossRef]
- Kwon, D.-N.; Choi, Y.-J.; Cho, S.-G.; Park, C.; Seo, H.G.; Song, H.; Kim, J.-H. CMP-Neu5Ac Hydroxylase Null Mice as a Model for Studying Metabolic Disorders Caused by the Evolutionary Loss of Neu5Gc in Humans. BioMed Res. Int. 2015, 2015, 830315. [Google Scholar] [CrossRef]
- Cluntun, A.A.; Visker, J.R.; Velasco-Silva, J.N.; Lang, M.J.; Cedeño-Rosario, L.; Shankar, T.S.; Hamouche, R.; Ling, J.; Kim, J.E.; Toshniwal, A.G.; et al. Direct mitochondrial import of lactate supports resilient carbohydrate oxidation. bioRxiv 2024. bioRxiv:2024.10.07.617073. [Google Scholar] [CrossRef]
- Brooks, G.A. The Science and Translation of Lactate Shuttle Theory. Cell Metab. 2018, 27, 757–785. [Google Scholar] [CrossRef]
- Pellerin, L.; Bouzier-Sore, A.-K.; Aubert, A.; Serres, S.; Merle, M.; Costalat, R.; Magistretti, P.J. Activity-dependent regulation of energy metabolism by astrocytes: An update. Glia 2007, 55, 1251–1262. [Google Scholar] [CrossRef] [PubMed]
- Schurr, A. Lactate: The ultimate cerebral oxidative energy substrate? J. Cereb. Blood Flow Metab. 2006, 26, 142–152. [Google Scholar] [CrossRef]
- Payne, R.S.; Schurr, A. Corticosterone disrupts glucose-, but not lactate-supported hippocampal PS-LTP. Neurosci. Lett. 2007, 424, 111–115. [Google Scholar] [CrossRef] [PubMed]
- Chamaa, F.; Magistretti, P.J.; Fiumelli, H. Astrocyte-derived lactate in stress disorders. Neurobiol. Dis. 2024, 192, 106417. [Google Scholar] [CrossRef]
- Karagiannis, A.; Gallopin, T.; Lacroix, A.; Plaisier, F.; Piquet, J.; Geoffroy, H.; Hepp, R.; Naudé, J.; Le Gac, B.; Egger, R.; et al. Lactate is an energy substrate for rodent cortical neurons and enhances their firing activity. elife 2021, 10, e71424. [Google Scholar] [CrossRef]
- Brooks, G.A.; Osmond, A.D.; Leija, R.G.; Curl, C.C.; Arevalo, J.A.; Duong, J.J.; Horning, M.A. The blood lactate/pyruvate equilibrium affair. Am. J. Physiol.-Endocrinol. Metab. 2022, 322, E34–E43. [Google Scholar] [CrossRef] [PubMed]
- Alexander, S.P.H.; Kelly, E.; Mathie, A.A.; Peters, J.A.; Veale, E.L.; Armstrong, J.F.; Buneman, O.P.; Faccenda, E.; Harding, S.D.; Spedding, M.; et al. The Concise Guide to PHARMACOLOGY 2023/24: Introduction and Other Protein Targets. Br. J. Pharmacol. 2023, 180 (Suppl. S2), S1–S22. [Google Scholar] [CrossRef]
- Griego, E.; Galván, E.J. BDNF and Lactate as Modulators of Hippocampal CA3 Network Physiology. Cell Mol. Neurobiol. 2023, 43, 4007–4022. [Google Scholar] [CrossRef]
- Herrera-López, G.; Griego, E.; Galván, E.J. Lactate induces synapse-specific potentiation on CA3 pyramidal cells of rat hippocampus. PLoS ONE 2020, 15, e0242309. [Google Scholar] [CrossRef] [PubMed]
- Brooks, G.A. Lactate as a fulcrum of metabolism. Redox Biol. 2020, 35, 101454. [Google Scholar] [CrossRef]
- Merkuri, F.; Rothstein, M.; Simoes-Costa, M. Histone lactylation couples cellular metabolism with developmental gene regulatory networks. Nat. Commun. 2024, 15, 90. [Google Scholar] [CrossRef]
- Monsorno, K.; Buckinx, A.; Paolicelli, R.C. Microglial metabolic flexibility: Emerging roles for lactate. Trends Endocrinol. Metab. 2022, 33, 186–195. [Google Scholar] [CrossRef] [PubMed]
- Shichkova, P.; Coggan, J.S.; Markram, H.; Keller, D. Brain Metabolism in Health and Neurodegeneration: The Interplay Among Neurons and Astrocytes. Cells 2024, 13, 1714. [Google Scholar] [CrossRef]
- Cauli, B.; Dusart, I.; Li, D. Lactate as a determinant of neuronal excitability, neuroenergetics and beyond. Neurobiol. Dis. 2023, 184, 106207. [Google Scholar] [CrossRef]
- Matsui, T.; Omuro, H.; Liu, Y.-F.; Soya, M.; Shima, T.; McEwen, B.S.; Soya, H. Astrocytic glycogen-derived lactate fuels the brain during exhaustive exercise to maintain endurance capacity. Proc. Natl. Acad. Sci. USA 2017, 114, 6358–6363. [Google Scholar] [CrossRef]
- Dodge, J.C.; Treleaven, C.M.; Fidler, J.A.; Tamsett, T.J.; Bao, C.; Searles, M.; Taksir, T.V.; Misra, K.; Sidman, R.L.; Cheng, S.H.; et al. Metabolic signatures of amyotrophic lateral sclerosis reveal insights into disease pathogenesis. Proc. Natl. Acad. Sci. USA 2013, 110, 10812–10817. [Google Scholar] [CrossRef]
- Morrison, B.M.; Lee, Y.; Rothstein, J.D. Oligodendroglia metabolically support axons and maintain structural integrity. Trends Cell Biol. 2013, 23, 644–651. [Google Scholar] [CrossRef]
- Traiffort, E.; Morisset-Lopez, S.; Moussaed, M.; Zahaf, A. Defective Oligodendroglial Lineage and Demyelination in Amyotrophic Lateral Sclerosis. Int. J. Mol. Sci. 2021, 22, 3426. [Google Scholar] [CrossRef] [PubMed]
- Lee, Y.; Morrison, B.M.; Li, Y.; Lengacher, S.; Farah, M.H.; Hoffman, P.N.; Liu, Y.; Tsingalia, A.; Jin, L.; Zhang, P.-W.; et al. Oligodendroglia metabolically support axons and contribute to neurodegeneration. Nature 2012, 487, 443–448. [Google Scholar] [CrossRef] [PubMed]
- Philips, T.; Mironova, Y.A.; Jouroukhin, Y.; Chew, J.; Vidensky, S.; Farah, M.H.; Pletnikov, M.V.; Bergles, D.E.; Morrison, B.M.; Rothstein, J.D. MCT1 Deletion in Oligodendrocyte Lineage Cells Causes Late-Onset Hypomyelination and Axonal Degeneration. Cell Rep. 2021, 34, 108610. [Google Scholar] [CrossRef]
- Bouçanova, F.; Pollmeier, G.; Sandor, K.; Morado Urbina, C.; Nijssen, J.; Médard, J.-J.; Bartesaghi, L.; Pellerin, L.; Svensson, C.I.; Hedlund, E.; et al. Disrupted function of lactate transporter MCT1, but not MCT4, in Schwann cells affects the maintenance of motor end-plate innervation. Glia 2021, 69, 124–136. [Google Scholar] [CrossRef]
- Susuki, K.; Baba, H.; Tohyama, K.; Kanai, K.; Kuwabara, S.; Hirata, K.; Furukawa, K.; Furukawa, K.; Rasband, M.N.; Yuki, N. Gangliosides contribute to stability of paranodal junctions and ion channel clusters in myelinated nerve fibers. Glia 2007, 55, 746–757. [Google Scholar] [CrossRef] [PubMed]
- Ganser, A.L.; Kirschner, D.A.; Willinger, M. Ganglioside localization on myelinated nerve fibres by cholera toxin binding. J. Neurocytol. 1983, 12, 921–938. [Google Scholar] [CrossRef]
- Ganser, A.L.; Kirschner, D.A. Differential expression of gangliosides on the surfaces of myelinated nerve fibers. J. Neurosci. Res. 1984, 12, 245–255. [Google Scholar] [CrossRef]
- Yuki, N.; Yanaka, C.; Sudo, M.; Funakoshi, M.; Ishida, H.; Mori, M.; Kanda, F.; Hirata, K. Lower motor neuron syndrome associated with IgG anti-GM1 antibodies revisited. J. Neuroimmunol. 2014, 272, 62–66. [Google Scholar] [CrossRef]
- Santoro, M.; Thomas, F.P.; Fink, M.E.; Lange, D.J.; Uncini, A.; Wadia, N.H.; Latov, N.; Hays, A.P. IgM deposits at nodes of Ranvier in a patient with amyotrophic lateral sclerosis, anti-GM1 antibodies, and multifocal motor conduction block. Ann. Neurol. 1990, 28, 373–377. [Google Scholar] [CrossRef]
- Deck, M.; Van Hameren, G.; Campbell, G.; Bernard-Marissal, N.; Devaux, J.; Berthelot, J.; Lattard, A.; Médard, J.-J.; Gautier, B.; Guelfi, S.; et al. Physiology of PNS axons relies on glycolytic metabolism in myelinating Schwann cells. PLoS ONE 2022, 17, e0272097. [Google Scholar] [CrossRef] [PubMed]
- Rich, L.R.; Ransom, B.R.; Brown, A.M. Energy Metabolism in Mouse Sciatic Nerve A Fibres during Increased Energy Demand. Metabolites 2022, 12, 505. [Google Scholar] [CrossRef]
- Riva, N.; Gentile, F.; Cerri, F.; Gallia, F.; Podini, P.; Dina, G.; Falzone, Y.M.; Fazio, R.; Lunetta, C.; Calvo, A.; et al. Phosphorylated TDP-43 aggregates in peripheral motor nerves of patients with amyotrophic lateral sclerosis. Brain 2022, 145, 276–284. [Google Scholar] [CrossRef] [PubMed]
- Quittmann, O.J.; Foitschik, T.; Vafa, R.; Freitag, F.J.; Sparmann, N.; Nolte, S.; Abel, T. Is Maximal Lactate Accumulation Rate Promising for Improving 5000-m Prediction in Running? Int. J. Sports Med. 2023, 44, 268–279. [Google Scholar] [CrossRef]
- Arevalo, J.A.; Leija, R.G.; Osmond, A.D.; Curl, C.C.; Duong, J.J.; Huie, M.J.; Masharani, U.; Brooks, G.A. Delayed and diminished postprandial lactate shuttling in healthy older men and women. Am. J. Physiol. Endocrinol. Metab. 2024, 327, E430–E440. [Google Scholar] [CrossRef]
- Marck, A.; Antero, J.; Berthelot, G.; Saulière, G.; Jancovici, J.-M.; Masson-Delmotte, V.; Boeuf, G.; Spedding, M.; Le Bourg, É.; Toussaint, J.-F. Are We Reaching the Limits of Homo sapiens? Front. Physiol. 2017, 8, 812. [Google Scholar] [CrossRef]
- Mezzani, A.; Pisano, F.; Cavalli, A.; Tommasi, M.A.; Corrà, U.; Colombo, S.; Grassi, B.; Marzorati, M.; Porcelli, S.; Morandi, L.; et al. Reduced exercise capacity in early-stage amyotrophic lateral sclerosis: Role of skeletal muscle. Amyotroph. Lateral Scler. 2012, 13, 87–94. [Google Scholar] [CrossRef]
- Goodwin, M.L.; Harris, J.E.; Hernández, A.; Gladden, L.B. Blood Lactate Measurements and Analysis during Exercise: A Guide for Clinicians. J. Diabetes Sci. Technol. 2007, 1, 558–569. [Google Scholar] [CrossRef] [PubMed]
- Siciliano, G.; Pastorini, E.; Pasquali, L.; Manca, M.L.; Iudice, A.; Murri, L. Impaired oxidative metabolism in exercising muscle from ALS patients. J. Neurol. Sci. 2001, 191, 61–65. [Google Scholar] [CrossRef]
- Abi-Saab, W.M.; Maggs, D.G.; Jones, T.; Jacob, R.; Srihari, V.; Thompson, J.; Kerr, D.; Leone, P.; Krystal, J.H.; Spencer, D.D.; et al. Striking differences in glucose and lactate levels between brain extracellular fluid and plasma in conscious human subjects: Effects of hyperglycemia and hypoglycemia. J. Cereb. Blood Flow Metab. 2002, 22, 271–279. [Google Scholar] [CrossRef] [PubMed]
- Pradat, P.-F.; Bruneteau, G.; Gordon, P.H.; Dupuis, L.; Bonnefont-Rousselot, D.; Simon, D.; Salachas, F.; Corcia, P.; Frochot, V.; Lacorte, J.-M.; et al. Impaired glucose tolerance in patients with amyotrophic lateral sclerosis. Amyotroph. Lateral Scler. 2010, 11, 166–171. [Google Scholar] [CrossRef]
- Huang, S.; Shangguan, R.; Chen, S.; Lai, X.; Han, H.; Sun, J. Mechanism of Fatty Acid Metabolism and Regulation by Lactate During Exercise in White Adipose and Skeletal Muscle Tissue: A Review. Sports Med. Open 2025, 11, 76. [Google Scholar] [CrossRef]
- Larrea, D.; Tamucci, K.A.; Kabra, K.; Velasco, K.R.; Yun, T.D.; Pera, M.; Montesinos, J.; Agrawal, R.R.; Paradas, C.; Smerdon, J.W.; et al. Altered mitochondria-associated ER membrane (MAM) function shifts mitochondrial metabolism in amyotrophic lateral sclerosis (ALS). Nat. Commun. 2025, 16, 379. [Google Scholar] [CrossRef] [PubMed]
- Vance, J.E. Historical perspective: Phosphatidylserine and phosphatidylethanolamine from the 1800s to the present. J. Lipid Res. 2018, 59, 923–944. [Google Scholar] [CrossRef]
- Vance, J.E. Newly made phosphatidylserine and phosphatidylethanolamine are preferentially translocated between rat liver mitochondria and endoplasmic reticulum. J. Biol. Chem. 1991, 266, 89–97. [Google Scholar] [CrossRef]
- Calzada, E.; Onguka, O.; Claypool, S.M. Phosphatidylethanolamine Metabolism in Health and Disease. Int. Rev. Cell Mol. Biol. 2016, 321, 29–88. [Google Scholar] [CrossRef]
- Boumezbeur, F.; Petersen, K.F.; Cline, G.W.; Mason, G.F.; Behar, K.L.; Shulman, G.I.; Rothman, D.L. The contribution of blood lactate to brain energy metabolism in humans measured by dynamic 13C nuclear magnetic resonance spectroscopy. J. Neurosci. 2010, 30, 13983–13991. [Google Scholar] [CrossRef]
- Magistretti, P.J.; Allaman, I. A Cellular Perspective on Brain Energy Metabolism and Functional Imaging. Neuron 2015, 86, 883–901. [Google Scholar] [CrossRef]
- Ngo, S.T.; Steyn, F.J. The interplay between metabolic homeostasis and neurodegeneration: Insights into the neurometabolic nature of amyotrophic lateral sclerosis. Cell Regen. 2015, 4, 5. [Google Scholar] [CrossRef]
- Cunnane, S.C.; Trushina, E.; Morland, C.; Prigione, A.; Casadesus, G.; Andrews, Z.B.; Beal, M.F.; Bergersen, L.H.; Brinton, R.D.; de la Monte, S.; et al. Brain energy rescue: An emerging therapeutic concept for neurodegenerative disorders of ageing. Nat. Rev. Drug Discov. 2020, 19, 609–633. [Google Scholar] [CrossRef]
- Flis, D.J.; Dzik, K.; Kaczor, J.J.; Halon-Golabek, M.; Antosiewicz, J.; Wieckowski, M.R.; Ziolkowski, W. Swim Training Modulates Skeletal Muscle Energy Metabolism, Oxidative Stress, and Mitochondrial Cholesterol Content in Amyotrophic Lateral Sclerosis Mice. Oxid. Med. Cell Longev. 2018, 2018, 5940748. [Google Scholar] [CrossRef] [PubMed]
- Ferraiuolo, L.; Higginbottom, A.; Heath, P.R.; Barber, S.; Greenald, D.; Kirby, J.; Shaw, P.J. Dysregulation of astrocyte-motoneuron cross-talk in mutant superoxide dismutase 1-related amyotrophic lateral sclerosis. Brain 2011, 134, 2627–2641. [Google Scholar] [CrossRef] [PubMed]
- Jourdain, P.; Allaman, I.; Rothenfusser, K.; Fiumelli, H.; Marquet, P.; Magistretti, P.J. L-Lactate protects neurons against excitotoxicity: Implication of an ATP-mediated signaling cascade. Sci. Rep. 2016, 6, 21250. [Google Scholar] [CrossRef]
- Gerou, M.; Hall, B.; Woof, R.; Allsop, J.; Kolb, S.J.; Meyer, K.; Shaw, P.J.; Allen, S.P. Amyotrophic lateral sclerosis alters the metabolic aging profile in patient derived fibroblasts. Neurobiol. Aging 2021, 105, 64–77. [Google Scholar] [CrossRef]
- Petit, J.-M.; Eren-Koçak, E.; Karatas, H.; Magistretti, P.; Dalkara, T. Brain glycogen metabolism: A possible link between sleep disturbances, headache and depression. Sleep Med. Rev. 2021, 59, 101449. [Google Scholar] [CrossRef] [PubMed]
- Ros, J.; Pecinska, N.; Alessandri, B.; Landolt, H.; Fillenz, M. Lactate reduces glutamate-induced neurotoxicity in rat cortex. J. Neurosci. Res. 2001, 66, 790–794. [Google Scholar] [CrossRef] [PubMed]
- Wahis, J.; Holt, M.G. Astrocytes, Noradrenaline, α1-Adrenoreceptors, and Neuromodulation: Evidence and Unanswered Questions. Front. Cell Neurosci. 2021, 15, 645691. [Google Scholar] [CrossRef] [PubMed]
- Bista, S.; Coffey, A.; Mitchell, M.; Fasano, A.; Dukic, S.; Buxo, T.; Giglia, E.; Heverin, M.; Muthuraman, M.; Carson, R.G.; et al. Abnormal EEG spectral power and coherence measures during pre-motor stage in Amyotrophic Lateral Sclerosis. IEEE Trans. Neural Syst. Rehabil. Eng. 2024, 33, 232–242. [Google Scholar] [CrossRef]
- Scekic-Zahirovic, J.; Benetton, C.; Brunet, A.; Ye, X.; Logunov, E.; Douchamps, V.; Megat, S.; Andry, V.; Kan, V.W.Y.; Stuart-Lopez, G.; et al. Cortical hyperexcitability in mouse models and patients with amyotrophic lateral sclerosis is linked to noradrenaline deficiency. Sci. Transl. Med. 2024, 16, eadg3665. [Google Scholar] [CrossRef]
- Turner, B.J.; Rembach, A.; Spark, R.; Lopes, E.C.; Cheema, S.S. Opposing effects of low and high-dose clozapine on survival of transgenic amyotrophic lateral sclerosis mice. J. Neurosci. Res. 2003, 74, 605–613. [Google Scholar] [CrossRef]
- Spedding, M.; Sebban, C.; Jay, T.M.; Rocher, C.; Tesolin-Decros, B.; Chazot, P.; Schenker, E.; Szénási, G.; Lévay, G.I.; Megyeri, K.; et al. Phenotypical Screening on Neuronal Plasticity in Hippocampal-Prefrontal Cortex Connectivity Reveals an Antipsychotic with a Novel Profile. Cells 2022, 11, 1181. [Google Scholar] [CrossRef]
- Vandoorne, T.; Veys, K.; Guo, W.; Sicart, A.; Vints, K.; Swijsen, A.; Moisse, M.; Eelen, G.; Gounko, N.V.; Fumagalli, L.; et al. Differentiation but not ALS mutations in FUS rewires motor neuron metabolism. Nat. Commun. 2019, 10, 4147. [Google Scholar] [CrossRef]
- Hor, J.-H.; Santosa, M.M.; Lim, V.J.W.; Ho, B.X.; Taylor, A.; Khong, Z.J.; Ravits, J.; Fan, Y.; Liou, Y.-C.; Soh, B.-S.; et al. ALS motor neurons exhibit hallmark metabolic defects that are rescued by SIRT3 activation. Cell Death Differ. 2021, 28, 1379–1397. [Google Scholar] [CrossRef] [PubMed]
- Mehta, A.R.; Gregory, J.M.; Dando, O.; Carter, R.N.; Burr, K.; Nanda, J.; Story, D.; McDade, K.; Smith, C.; Morton, N.M.; et al. Mitochondrial bioenergetic deficits in C9orf72 amyotrophic lateral sclerosis motor neurons cause dysfunctional axonal homeostasis. Acta Neuropathol. 2021, 141, 257–279. [Google Scholar] [CrossRef] [PubMed]
- Jankovic, M.; Novakovic, I.; Gamil Anwar Dawod, P.; Gamil Anwar Dawod, A.; Drinic, A.; Abdel Motaleb, F.I.; Ducic, S.; Nikolic, D. Current Concepts on Genetic Aspects of Mitochondrial Dysfunction in Amyotrophic Lateral Sclerosis. Int. J. Mol. Sci. 2021, 22, 9832. [Google Scholar] [CrossRef]
- Cieminski, K.; Flis, D.J.; Dzik, K.; Kaczor, J.J.; Czyrko, E.; Halon-Golabek, M.; Wieckowski, M.R.; Antosiewicz, J.; Ziolkowski, W. Swim training affects Akt signaling and ameliorates loss of skeletal muscle mass in a mouse model of amyotrophic lateral sclerosis. Sci. Rep. 2021, 11, 20899. [Google Scholar] [CrossRef] [PubMed]
- Cieminski, K.; Flis, D.J.; Dzik, K.P.; Kaczor, J.J.; Wieckowski, M.R.; Antosiewicz, J.; Ziolkowski, W. Swim Training Affects on Muscle Lactate Metabolism, Nicotinamide Adenine Dinucleotides Concentration, and the Activity of NADH Shuttle Enzymes in a Mouse Model of Amyotrophic Lateral Sclerosis. Int. J. Mol. Sci. 2022, 23, 11504. [Google Scholar] [CrossRef]
- Coco, M. The brain behaves as a muscle? Neurol. Sci. 2017, 38, 1865–1868. [Google Scholar] [CrossRef]
- Coco, M.; Buscemi, A.; Ramaci, T.; Tusak, M.; Corrado, D.D.; Perciavalle, V.; Maugeri, G.; Perciavalle, V.; Musumeci, G. Influences of Blood Lactate Levels on Cognitive Domains and Physical Health during a Sports Stress. Brief Review. Int. J. Environ. Res. Public Health 2020, 17, 9043. [Google Scholar] [CrossRef]
- Ranieri, F.; Senerchia, G.; Bonan, L.; Casali, S.; Cabona, C.; Cantone, M.; De Marchi, F.; Diamanti, L.; Doretti, A.; Fini, N.; et al. Cortical Excitability as a Prognostic and Phenotypic Stratification Biomarker in Amyotrophic Lateral Sclerosis. Ann. Neurol. 2025. [Google Scholar] [CrossRef]
- Vucic, S.; Nicholson, G.A.; Kiernan, M.C. Cortical hyperexcitability may precede the onset of familial amyotrophic lateral sclerosis. Brain 2008, 131, 1540–1550. [Google Scholar] [CrossRef]
- Coco, M.; Alagona, G.; Rapisarda, G.; Costanzo, E.; Calogero, R.A.; Perciavalle, V.; Perciavalle, V. Elevated blood lactate is associated with increased motor cortex excitability. Somatosens. Mot. Res. 2010, 27, 1–8. [Google Scholar] [CrossRef]
- Alagona, G.; Coco, M.; Rapisarda, G.; Costanzo, E.; Maci, T.; Restivo, D.; Maugeri, A.; Perciavalle, V. Changes of blood lactate levels after repetitive transcranial magnetic stimulation. Neurosci. Lett. 2009, 450, 111–113. [Google Scholar] [CrossRef]
- Chapman, L.; Cooper-Knock, J.; Shaw, P.J. Physical activity as an exogenous risk factor for amyotrophic lateral sclerosis: A review of the evidence. Brain 2023, 146, 1745–1757. [Google Scholar] [CrossRef]
- Gallo, V.; Vanacore, N.; Bueno-de-Mesquita, H.B.; Vermeulen, R.; Brayne, C.; Pearce, N.; Wark, P.A.; Ward, H.A.; Ferrari, P.; Jenab, M.; et al. Physical activity and risk of Amyotrophic Lateral Sclerosis in a prospective cohort study. Eur. J. Epidemiol. 2016, 31, 255–266. [Google Scholar] [CrossRef]
- Vaage, A.M.; Meyer, H.E.; Landgraff, I.K.; Myrstad, M.; Holmøy, T.; Nakken, O. Physical Activity, Fitness, and Long-Term Risk of Amyotrophic Lateral Sclerosis: A Prospective Cohort Study. Neurology 2024, 103, e209575. [Google Scholar] [CrossRef] [PubMed]
- Julian, T.H.; Glascow, N.; Barry, A.D.F.; Moll, T.; Harvey, C.; Klimentidis, Y.C.; Newell, M.; Zhang, S.; Snyder, M.P.; Cooper-Knock, J.; et al. Physical exercise is a risk factor for amyotrophic lateral sclerosis: Convergent evidence from Mendelian randomisation, transcriptomics and risk genotypes. EBioMedicine 2021, 68, 103397. [Google Scholar] [CrossRef] [PubMed]
- Zulueta, A.; Piras, R.; Azzolino, D.; Mariani, P.; Sideri, R.; Garrè, C.; Federico, G.; Lucchi, T.; Magni, P.; Parati, E.A.; et al. Neurofilament Light Chain Levels, Skeletal Muscle Loss, and Nutritional Decline: Key Prognostic Factors in Amyotrophic Lateral Sclerosis. Muscle Nerve 2025, 72, 49–55. [Google Scholar] [CrossRef] [PubMed]
- Tosolini, A.P.; Sleigh, J.N.; Surana, S.; Rhymes, E.R.; Cahalan, S.D.; Schiavo, G. BDNF-dependent modulation of axonal transport is selectively impaired in ALS. Acta Neuropathol. Commun. 2022, 10, 121. [Google Scholar] [CrossRef]
- Sleigh, J.N.; Villarroel-Campos, D.; Surana, S.; Wickenden, T.; Tong, Y.; Simkin, R.L.; Vargas, J.N.S.; Rhymes, E.R.; Tosolini, A.P.; West, S.J.; et al. Boosting peripheral BDNF rescues impaired in vivo axonal transport in CMT2D mice. JCI Insight 2023, 8, e157191. [Google Scholar] [CrossRef]
- Mc Cluskey, M.; Dubouchaud, H.; Nicot, A.-S.; Saudou, F. A vesicular Warburg effect: Aerobic glycolysis occurs on axonal vesicles for local NAD+ recycling and transport. Traffic 2024, 25, e12926. [Google Scholar] [CrossRef]
- Ni, Y.-F.; Zhang, W.; Bao, X.-F.; Wang, W.; Song, L.; Jiang, B. GM1 ganglioside reverses the cognitive deficits induced by MK801 in mice. Behav. Pharmacol. 2016, 27, 451–459. [Google Scholar] [CrossRef]
- Valdomero, A.; Perondi, M.C.; Orsingher, O.A.; Cuadra, G.R. Exogenous GM1 ganglioside increases accumbal BDNF levels in rats. Behav. Brain Res. 2015, 278, 303–306. [Google Scholar] [CrossRef] [PubMed]
- Lim, S.T.; Esfahani, K.; Avdoshina, V.; Mocchetti, I. Exogenous gangliosides increase the release of brain-derived neurotrophic factor. Neuropharmacology 2011, 60, 1160–1167. [Google Scholar] [CrossRef] [PubMed]
- Harvey, C.; Weinreich, M.; Lee, J.A.K.; Shaw, A.C.; Ferraiuolo, L.; Mortiboys, H.; Zhang, S.; Hop, P.J.; Zwamborn, R.A.J.; van Eijk, K.; et al. Rare and common genetic determinants of mitochondrial function determine severity but not risk of amyotrophic lateral sclerosis. Heliyon 2024, 10, e24975. [Google Scholar] [CrossRef]
- Rifai, O.M.; O’Shaughnessy, J.; Dando, O.R.; Munro, A.F.; Sewell, M.D.E.; Abrahams, S.; Waldron, F.M.; Sibley, C.R.; Gregory, J.M. Distinct neuroinflammatory signatures exist across genetic and sporadic amyotrophic lateral sclerosis cohorts. Brain 2023, 146, 5124–5138. [Google Scholar] [CrossRef]
- Lehman, E.J.; Hein, M.J.; Baron, S.L.; Gersic, C.M. Neurodegenerative causes of death among retired National Football League players. Neurology 2012, 79, 1970–1974. [Google Scholar] [CrossRef]
- Daneshvar, D.H.; Mez, J.; Alosco, M.L.; Baucom, Z.H.; Mahar, I.; Baugh, C.M.; Valle, J.P.; Weuve, J.; Paganoni, S.; Cantu, R.C.; et al. Incidence of and Mortality from Amyotrophic Lateral Sclerosis in National Football League Athletes. JAMA Netw. Open 2021, 4, e2138801. [Google Scholar] [CrossRef]
- Chio, A.; Calvo, A.; Dossena, M.; Ghiglione, P.; Mutani, R.; Mora, G. ALS in Italian professional soccer players: The risk is still present and could be soccer-specific. Amyotroph. Lateral Scler. 2009, 10, 205–209. [Google Scholar] [CrossRef] [PubMed]
- Pupillo, E.; Messina, P.; Giussani, G.; Logroscino, G.; Zoccolella, S.; Chiò, A.; Calvo, A.; Corbo, M.; Lunetta, C.; Marin, B.; et al. Physical activity and amyotrophic lateral sclerosis: A European population-based case-control study. Ann. Neurol. 2014, 75, 708–716. [Google Scholar] [CrossRef]
- McKee, A.C.; Gavett, B.E.; Stern, R.A.; Nowinski, C.J.; Cantu, R.C.; Kowall, N.W.; Perl, D.P.; Hedley-Whyte, E.T.; Price, B.; Sullivan, C.; et al. TDP-43 proteinopathy and motor neuron disease in chronic traumatic encephalopathy. J. Neuropathol. Exp. Neurol. 2010, 69, 918–929. [Google Scholar] [CrossRef]
- Blecher, R.; Elliott, M.A.; Yilmaz, E.; Dettori, J.R.; Oskouian, R.J.; Patel, A.; Clarke, A.; Hutton, M.; McGuire, R.; Dunn, R.; et al. Contact Sports as a Risk Factor for Amyotrophic Lateral Sclerosis: A Systematic Review. Glob. Spine J. 2019, 9, 104–118. [Google Scholar] [CrossRef]
- Fang, F.; Hållmarker, U.; James, S.; Ingre, C.; Michaëlsson, K.; Ahlbom, A.; Feychting, M. Amyotrophic lateral sclerosis among cross-country skiers in Sweden. Eur. J. Epidemiol. 2016, 31, 247–253. [Google Scholar] [CrossRef] [PubMed]
- Schmitt, F.; Hussain, G.; Dupuis, L.; Loeffler, J.-P.; Henriques, A. A plural role for lipids in motor neuron diseases: Energy, signaling and structure. Front. Cell. Neurosci. 2014, 8, 25. [Google Scholar] [CrossRef] [PubMed]
- Nilsson, J.; Thorstensson, A. Ground reaction forces at different speeds of human walking and running. Acta Physiol. Scand. 1989, 136, 217–227. [Google Scholar] [CrossRef]
- Standley, R.A.; Distefano, G.; Trevino, M.B.; Chen, E.; Narain, N.R.; Greenwood, B.; Kondakci, G.; Tolstikov, V.V.; Kiebish, M.A.; Yu, G.; et al. Skeletal Muscle Energetics and Mitochondrial Function Are Impaired Following 10 Days of Bed Rest in Older Adults. J. Gerontol. A Biol. Sci. Med. Sci. 2020, 75, 1744–1753. [Google Scholar] [CrossRef] [PubMed]
- Dittmer, D.K.; Teasell, R. Complications of immobilization and bed rest. Part 1: Musculoskeletal and cardiovascular complications. Can. Fam. Physician 1993, 39, 1428–1437. [Google Scholar]
- Eggelbusch, M.; Charlton, B.T.; Bosutti, A.; Ganse, B.; Giakoumaki, I.; Grootemaat, A.E.; Hendrickse, P.W.; Jaspers, Y.; Kemp, S.; Kerkhoff, T.J.; et al. The impact of bed rest on human skeletal muscle metabolism. Cell Rep. Med. 2024, 5, 101372. [Google Scholar] [CrossRef]
- Siciliano, G.; D’Avino, C.; Del Corona, A.; Barsacchi, R.; Kusmic, C.; Rocchi, A.; Pastorini, E.; Murri, L. Impaired oxidative metabolism and lipid peroxidation in exercising muscle from ALS patients. Amyotroph. Lateral Scler. Other Motor Neuron Disord. 2002, 3, 57–62. [Google Scholar] [CrossRef]
- Bayer, H.; Lang, K.; Buck, E.; Higelin, J.; Barteczko, L.; Pasquarelli, N.; Sprissler, J.; Lucas, T.; Holzmann, K.; Demestre, M.; et al. ALS-causing mutations differentially affect PGC-1α expression and function in the brain vs. peripheral tissues. Neurobiol. Dis. 2017, 97, 36–45. [Google Scholar] [CrossRef]
- Park, D.; Kwak, S.G.; Park, J.-S.; Choo, Y.J.; Chang, M.C. Can Therapeutic Exercise Slow Down Progressive Functional Decline in Patients with Amyotrophic Lateral Sclerosis? A Meta-Analysis. Front. Neurol. 2020, 11, 853. [Google Scholar] [CrossRef]
- Kitano, K.; Asakawa, T.; Kamide, N.; Yorimoto, K.; Yoneda, M.; Kikuchi, Y.; Sawada, M.; Komori, T. Effectiveness of Home-Based Exercises Without Supervision by Physical Therapists for Patients with Early-Stage Amyotrophic Lateral Sclerosis: A Pilot Study. Arch. Phys. Med. Rehabil. 2018, 99, 2114–2117. [Google Scholar] [CrossRef]
- Bello-Haas, V.D.; Florence, J.M.; Kloos, A.D.; Scheirbecker, J.; Lopate, G.; Hayes, S.M.; Pioro, E.P.; Mitsumoto, H. A randomized controlled trial of resistance exercise in individuals with ALS. Neurology 2007, 68, 2003–2007. [Google Scholar] [CrossRef]
- Dal Bello-Haas, V.; Florence, J.M. Therapeutic exercise for people with amyotrophic lateral sclerosis or motor neuron disease. Cochrane Database Syst. Rev. 2013, 2013, CD005229. [Google Scholar] [CrossRef] [PubMed]
- Ferri, A.; Lanfranconi, F.; Corna, G.; Bonazzi, R.; Marchese, S.; Magnoni, A.; Tremolizzo, L. Tailored Exercise Training Counteracts Muscle Disuse and Attenuates Reductions in Physical Function in Individuals with Amyotrophic Lateral Sclerosis. Front. Physiol. 2019, 10, 1537. [Google Scholar] [CrossRef]
- Fenili, G.; Scaricamazza, S.; Ferri, A.; Valle, C.; Paronetto, M.P. Physical exercise in amyotrophic lateral sclerosis: A potential co-adjuvant therapeutic option to counteract disease progression. Front. Cell Dev. Biol. 2024, 12, 1421566. [Google Scholar] [CrossRef] [PubMed]
- Scaricamazza, S.; Nesci, V.; Salvatori, I.; Fenili, G.; Rosina, M.; Gloriani, M.; Paronetto, M.P.; Madaro, L.; Ferri, A.; Valle, C. Endurance exercise has a negative impact on the onset of SOD1-G93A ALS in female mice and affects the entire skeletal muscle-motor neuron axis. Front. Pharmacol. 2024, 15, 1360099. [Google Scholar] [CrossRef] [PubMed]
- Markham, A.; Cameron, I.; Franklin, P.; Spedding, M. BDNF increases rat brain mitochondrial respiratory coupling at complex I, but not complex II. Eur. J. Neurosci. 2004, 20, 1189–1196. [Google Scholar] [CrossRef]
- McCluskey, G.; Morrison, K.E.; Donaghy, C.; Rene, F.; Duddy, W.; Duguez, S. Extracellular Vesicles in Amyotrophic Lateral Sclerosis. Life 2022, 13, 121. [Google Scholar] [CrossRef]
- Spendiff, S.; Vuda, M.; Gouspillou, G.; Aare, S.; Perez, A.; Morais, J.A.; Jagoe, R.T.; Filion, M.-E.; Glicksman, R.; Kapchinsky, S.; et al. Denervation drives mitochondrial dysfunction in skeletal muscle of octogenarians. J. Physiol. 2016, 594, 7361–7379. [Google Scholar] [CrossRef]
- Power, G.A.; Allen, M.D.; Gilmore, K.J.; Stashuk, D.W.; Doherty, T.J.; Hepple, R.T.; Taivassalo, T.; Rice, C.L. Motor unit number and transmission stability in octogenarian world class athletes: Can age-related deficits be outrun? J. Appl. Physiol. (1985) 2016, 121, 1013–1020. [Google Scholar] [CrossRef]
- Sonjak, V.; Jacob, K.; Morais, J.A.; Rivera-Zengotita, M.; Spendiff, S.; Spake, C.; Taivassalo, T.; Chevalier, S.; Hepple, R.T. Fidelity of muscle fibre reinnervation modulates ageing muscle impact in elderly women. J. Physiol. 2019, 597, 5009–5023. [Google Scholar] [CrossRef]
- Potvin-Desrochers, A.; Atri, A.; Clouette, J.; Hepple, R.T.; Taivassalo, T.; Paquette, C. Resting-state Functional Connectivity of the Motor and Cognitive Areas is Preserved in Masters Athletes. Neuroscience 2024, 546, 53–62. [Google Scholar] [CrossRef]
- Mazza, J.C.; Festa, R.R.; Gurovich, A.N.; Jannas-Vela, S. Blood Lactate Steady State during Interval Training: New Perspectives on Something Already Known. Int. J. Exerc. Sci. 2024, 17, 941–953. [Google Scholar] [CrossRef] [PubMed]
- Gold, S.M.; Schulz, K.-H.; Hartmann, S.; Mladek, M.; Lang, U.E.; Hellweg, R.; Reer, R.; Braumann, K.-M.; Heesen, C. Basal serum levels and reactivity of nerve growth factor and brain-derived neurotrophic factor to standardized acute exercise in multiple sclerosis and controls. J. Neuroimmunol. 2003, 138, 99–105. [Google Scholar] [CrossRef]
- El Hayek, L.; Khalifeh, M.; Zibara, V.; Abi Assaad, R.; Emmanuel, N.; Karnib, N.; El-Ghandour, R.; Nasrallah, P.; Bilen, M.; Ibrahim, P.; et al. Lactate Mediates the Effects of Exercise on Learning and Memory through SIRT1-Dependent Activation of Hippocampal Brain-Derived Neurotrophic Factor (BDNF). J. Neurosci. 2019, 39, 2369–2382. [Google Scholar] [CrossRef]
- Paquet Luzy, C.; Doppler, E.; Polasek, T.M.; Giorgino, R. First-in-human single-dose study of nizubaglustat, a dual inhibitor of ceramide glucosyltransferase and non-lysosomal glucosylceramidase: Safety, tolerability, pharmacokinetics, and pharmacodynamics of single ascending and multiple doses in healthy adults. Mol. Genet. Metab. 2024, 141, 108113. [Google Scholar] [CrossRef]
- Su, Q.; Louwerse, M.; Lammers, R.F.; Maurits, E.; Janssen, M.; Boot, R.G.; Borlandelli, V.; Offen, W.A.; Linzel, D.; Schröder, S.P.; et al. Selective labelling of GBA2 in cells with fluorescent β-d-arabinofuranosyl cyclitol aziridines. Chem. Sci. 2024, 15, 15212–15220. [Google Scholar] [CrossRef]
- Ambrosi, G.; Ghezzi, C.; Zangaglia, R.; Levandis, G.; Pacchetti, C.; Blandini, F. Ambroxol-induced rescue of defective glucocerebrosidase is associated with increased LIMP-2 and saposin C levels in GBA1 mutant Parkinson’s disease cells. Neurobiol Dis. 2015, 82, 235–242. [Google Scholar] [CrossRef]
- Gojda, J.; Waldauf, P.; Hrušková, N.; Blahutová, B.; Krajčová, A.; Urban, T.; Tůma, P.; Řasová, K.; Duška, F. Lactate production without hypoxia in skeletal muscle during electrical cycling: Crossover study of femoral venous-arterial differences in healthy volunteers. PLoS ONE 2019, 14, e0200228. [Google Scholar] [CrossRef]
- Maier, A.; Gaudlitz, M.; Grehl, T.; Weyen, U.; Steinbach, R.; Grosskreutz, J.; Rödiger, A.; Koch, J.C.; Lengenfeld, T.; Weydt, P.; et al. Use and subjective experience of the impact of motor-assisted movement exercisers in people with amyotrophic lateral sclerosis: A multicenter observational study. Sci. Rep. 2022, 12, 9657. [Google Scholar] [CrossRef] [PubMed]
- Noll, K.; Dowdell, B.T.; Ridgel, A.L. Mobility Improvements After a High-cadence Dynamic Cycling Intervention in an Individual with Motor Neuron Disease: A Case Study. Int. J. Exerc. Sci. 2021, 14, 791–801. [Google Scholar] [CrossRef] [PubMed]
- Joshi, P.; Shigo, L.; Smith, B.; Kilbane, C.W.; Guha, A.; Loparo, K.; Ridgel, A.L.; Shaikh, A.G. Electrophysiological correlates of dynamic cycling in Parkinson’s disease. Clin. Neurophysiol. 2025, 174, 17–27. [Google Scholar] [CrossRef]
- Kathmann, I.; Cizinauskas, S.; Doherr, M.G.; Steffen, F.; Jaggy, A. Daily Controlled Physiotherapy Increases Survival Time in Dogs with Suspected Degenerative Myelopathy. J. Vet. Intern. Med. 2006, 20, 927–932. [Google Scholar] [CrossRef]
- Perera, R.; Riley, C.; Isaac, G.; Hopf-Jannasch, A.S.; Moore, R.J.; Weitz, K.W.; Pasa-Tolic, L.; Metz, T.O.; Adamec, J.; Kuhn, R.J. Dengue virus infection perturbs lipid homeostasis in infected mosquito cells. PLoS Pathog. 2012, 8, e1002584. [Google Scholar] [CrossRef]
- Bjornevik, K.; Münz, C.; Cohen, J.I.; Ascherio, A. Epstein–Barr virus as a leading cause of multiple sclerosis: Mechanisms and implications. Nat. Rev. Neurol. 2023, 19, 160–171. [Google Scholar] [CrossRef] [PubMed]
- Gao, J.; Sterling, E.; Hankin, R.; Sikal, A.; Yao, Y. Therapeutics Targeting Skeletal Muscle in Amyotrophic Lateral Sclerosis. Biomolecules 2024, 14, 878. [Google Scholar] [CrossRef] [PubMed]
- Bouscary, A.; Quessada, C.; René, F.; Spedding, M.; Henriques, A.; Ngo, S.; Loeffler, J.-P. Drug repositioning in neurodegeneration: An overview of the use of ambroxol in neurodegenerative diseases. Eur. J. Pharmacol. 2020, 884, 173446. [Google Scholar] [CrossRef] [PubMed]
- Corr, P.B.; Gross, R.W.; Sobel, B.E. Amphipathic metabolites and membrane dysfunction in ischemic myocardium. Circ. Res. 1984, 55, 135–154. [Google Scholar] [CrossRef]
- Corr, P.B.; Saffitz, J.E.; Sobel, B.E. Lysophospholipids, long chain acylcarnitines and membrane dysfunction in the ischaemic heart. Basic Res. Cardiol. 1987, 82 (Suppl. S1), 199–208. [Google Scholar] [CrossRef]
- Corr, P.B.; Creer, M.H.; Yamada, K.A.; Saffitz, J.E.; Sobel, B.E. Prophylaxis of early ventricular fibrillation by inhibition of acylcarnitine accumulation. J. Clin. Investig. 1989, 83, 927–936. [Google Scholar] [CrossRef]
- Clarke, B.; Spedding, M.; Patmore, L.; McCormack, J.G. Protective effects of ranolazine in guinea-pig hearts during low-flow ischaemia and their association with increases in active pyruvate dehydrogenase. Br. J. Pharmacol. 1993, 109, 748–750. [Google Scholar] [CrossRef]
- Stanley, W.C. Partial fatty acid oxidation inhibitors for stable angina. Expert Opin. Investig. Drugs 2002, 11, 615–629. [Google Scholar] [CrossRef]
- Salvatori, I.; Nesci, V.; Spalloni, A.; Marabitti, V.; Muzzi, M.; Zenuni, H.; Scaricamazza, S.; Rosina, M.; Fenili, G.; Goglia, M.; et al. Trimetazidine Improves Mitochondrial Dysfunction in SOD1G93A Cellular Models of Amyotrophic Lateral Sclerosis through Autophagy Activation. Int. J. Mol. Sci. 2024, 25, 3251. [Google Scholar] [CrossRef] [PubMed]
- Kantor, P.F.; Lucien, A.; Kozak, R.; Lopaschuk, G.D. The antianginal drug trimetazidine shifts cardiac energy metabolism from fatty acid oxidation to glucose oxidation by inhibiting mitochondrial long-chain 3-ketoacyl coenzyme A thiolase. Circ. Res. 2000, 86, 580–588. [Google Scholar] [CrossRef]
- Lopaschuk, G.D.; Barr, R.; Thomas, P.D.; Dyck, J.R.B. Beneficial effects of trimetazidine in ex vivo working ischemic hearts are due to a stimulation of glucose oxidation secondary to inhibition of long-chain 3-ketoacyl coenzyme a thiolase. Circ. Res. 2003, 93, e33–e37. [Google Scholar] [CrossRef]
- Pușcaș, A.; Ștefănescu, R.; Vari, C.-E.; Ősz, B.-E.; Filip, C.; Bitzan, J.K.; Buț, M.-G.; Tero-Vescan, A. Biochemical Aspects That Lead to Abusive Use of Trimetazidine in Performance Athletes: A Mini-Review. Int. J. Mol. Sci. 2024, 25, 1605. [Google Scholar] [CrossRef] [PubMed]
- Dy, A.M.B.; Limjoco, L.L.G.; Jamora, R.D.G. Trimetazidine-Induced Parkinsonism: A Systematic Review. Front. Neurol. 2020, 11, 44. [Google Scholar] [CrossRef]
- van der Pol, K.H.; Aljofan, M.; Blin, O.; Cornel, J.H.; Rongen, G.A.; Woestelandt, A.-G.; Spedding, M. Drug Repurposing of Generic Drugs: Challenges and the Potential Role for Government. Appl. Health Econ. Health Policy 2023, 21, 831–840. [Google Scholar] [CrossRef] [PubMed]




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Spedding, M. Does Amyotrophic Lateral Sclerosis (ALS) Have Metabolic Causes from Human Evolution? Cells 2025, 14, 1734. https://doi.org/10.3390/cells14211734
Spedding M. Does Amyotrophic Lateral Sclerosis (ALS) Have Metabolic Causes from Human Evolution? Cells. 2025; 14(21):1734. https://doi.org/10.3390/cells14211734
Chicago/Turabian StyleSpedding, Michael. 2025. "Does Amyotrophic Lateral Sclerosis (ALS) Have Metabolic Causes from Human Evolution?" Cells 14, no. 21: 1734. https://doi.org/10.3390/cells14211734
APA StyleSpedding, M. (2025). Does Amyotrophic Lateral Sclerosis (ALS) Have Metabolic Causes from Human Evolution? Cells, 14(21), 1734. https://doi.org/10.3390/cells14211734
