Durant ces 3 années, j’ai voulu mettre en relation les phénotypes moteurs et cognitifs observés dans notre modèle murin Spg11-/- avec, d’une part, la fonctionnalité des réseaux neuronaux sous-tendants ces fonctions, et d’autre part, les dysfonctions intracellulaires révélées par les études précédentes. Dans cette optique, l’utilisation de techniques d’électrophysiologie m’a paru le plus judicieux malgré les contraintes et limitations qu’elles peuvent présenter. A ma connaissance, c’est la première fois que des données d’EEG sont générées à partir d’un modèle murin de PSH et qui permettent de révéler des évènements anormaux à l’EEG. Ces résultats ont d’ailleurs poussé les cliniciens de notre Institut à systématiser la mesure de l’activité corticale spontanée des patients SPG11 par EEG de surface. Si ces évènements EEG anormaux sont retrouvés chez les patients, et s’il se trouve qu’ils répondent à la thérapie génique expérimentée en ce moment dans notre laboratoire, nous pourrons proposer ces signaux comme un marqueur (non spécifique mais indicatif) de la PSH-SPG11 mais aussi comme une évaluation non invasive de l’efficacité d’essais thérapeutiques. De plus, ces résultats nous amènent à étudier l’ethosuximide comme molécule thérapeutique potentielle pour le traitement de certains symptôme de la PSH-SPG11. Si les données obtenues sur le réseau hippocampique Spg11-/- nécessitent d’autres expériences pour être expoitables, elles corroborent les données comportementales et histologiques précédentes et fournissent des pistes dans les mécanismes d’apparition des symptômes cognitifs observés dans notre modèle.
En conclusion, ce projet est le premier à s’intéresser à la PSH-SPG11 d’un point de vue plus électrophysiologique et réseautique que les études précédentes qui se concentraient plus sur la pathologie d’un point de vue biologique et intracellulaire. De futures études seront nécessaires pour valider ou invalider nos résultats et se pencher sur les nombreuses questions intéressantes soulevées par notre approche.
Références
1. Ellenbroek, B. & Youn, J. Rodent models in neuroscience research: is it a rat race? Dis. Model. Mech. 9, 1079–1087 (2016).
2. Barker, A. T., Jalinous, R. & Freeston, I. L. Non-invasive magnetic stimulation of human motor cortex. Lancet Lond. Engl. 1, 1106–1107 (1985).
3. Fortin, D. et al. Tractography in the Study of the Human Brain: A Neurosurgical Perspective. Can. J. Neurol. Sci. J. Can. Sci. Neurol. 39, 747–56 (2012).
4. Mountcastle, V. B. The columnar organization of the neocortex. Brain J. Neurol. 120 ( Pt 4), 701–722 (1997).
5. Nudo, R. J. & Masterton, R. B. Descending pathways to the spinal cord, III: Sites of origin of the corticospinal tract. J. Comp. Neurol. 296, 559–583 (1990).
6. Nudo, R. J. & Masterton, R. B. Descending pathways to the spinal cord, IV: Some factors related to the amount of cortex devoted to the corticospinal tract. J. Comp. Neurol. 296, 584–597 (1990).
7. Akintunde, A. & Buxton, D. F. Origins and collateralization of corticospinal, corticopontine, corticorubral and corticostriatal tracts: a multiple retrograde fluorescent tracing study. Brain Res. 586, 208–218 (1992).
8. Tennant, K. A. et al. The organization of the forelimb representation of the C57BL/6 mouse motor cortex as defined by intracortical microstimulation and cytoarchitecture. Cereb. Cortex N. Y. N 1991 21, 865–876 (2011).
9. Kamiyama, T. et al. Corticospinal tract development and spinal cord innervation differ between cervical and lumbar targets. J. Neurosci. Off. J. Soc. Neurosci. 35, 1181–1191 (2015). 10. Seo, J. P. & Jang, S. H. Different characteristics of the corticospinal tract according to the
cerebral origin: DTI study. AJNR Am. J. Neuroradiol. 34, 1359–1363 (2013).
11. Welniarz, Q., Dusart, I. & Roze, E. The corticospinal tract: Evolution, development, and human disorders. Dev. Neurobiol. 77, 810–829 (2017).
13. Tasic, B. et al. Adult Mouse Cortical Cell Taxonomy by Single Cell Transcriptomics. Nat. Neurosci. 19, 335–346 (2016).
14. Zeisel, A. et al. Brain structure. Cell types in the mouse cortex and hippocampus revealed by single-cell RNA-seq. Science 347, 1138–1142 (2015).
15. Sultan, K. T., Brown, K. N. & Shi, S.-H. Production and organization of neocortical interneurons. Front. Cell. Neurosci. 7, 221 (2013).
16. Sloviter, R. S. & Nilaver, G. Immunocytochemical localization of GABA-, cholecystokinin-, vasoactive intestinal polypeptide-, and somatostatin-like immunoreactivity in the area dentata and hippocampus of the rat. J. Comp. Neurol. 256, 42–60 (1987).
17. Cohen, I., Navarro, V., Clemenceau, S., Baulac, M. & Miles, R. On the origin of interictal activity in human temporal lobe epilepsy in vitro. Science 298, 1418–1421 (2002).
18. Katsarou, A.-M., Moshé, S. L. & Galanopoulou, A. S. INTERNEURONOPATHIES AND THEIR ROLE IN EARLY LIFE EPILEPSIES AND NEURODEVELOPMENTAL DISORDERS. Epilepsia Open 2, 284–306 (2017).
19. Lewis, D. A., Hashimoto, T. & Volk, D. W. Cortical inhibitory neurons and schizophrenia. Nat. Rev. Neurosci. 6, 312–324 (2005).
20. Rossignol, E. Genetics and function of neocortical GABAergic interneurons in neurodevelopmental disorders. Neural Plast. 2011, 649325 (2011).
21. Joosten, E. A., Schuitman, R. L., Vermelis, M. E. & Dederen, P. J. Postnatal development of the ipsilateral corticospinal component in rat spinal cord: a light and electron microscopic anterograde HRP study. J. Comp. Neurol. 326, 133–146 (1992).
22. Jang, S. H. The corticospinal tract from the viewpoint of brain rehabilitation. J. Rehabil. Med.
46, 193–199 (2014).
23. Vulliemoz, S., Raineteau, O. & Jabaudon, D. Reaching beyond the midline: why are human brains cross wired? Lancet Neurol. 4, 87–99 (2005).
24. Delwaide, P. J. & Oliver, E. Short-latency autogenic inhibition (IB inhibition) in human spasticity. J. Neurol. Neurosurg. Psychiatry 51, 1546–1550 (1988).
25. Tesson, C., Koht, J. & Stevanin, G. Delving into the complexity of hereditary spastic paraplegias: how unexpected phenotypes and inheritance modes are revolutionizing their nosology. Hum. Genet. 134, 511–538 (2015).
26. Oates, E. C. et al. Mutations in BICD2 cause dominant congenital spinal muscular atrophy and hereditary spastic paraplegia. Am. J. Hum. Genet. 92, 965–973 (2013).
27. Neveling, K. et al. Mutations in BICD2, which encodes a golgin and important motor adaptor, cause congenital autosomal-dominant spinal muscular atrophy. Am. J. Hum. Genet. 92, 946– 954 (2013).
28. Alazami, A. M., Adly, N., Al Dhalaan, H. & Alkuraya, F. S. A nullimorphic ERLIN2 mutation defines a complicated hereditary spastic paraplegia locus (SPG18). Neurogenetics 12, 333– 336 (2011).
29. Al-Saif, A., Bohlega, S. & Al-Mohanna, F. Loss of ERLIN2 function leads to juvenile primary lateral sclerosis. Ann. Neurol. 72, 510–516 (2012).
30. Parodi, L., Fenu, S., Stevanin, G. & Durr, A. Hereditary spastic paraplegia: More than an upper motor neuron disease. Rev. Neurol. (Paris) 173, 352–360 (2017).
31. Ruano, L., Melo, C., Silva, M. C. & Coutinho, P. The global epidemiology of hereditary ataxia and spastic paraplegia: a systematic review of prevalence studies. Neuroepidemiology 42, 174–183 (2014).
32. Erichsen, A. K., Koht, J., Stray-Pedersen, A., Abdelnoor, M. & Tallaksen, C. M. E. Prevalence of hereditary ataxia and spastic paraplegia in southeast Norway: a population-based study. Brain J. Neurol. 132, 1577–1588 (2009).
33. Boukhris, A. et al. Tunisian hereditary spastic paraplegias: clinical variability supported by genetic heterogeneity. Clin. Genet. 75, 527–536 (2009).
34. Blackstone, C. Hereditary spastic paraplegia. Handb. Clin. Neurol. 148, 633–652 (2018). 35. Deluca, G. C., Ebers, G. C. & Esiri, M. M. The extent of axonal loss in the long tracts in
hereditary spastic paraplegia. Neuropathol. Appl. Neurobiol. 30, 576–584 (2004).
36. Tesson, C., Koht, J. & Stevanin, G. Delving into the complexity of hereditary spastic paraplegias: how unexpected phenotypes and inheritance modes are revolutionizing their nosology. Hum. Genet. 134, 511–538 (2015).
37. Stevanin, G. et al. Mutations in SPG11, encoding spatacsin, are a major cause of spastic paraplegia with thin corpus callosum. Nat. Genet. 39, 366–372 (2007).
38. Stevanin, G. et al. Mutations in SPG11 are frequent in autosomal recessive spastic paraplegia with thin corpus callosum, cognitive decline and lower motor neuron degeneration. Brain J. Neurol. 131, 772–784 (2008).
39. Burke, D. & Ashby, P. Are spinal ‘presynaptic’ inhibitory mechanisms suppressed in spasticity? J. Neurol. Sci. 15, 321–326 (1972).
40. Rekling, J. C., Funk, G. D., Bayliss, D. A., Dong, X. W. & Feldman, J. L. Synaptic control of motoneuronal excitability. Physiol. Rev. 80, 767–852 (2000).
41. Heckman, C. J., Lee, R. H. & Brownstone, R. M. Hyperexcitable dendrites in motoneurons and their neuromodulatory control during motor behavior. Trends Neurosci. 26, 688–695 (2003).
42. Heckmann, C. J., Gorassini, M. A. & Bennett, D. J. Persistent inward currents in motoneuron dendrites: implications for motor output. Muscle Nerve 31, 135–156 (2005).
43. Burke, D., Knowles, L., Andrews, C. & Ashby, P. Spasticity, decerebrate rigidity and the clasp-knife phenomenon: an experimental study in the cat. Brain J. Neurol. 95, 31–48 (1972). 44. Nielsen, J., Petersen, N. & Crone, C. Changes in transmission across synapses of Ia afferents
in spastic patients. Brain J. Neurol. 118 ( Pt 4), 995–1004 (1995).
45. Nielsen, J. F., Klemar, B., Hansen, H. J. & Sinkjaer, T. A new treatment of spasticity with repetitive magnetic stimulation in multiple sclerosis. J. Neurol. Neurosurg. Psychiatry 58, 254–255 (1995).
46. Fries, W., Danek, A., Scheidtmann, K. & Hamburger, C. Motor recovery following capsular stroke. Role of descending pathways from multiple motor areas. Brain J. Neurol. 116 ( Pt 2), 369–382 (1993).
47. Synofzik, M. & Schüle, R. Overcoming the divide between ataxias and spastic paraplegias: Shared phenotypes, genes, and pathways. Mov. Disord. Off. J. Mov. Disord. Soc. 32, 332–345 (2017).
48. Blackstone, C. Converging cellular themes for the hereditary spastic paraplegias. Curr. Opin. Neurobiol. 51, 139–146 (2018).
49. Allison, R. et al. Defects in ER-endosome contacts impact lysosome function in hereditary spastic paraplegia. J. Cell Biol. 216, 1337–1355 (2017).
50. Lindsey, J. C. et al. Mutation analysis of the spastin gene (SPG4) in patients with hereditary spastic paraparesis. J. Med. Genet. 37, 759–765 (2000).
51. Evans, K. et al. Interaction of two hereditary spastic paraplegia gene products, spastin and atlastin, suggests a common pathway for axonal maintenance. Proc. Natl. Acad. Sci. U. S. A.
103, 10666–10671 (2006).
52. Mannan, A. U. et al. Spastin, the most commonly mutated protein in hereditary spastic paraplegia interacts with Reticulon 1 an endoplasmic reticulum protein. Neurogenetics 7, 93–103 (2006).
53. Sanderson, C. M. et al. Spastin and atlastin, two proteins mutated in autosomal-dominant hereditary spastic paraplegia, are binding partners. Hum. Mol. Genet. 15, 307–318 (2006). 54. Park, S. H., Zhu, P.-P., Parker, R. L. & Blackstone, C. Hereditary spastic paraplegia proteins
REEP1, spastin, and atlastin-1 coordinate microtubule interactions with the tubular ER network. J. Clin. Invest. 120, 1097–1110 (2010).
55. Durand, C. M. et al. CYP2U1 activity is altered by missense mutations in hereditary spastic paraplegia 56. Hum. Mutat. 39, 140–151 (2018).
56. Citterio, A. et al. Mutations in CYP2U1, DDHD2 and GBA2 genes are rare causes of complicated forms of hereditary spastic paraparesis. J. Neurol. 261, 373–381 (2014).
57. Tesson, C. et al. Alteration of fatty-acid-metabolizing enzymes affects mitochondrial form and function in hereditary spastic paraplegia. Am. J. Hum. Genet. 91, 1051–1064 (2012). 58. Cesnekova, J., Rodinova, M., Hansikova, H., Zeman, J. & Stiburek, L. Loss of Mitochondrial
AAA Proteases AFG3L2 and YME1L Impairs Mitochondrial Structure and Respiratory Chain Biogenesis. Int. J. Mol. Sci. 19, (2018).
59. Coarelli, G. et al. Loss of paraplegin drives spasticity rather than ataxia in a cohort of 241 patients with SPG7. Neurology 92, e2679–e2690 (2019).
60. Abel, A. et al. Early onset autosomal dominant spastic paraplegia caused by novel mutations in SPG3A. Neurogenetics 5, 239–243 (2004).
61. Zhu, P.-P., Soderblom, C., Tao-Cheng, J.-H., Stadler, J. & Blackstone, C. SPG3A protein atlastin-1 is enriched in growth cones and promotes axon elongation during neuronal development. Hum. Mol. Genet. 15, 1343–1353 (2006).
62. Zhu, P.-P., Denton, K. R., Pierson, T. M., Li, X.-J. & Blackstone, C. Pharmacologic rescue of axon growth defects in a human iPSC model of hereditary spastic paraplegia SPG3A. Hum. Mol. Genet. 23, 5638–5648 (2014).
63. de Bot, S. T. et al. Rapidly deteriorating course in Dutch hereditary spastic paraplegia type 11 patients. Eur. J. Hum. Genet. EJHG 21, 1312–1315 (2013).
64. Anheim, M. et al. SPG11 spastic paraplegia. A new cause of juvenile parkinsonism. J. Neurol.
256, 104–108 (2009).
65. Guidubaldi, A. et al. Novel mutations in SPG11 cause hereditary spastic paraplegia associated with early-onset levodopa-responsive Parkinsonism. Mov. Disord. Off. J. Mov. Disord. Soc. 26, 553–556 (2011).
66. Stevanin, G. et al. Mutations in SPG11 are frequent in autosomal recessive spastic paraplegia with thin corpus callosum, cognitive decline and lower motor neuron degeneration. Brain J. Neurol. 131, 772–784 (2008).
67. Al-Yahyaee, S. et al. A novel locus for hereditary spastic paraplegia with thin corpus callosum and epilepsy. Neurology 66, 1230–1234 (2006).
68. Karle, K. N. et al. Electrophysiological characterisation of motor and sensory tracts in patients with hereditary spastic paraplegia (HSP). Orphanet J. Rare Dis. 8, 158 (2013).
69. Orlacchio, A. et al. SPATACSIN mutations cause autosomal recessive juvenile amyotrophic lateral sclerosis. Brain J. Neurol. 133, 591–598 (2010).
70. Mukai, M. et al. Novel SPG11 Mutations in a Patient with Symptoms Mimicking Multiple Sclerosis. Intern. Med. Tokyo Jpn. 57, 3183–3186 (2018).
71. Kara, E. et al. Genetic and phenotypic characterization of complex hereditary spastic paraplegia. Brain J. Neurol. (2016) doi:10.1093/brain/aww111.
72. Faber, I. et al. SPG11-related parkinsonism: Clinical profile, molecular imaging and l-dopa response. Mov. Disord. Off. J. Mov. Disord. Soc. 33, 1650–1656 (2018).
73. Denora, P. S. et al. Motor neuron degeneration in spastic paraplegia 11 mimics amyotrophic lateral sclerosis lesions. Brain J. Neurol. 139, 1723–1734 (2016).
74. Pensato, V. et al. Overlapping phenotypes in complex spastic paraplegias SPG11, SPG15, SPG35 and SPG48. Brain J. Neurol. 137, 1907–1920 (2014).
75. Chang, J., Lee, S. & Blackstone, C. Spastic paraplegia proteins spastizin and spatacsin mediate autophagic lysosome reformation. J. Clin. Invest. 124, 5249–5262 (2014).
76. Varga, R.-E. et al. In Vivo Evidence for Lysosome Depletion and Impaired Autophagic Clearance in Hereditary Spastic Paraplegia Type SPG11. PLoS Genet. 11, e1005454 (2015). 77. Hanein, S. et al. Identification of the SPG15 Gene, Encoding Spastizin, as a Frequent Cause
of Complicated Autosomal-Recessive Spastic Paraplegia, Including Kjellin Syndrome. Am. J. Hum. Genet. 82, 992–1002 (2008).
78. Murmu, R. P. et al. Cellular distribution and subcellular localization of spatacsin and spastizin, two proteins involved in hereditary spastic paraplegia. Mol. Cell. Neurosci. 47, 191– 202 (2011).
79. Hirst, J. et al. Interaction between AP-5 and the hereditary spastic paraplegia proteins SPG11 and SPG15. Mol. Biol. Cell 24, 2558–2569 (2013).
80. Khundadze, M. et al. A hereditary spastic paraplegia mouse model supports a role of ZFYVE26/SPASTIZIN for the endolysosomal system. PLoS Genet. 9, e1003988 (2013).
81. Branchu, J. et al. Loss of spatacsin function alters lysosomal lipid clearance leading to upper and lower motor neuron degeneration. Neurobiol. Dis. 102, 21–37 (2017).
82. Boutry, M. et al. Inhibition of Lysosome Membrane Recycling Causes Accumulation of Gangliosides that Contribute to Neurodegeneration. Cell Rep. 23, 3813–3826 (2018). 83. Hirst, J. et al. Loss of AP-5 results in accumulation of aberrant endolysosomes: defining a
new type of lysosomal storage disease. Hum. Mol. Genet. 24, 4984–4996 (2015).
84. Renvoisé, B. et al. Lysosomal abnormalities in hereditary spastic paraplegia types SPG15 and SPG11. Ann. Clin. Transl. Neurol. 1, 379–389 (2014).
85. Kiernan, M. C. et al. Amyotrophic lateral sclerosis. Lancet Lond. Engl. 377, 942–955 (2011). 86. Vucic, S., Rothstein, J. D. & Kiernan, M. C. Advances in treating amyotrophic lateral sclerosis:
87. Al-Chalabi, A. et al. An estimate of amyotrophic lateral sclerosis heritability using twin data. J. Neurol. Neurosurg. Psychiatry 81, 1324–1326 (2010).
88. Keller, M. F. et al. Genome-wide analysis of the heritability of amyotrophic lateral sclerosis. JAMA Neurol. 71, 1123–1134 (2014).
89. Delestrée, N. et al. Adult spinal motoneurones are not hyperexcitable in a mouse model of inherited amyotrophic lateral sclerosis. J. Physiol. 592, 1687–1703 (2014).
90. Vucic, S., Cheah, B. C. & Kiernan, M. C. Defining the mechanisms that underlie cortical hyperexcitability in amyotrophic lateral sclerosis. Exp. Neurol. 220, 177–182 (2009).
91. Zhang, W. et al. Hyperactive somatostatin interneurons contribute to excitotoxicity in neurodegenerative disorders. Nat. Neurosci. 19, 557–559 (2016).
92. Ferraiuolo, L. et al. Oligodendrocytes contribute to motor neuron death in ALS via SOD1-dependent mechanism. Proc. Natl. Acad. Sci. U. S. A. 113, E6496–E6505 (2016).
93. Wainger, B. J. et al. Intrinsic Membrane Hyperexcitability of Amyotrophic Lateral Sclerosis Patient-Derived Motor Neurons. Cell Rep. 7, 1–11 (2014).
94. Majounie, E. et al. Frequency of the C9orf72 hexanucleotide repeat expansion in patients with amyotrophic lateral sclerosis and frontotemporal dementia: a cross-sectional study. Lancet Neurol. 11, 323–330 (2012).
95. Haeusler, A. R. et al. C9orf72 nucleotide repeat structures initiate molecular cascades of disease. Nature 507, 195–200 (2014).
96. Rosen, D. R. et al. Mutations in Cu/Zn superoxide dismutase gene are associated with familial amyotrophic lateral sclerosis. Nature 362, 59–62 (1993).
97. Ilieva, H., Polymenidou, M. & Cleveland, D. W. Non-cell autonomous toxicity in neurodegenerative disorders: ALS and beyond. J. Cell Biol. 187, 761–772 (2009).
98. Guo, W. et al. An ALS-associated mutation affecting TDP-43 enhances protein aggregation, fibril formation and neurotoxicity. Nat. Struct. Mol. Biol. 18, 822–830 (2011).
99. Heath, P. R. & Shaw, P. J. Update on the glutamatergic neurotransmitter system and the role of excitotoxicity in amyotrophic lateral sclerosis. Muscle Nerve 26, 438–458 (2002).
100. Rothstein, J. D., Van Kammen, M., Levey, A. I., Martin, L. J. & Kuncl, R. W. Selective loss of glial glutamate transporter GLT-1 in amyotrophic lateral sclerosis. Ann. Neurol. 38, 73–84 (1995).
101. Trotti, D., Rolfs, A., Danbolt, N. C., Brown, R. H. & Hediger, M. A. SOD1 mutants linked to amyotrophic lateral sclerosis selectively inactivate a glial glutamate transporter. Nat. Neurosci. 2, 427–433 (1999).
102. Rothstein, J. D. et al. Beta-lactam antibiotics offer neuroprotection by increasing glutamate transporter expression. Nature 433, 73–77 (2005).
103. Spalloni, A., Nutini, M. & Longone, P. Role of the N-methyl-d-aspartate receptors complex in amyotrophic lateral sclerosis. Biochim. Biophys. Acta 1832, 312–322 (2013).
104. Jaiswal, M. K. Selective vulnerability of motoneuron and perturbed mitochondrial calcium homeostasis in amyotrophic lateral sclerosis: implications for motoneurons specific calcium dysregulation. Mol. Cell. Ther. 2, 26 (2014).
105. Urushitani, M. et al. N-methyl-D-aspartate receptor-mediated mitochondrial Ca(2+) overload in acute excitotoxic motor neuron death: a mechanism distinct from chronic neurotoxicity after Ca(2+) influx. J. Neurosci. Res. 63, 377–387 (2001).
106. Sanelli, T., Ge, W., Leystra-Lantz, C. & Strong, M. J. Calcium mediated excitotoxicity in neurofilament aggregate-bearing neurons in vitro is NMDA receptor dependant. J. Neurol. Sci. 256, 39–51 (2007).
107. Van Damme, P., Braeken, D., Callewaert, G., Robberecht, W. & Van Den Bosch, L. GluR2 deficiency accelerates motor neuron degeneration in a mouse model of amyotrophic lateral sclerosis. J. Neuropathol. Exp. Neurol. 64, 605–612 (2005).
108. Kwak, S. & Kawahara, Y. Deficient RNA editing of GluR2 and neuronal death in amyotropic lateral sclerosis. J. Mol. Med. Berl. Ger. 83, 110–120 (2005).
109. Takuma, H., Kwak, S., Yoshizawa, T. & Kanazawa, I. Reduction of GluR2 RNA editing, a molecular change that increases calcium influx through AMPA receptors, selective in the spinal ventral gray of patients with amyotrophic lateral sclerosis. Ann. Neurol. 46, 806–815 (1999).
110. Saxena, S. et al. Neuroprotection through excitability and mTOR required in ALS motoneurons to delay disease and extend survival. Neuron 80, 80–96 (2013).
111. Leroy, F., Lamotte d’Incamps, B., Imhoff-Manuel, R. D. & Zytnicki, D. Early intrinsic hyperexcitability does not contribute to motoneuron degeneration in amyotrophic lateral sclerosis. eLife 3, (2014).
112. Wong, M. & Martin, L. J. Skeletal muscle-restricted expression of human SOD1 causes motor neuron degeneration in transgenic mice. Hum. Mol. Genet. 19, 2284–2302 (2010).
113. Bird, C. M. & Burgess, N. The hippocampus and memory: insights from spatial processing. Nat. Rev. Neurosci. 9, 182–194 (2008).
114. Hering, H. & Sheng, M. Dendritic spines: structure, dynamics and regulation. Nat. Rev. Neurosci. 2, 880–888 (2001).
115. Fiala, J. C., Feinberg, M., Popov, V. & Harris, K. M. Synaptogenesis via dendritic filopodia in developing hippocampal area CA1. J. Neurosci. Off. J. Soc. Neurosci. 18, 8900–8911 (1998). 116. Ziv, N. E. & Smith, S. J. Evidence for a role of dendritic filopodia in synaptogenesis and spine
formation. Neuron 17, 91–102 (1996).
117. Okamoto, K., Bosch, M. & Hayashi, Y. The roles of CaMKII and F-actin in the structural plasticity of dendritic spines: a potential molecular identity of a synaptic tag? Physiol. Bethesda Md 24, 357–366 (2009).
118. Androuin, A. et al. Evidence for altered dendritic spine compartmentalization in Alzheimer’s disease and functional effects in a mouse model. Acta Neuropathol. (Berl.) 135, 839–854 (2018).
119. Koch, G. et al. Impaired LTP- but not LTD-like cortical plasticity in Alzheimer’s disease patients. J. Alzheimers Dis. JAD 31, 593–599 (2012).
120. Pontrello, C. G. et al. Cofilin under control of β-arrestin-2 in NMDA-dependent dendritic spine plasticity, long-term depression (LTD), and learning. Proc. Natl. Acad. Sci. U. S. A. 109, E442–E451 (2012).
121. Fà, M., Staniszewski, A., Saeed, F., Francis, Y. I. & Arancio, O. Dynamin 1 is required for memory formation. PloS One 9, e91954 (2014).
122. Sheng, M., Sabatini, B. L. & Südhof, T. C. Synapses and Alzheimer’s disease. Cold Spring Harb. Perspect. Biol. 4, (2012).
123. Smith-Dijak, A. I. et al. Impairment and Restoration of Homeostatic Plasticity in Cultured Cortical Neurons From a Mouse Model of Huntington Disease. Front. Cell. Neurosci. 13, 209 (2019).
124. Smith-Dijak, A. I., Sepers, M. D. & Raymond, L. A. Alterations in synaptic function and plasticity in Huntington disease. J. Neurochem. 150, 346–365 (2019).
125. Walsh, D. M. et al. Naturally secreted oligomers of amyloid beta protein potently inhibit hippocampal long-term potentiation in vivo. Nature 416, 535–539 (2002).
126. Kamenetz, F. et al. APP processing and synaptic function. Neuron 37, 925–937 (2003). 127. Ungless, M. A., Whistler, J. L., Malenka, R. C. & Bonci, A. Single cocaine exposure in vivo
induces long-term potentiation in dopamine neurons. Nature 411, 583–587 (2001).
128. Mameli, M., Bellone, C., Brown, M. T. C. & Lüscher, C. Cocaine inverts rules for synaptic plasticity of glutamate transmission in the ventral tegmental area. Nat. Neurosci. 14, 414– 416 (2011).
129. Kolodziejczyk, K., Parsons, M. P., Southwell, A. L., Hayden, M. R. & Raymond, L. A. Striatal synaptic dysfunction and hippocampal plasticity deficits in the Hu97/18 mouse model of Huntington disease. PloS One 9, e94562 (2014).
130. Cummings, D. M. et al. Abnormal cortical synaptic plasticity in a mouse model of Huntington’s disease. Brain Res. Bull. 72, 103–107 (2007).
131. Begeti, F., Schwab, L. C., Mason, S. L. & Barker, R. A. Hippocampal dysfunction defines disease onset in Huntington’s disease. J. Neurol. Neurosurg. Psychiatry 87, 975–981 (2016). 132. Słabicki, M. et al. A genome-scale DNA repair RNAi screen identifies SPG48 as a novel gene
associated with hereditary spastic paraplegia. PLoS Biol. 8, e1000408 (2010).
133. Pérez-Brangulí, F. et al. Dysfunction of spatacsin leads to axonal pathology in SPG11-linked hereditary spastic paraplegia. Hum. Mol. Genet. 23, 4859–4874 (2014).
134. Duchesne, A. et al. Progressive ataxia of Charolais cattle highlights a role of KIF1C in sustainable myelination. PLoS Genet. 14, (2018).
135. Montecchiani, C. et al. ALS5/SPG11/KIAA1840 mutations cause autosomal recessive axonal Charcot-Marie-Tooth disease. Brain J. Neurol. 139, 73–85 (2016).
136. Sartucci, F., Tovani, S., Murri, L. & Sagliocco, L. Motor and somatosensory evoked potentials in Autosomal Dominant Hereditary Spastic Paraparesis (ADHSP) linked to chromosome 2p, SPG4. Brain Res. Bull. 74, 243–249 (2007).
137. Bönsch, D. et al. Motor system abnormalities in hereditary spastic paraparesis type 4 (SPG4) depend on the type of mutation in the spastin gene. J. Neurol. Neurosurg. Psychiatry 74, 1109–1112 (2003).
138. Vitner, E. B., Platt, F. M. & Futerman, A. H. Common and uncommon pathogenic cascades in lysosomal storage diseases. J. Biol. Chem. 285, 20423–20427 (2010).
139. Lawson, C. A. & Martin, D. R. Animal models of GM2 gangliosidosis: utility and limitations. Appl. Clin. Genet. 9, 111–120 (2016).
140. Vivas, O., Tiscione, S. A., Dixon, R. E., Ory, D. S. & Dickson, E. J. Niemann-Pick Type C Disease Reveals a Link between Lysosomal Cholesterol and PtdIns(4,5)P2 That Regulates Neuronal Excitability. Cell Rep. 27, 2636–2648.e4 (2019).
141. Avchalumov, Y. et al. Increased excitability and compromised long-term potentiation in the neocortex of NPC1(-/-) mice. Brain Res. 1444, 20–26 (2012).
142. Eckhardt, M. et al. Sulfatide Storage in Neurons Causes Hyperexcitability and Axonal Degeneration in a Mouse Model of Metachromatic Leukodystrophy. J. Neurosci. 27, 9009– 9021 (2007).
143. Ortu, E. et al. Primary motor cortex hyperexcitability in Fabry’s disease. Clin. Neurophysiol. Off. J. Int. Fed. Clin. Neurophysiol. 124, 1381–1389 (2013).
144. Hassan, S. S., Trenado, C., Elben, S., Schnitzler, A. & Groiss, S. J. Alteration of cortical excitability and its modulation by Miglustat in Niemann-Pick disease type C. J. Clin. Neurosci. Off. J. Neurosurg. Soc. Australas. 47, 214–217 (2018).
145. Marcuzzo, S. et al. Hyperexcitability in Cultured Cortical Neuron Networks from the G93A-SOD1 Amyotrophic Lateral Sclerosis Model Mouse and its Molecular Correlates. Neuroscience 416, 88–99 (2019).
146. Clark, R. M., Brizuela, M., Blizzard, C. A. & Dickson, T. C. Reduced Excitability and Increased Neurite Complexity of Cortical Interneurons in a Familial Mouse Model of Amyotrophic Lateral Sclerosis. Front. Cell. Neurosci. 12, 328 (2018).
147. Vollrath, J. T. et al. Loss of function of the ALS protein SigR1 leads to ER pathology associated