However, C9orf72-deficient mice do not display overt neurodegenerative phenotypes 14,18,19,22. Moreover, no studies have shown that reduced C9ORF72 activity leads to the degeneration of C9ORF72 ALS patient-derived motor neurons, nor have any provided direct evidence identifying a cellular pathway through which C9ORF72 activity modulates neuronal survival. Additionally, a patient homozygous for the C9ORF72 repeat expansion had clinical and pathological phenotypes that were severe but nonetheless did not fall outside the range of heterozygous patients, leaving it uncertain if reductions in C9ORF72 protein levels directly correlate with disease severity 23. Thus, the role of the C9ORF72 protein in C9ORF72 ALS/FTD disease pathogenesis remains unclear.
IPSC-MNs at differentiation D35 were harvested in cold Hypotonic buffer (20 mM HEPES pH 7.4, 10 mM KCl, 2 mM MgCl2, 1 mM EDTA, 1mM EGTA, 1 mM DTT and protease inhibitor cocktail (Roche)) and lysed by passing through G25 needles 25 times and then spun down at 700 x g for 10min at 4℃. The Supernatant was loaded onto pre-made 30% Percoll solution and re-centrifuged at 33,000 RPM using Beckman rotor SWI55 for 50min at 4℃. 300 ul aliquots were taken from top to bottom as fractions and all the collected samples were boiled with SDS-PAGE sample buffer and analyzed by western blot.
Hb9::RFP+ iMNs appeared between days 13–16 after retroviral transduction. RepSox was removed at day 17 and the survival assay was initiated. For the glutamate treatment condition, 10 µM glutamate was added to the culture medium on day 17 and removed after 12 hours. Cells were then maintained in N3 medium with neurotrophic factors without RepSox. For the glutamate treatment condition with glutamate receptor antagonists, cultures were co-treated with 10 μM MK801 and CNQX, and 2 μM Nimodipine during the 12 hour glutamate treatment. The antagonists were maintained for the remainder of the experiment. For the neurotrophic factor withdrawal condition, BDNF, GDNF, and CNTF were removed from the culture medium starting at day 17. Longitudinal tracking was performed by imaging neuronal cultures in a Nikon Biostation CT or Molecular Devices ImageExpress once every 24–72 hours starting at day 17. Tracking of neuronal survival was performed using SVcell 3.0 (DRVision Technologies). Neurons were scored as dead when their soma was no longer detectable by RFP fluorescence. All neuron survival assays were performed at least twice, with equal numbers of neurons from three individual replicates from one of the trials being used for the quantification shown. All trials quantified were representative of other trials of the same experiment. When iMNs from multiple independent donors are combined into one survival trace in the Kaplan-Meier plots for clarity, the number of iMNs tracked from each line can be found in Supplementary Table 5.
The GGGGCC repeat expansion in C9ORF72 is the most common cause of amyotrophic lateral sclerosis (ALS) and frontotemporal dementia (FTD), accounting for about 10% of each disease worldwide 1–4. In the central nervous system (CNS), neurons and microglia express the highest levels of C9ORF72 5, suggesting that C9ORF72 acts in part cell autonomously and effects in neurons are a key source of disease etiology. Studies showing that the repeat expansion generates neurotoxic species including nuclear RNA foci 6–8, RNA/DNA G-quadruplexes 9, and dipeptide repeat proteins (DPRs) 10–12 have oriented the field towards a therapeutic focus on blocking the toxicity of these products 6–8,13,14. However, these strategies have not fully rescued the degeneration of patient-derived neurons 7,13. Moreover, tandem GGGGCC repeats are transcribed from over 80 other genomic locations within human spinal motor neurons (Supplementary Tables 1 and 2), yet genetic studies have not linked repeat expansions in these regions to ALS/FTD. In addition, hexanucleotide repeat-mediated toxicity in mice requires supraphysiological expression levels or a specific genetic background 14–16. These observations suggest that there are additional pathogenic triggers caused by repeat expansion within C9ORF72.

Shi Y1,2,3, Lin S1,2,3, Staats KA1,2,3, Li Y1,2,3, Chang WH1,2,3, Hung ST1,2,3, Hendricks E1,2,3, Linares GR1,2,3, Wang Y3,4, Son EY5, Wen X6, Kisler K3,4, Wilkinson B3, Menendez L1,2,3, Sugawara T1,2,3, Woolwine P1,2,3, Huang M1,2,3, Cowan MJ1,2,3, Ge B1,2,3, Koutsodendris N1,2,3, Sandor KP1,2,3, Komberg J1,2,3, Vangoor VR7, Senthilkumar K7, Hennes V1,2,3, Seah C1,2,3, Nelson AR3,4, Cheng TY8, Lee SJ8, August PR9, Chen JA10, Wisniewski N10, Hanson-Smith V10, Belgard TG10, Zhang A10, Coba M3,11, Grunseich C12, Ward ME12, van den Berg LH13, Pasterkamp RJ7, Trotti D6, Zlokovic BV3,4, Ichida JK1,2,3.


Our results indicate that haploinsufficiency for C9ORF72 activity triggers neurodegeneration in C9ORF72 ALS, and this occurs by at least two mechanisms. First, reduced C9ORF72 activity causes the accumulation of glutamate receptors and excitotoxicity in response to glutamate. Although C9orf72 knockout mice do not display overt neurodegeneration14,18,22, these mice may be protected from excitotoxicity because they lack gain-of-function disease processes such as DPRs, which induce aberrant splicing and dysfunction of the EAAT2 glutamate transporter in astrocytes in vitro 12 and in C9ORF72 ALS patients 4,27. EAAT2 dysfunction causes glutamate accumulation in the cerebrospinal fluid of ALS patients 27, and consistent with this notion, we found that poly(PR) expression in human astrocytes reduced their rate of glutamate uptake. By using human iMNs, mice, and human post mortem tissue, we show for the first time that reduced C9ORF72 activity modulates the vulnerability of human motor neurons to degenerative stimuli and establish a mechanistic link between the C9ORF72 repeat expansion and glutamate-induced excitotoxicity
To study the pathogenic mechanism of the C9ORF72 repeat expansion in human motor neurons, we used the forced expression of the transcription factors Ngn2, Isl1, Lhx3, NeuroD1, Brn2, Ascl1, and Myt1l, to convert control and C9ORF72 ALS/FTD patient induced pluripotent stem cells (iPSCs)(for iPSC characterization, see Supplementary Fig. 1 and Supplementary Tables 3, 4) into iMNs (Supplementary Fig. 2a, b) 10,24. Control and patient iMNs labeled with an Hb9::RFP+ lentiviral reporter construct (Supplementary Fig. 2b-d)25 co-expressed spinal motor neuron markers including TUJ1, HB9, and VACHT; were produced at similar rates amongst different iPSC lines; and possessed electrophysiological properties of motor neurons (Supplementary Fig. 2c-i). Depolarizing voltage steps induced currents characteristic of sodium and potassium channels and iMNs fired single or repetitive action potentials (patient - 90%, n=10; control – 100%, n=10)(Supplementary Fig. 2g-i). When co-cultured with primary chick muscle, channel rhodopsin-expressing control and patient iMNs repeatedly induced myotube contraction upon depolarization with green light, indicating they formed neuromuscular junctions and actuated muscle contraction (Supplementary Fig. 2j and Supplementary Videos 1, 2).
Complementary DNAs (cDNAs) for the iMN factors (Ngn2, Lhx3, Isl1, NeuroD1, Ascl1, Myt1l, and Brn2) and iDA neuron factors (Ascl1, Brn2, Myt1l, Lmx1a, and Foxa2), were purchased from Addgene. cDNA for C9ORF72 was purchased from Thermo Scientific. Each cDNA was cloned into the pMXs retroviral expression vector using Gateway cloning technology (Invitrogen). The Hb9::RFP lentiviral vector was also purchased from Addgene (ID: 37081). Viruses were produced as follows. HEK293 cells were transfected at 80–90% confluency with viral vectors containing genes of interest and viral packaging plasmids (PIK-MLV-gp and pHDM for retrovirus; pPAX2 and VSVG for lentivirus) using polyethylenimine (PEI)(Sigma-Aldrich). The medium was changed 24h after transfection. Viruses were harvested at 48h and 72 h after transfection. Viral supernatants were filtered with 0.45 µM filters, incubated with Lenti-X concentrator (Clontech) for 24 h at 4 ºC, and centrifuged at 1,500 x g at 4ºC for 45 min. The pellets were resuspended in 300 µl DMEM + 10% FBS and stored at −80 ºC.

To measure the effect of dipeptide repeat protein expression on iMN survival, PR50 and GR50 were cloned into the pHAGE lentiviral vector as fusions with GFP to allow tracking of protein expression. iMN cultures were transduced with PR50 and GR50 lentiviruses at day 17 of reprogramming and longitudinal survival analysis was started the same day. 10 ng/ml of GDNF, BDNF, and CNTF was maintained throughout the experiment, and glutamate treatment was not performed. To measure PR50 turnover, PR50 was cloned into the pHAGE lentiviral vector as a fusion with Dendra2 (Addgene). iPSC-derived fibroblasts were generated according to Daley and colleagues64. Briefly, when C9ORF72−/− iPSC cultures reached 80% confluence, the medium was switched from mTeSR1 (Stem Cell Technologies) to human fibroblast medium containing DMEM (Life Technologies), 10% fetal bovine serum (FBS)(Thermo Fisher Scientific), and 1% penicillin/streptomycin (Life Technologies). Cells were passaged 2 to 3 times using Accutase (Life Technologies) before use in experiments. iPSC-derived fibroblasts were transduced with either pMXs-eGFP or pMXs-C9ORF72 isoform B-T2A-eGFP retrovirus and treated with 10 μg/ml mitomycin C for 3 hrs to inhibit cell proliferation. The cells were then transduced with the PR50–Dendra2 lentivirus and exposed to blue light for 1.5 sec using a lumencor LED light source to initiate photoconversion. The amount of decay (as a fraction of the starting level) of the red fluorescent punctae was monitored by longitudinal time lapse imaging in a Molecular Devices ImageExpress and analyzed using SVCell 2.0 (DRVision Technologies). Fluorescence was quantified at t = 0 and 12 hours after photoconversion. Distinct photoconverted punctae were treated as discrete objects for analysis (n = 20 each for +eGFP and +C9ORF72-T2A-eGFP). For each object, background fluorescence was subtracted and fluorescence was normalized according to object size. The fractional decay was statistically analyzed by two-tailed Student’s t-test. ** - p<.01.

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