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Neuropathy-associated Fars2 deficiency affects neuronal development and potentiates neuronal apoptosis by impairing mitochondrial function
Cell & Bioscience volume 12, Article number: 103 (2022)
Neurodegenerative diseases encompass an extensive and heterogeneous group of nervous system disorders which are characterized by progressive degeneration and death of neurons. Many lines of evidence suggest the participation of mitochondria dysfunction in these diseases. Mitochondrial phenylalanyl-tRNA synthetase, encoded by FARS2, catalyzes the transfer of phenylalanine to its cognate tRNA for protein synthesis. As a member of mt-aaRSs genes, FARS2 missense homozygous mutation c.424G > T (p.D142Y) found in a Chinese consanguineous family first built the relationship between pure hereditary spastic paraplegia (HSP) and FARS2 gene. More FARS2 variations were subsequently found to cause heterogeneous group of neurologic disorders presenting three main phenotypic manifestations: infantile-onset epileptic mitochondrial encephalopathy, later-onset spastic paraplegia and juvenile onset refractory epilepsy. Studies showed that aminoacylation activity is frequently disrupt in cases with FARS2 mutations, indicating a loss-of-function mechanism. However, the underlying pathogenesis of neuropathy-associated Fars2 deficiency is still largely unknown.
Early gestation lethality of global Fars2 knockout mice was observed prior to neurogenesis. The conditional Fars2 knockout-mouse model delayed lethality to late-gestation, resulting in a thinner cortex and an enlarged ventricle which is consist with the MRI results revealing cortical atrophy and reduced cerebral white matter volume in FARS2-deficient patients. Delayed development of neurite outgrowth followed by neuronal apoptosis was confirmed in Fars2-knockdown mouse primary cultured neurons. Zebrafish, in which fars2 was knocked down, exhibited aberrant motor neuron function including reduced locomotor capacity which well restored the spastic paraplegia phenotype of FARS2-deficient patients. Altered mitochondrial protein synthesis and reduced levels of oxidative phosphorylation complexes were detected in Fars2-deficient samples. And thus, reduced ATP, total NAD levels and mitochondrial membrane potential, together with increased ROS production, revealed mitochondrial dysfunction both in vitro and in vivo. Dctn3 is a potential downstream molecule in responds to Fars2 deficient in neurons, which may provide some evidence for the development of pathogenesis study and therapeutic schedule.
The Fars2 deficiency genetic models developed in this study cover the typical clinical manifestations in FARS2 patients, and help clarify how neuropathy-associated Fars2 deficiency, by damaging the mitochondrial respiratory chain and impairing mitochondrial function, affects neuronal development and potentiates neuronal cell apoptosis.
Mitochondrial aminoacyl-tRNA synthetases (mt-aaRSs) are a family of enzymes that are encoded by the nuclear genome, translated by cytosolic ribosomes, and imported into the mitochondria where they charge specific tRNAs for protein synthesis . Thirteen mitochondrial DNA (mt-DNA) encoded proteins are synthesized via the mitochondrial translation machinery. These proteins are subunits of respiratory chain complexes that play an important role in oxidative phosphorylation (OXPHOS), a crucial determinant of mitochondrial function . Mutations in mt-aaRSs genes including DARS2 , EARS2 , FARS2 , MARS2  and RARS2  can lead to central nervous system (CNS) disorders and/or muscle-related pathologies [8, 9]. Infantile-onset epileptic encephalopathy, later-onset spastic paraplegia and recently identified juvenile onset refractory epilepsy are three main phenotypes exhibited by patients harboring FARS2 mutations . To date, 39 pathogenic FARS2 variants have been implicated across various disease types. We have previously shown that a missense homozygous mutation c.424G > T (p.D142Y) in the FARS2 gene is the cause of pure-form hereditary spastic paraplegia (HSP) .
FARS2 encodes mitochondrial phenylalanyl-tRNA synthetase (mtPheRS) that contains a class II catalytic domain and an anti-codon binding domain  that drive the aminoacylation reaction during protein synthesis in the mitochondria . Patients with FARS2 mutations who received enzymatic tests all showed reduced or even absent aminoacylation activity . These findings suggest that mtPheRS loss-of-function variants that cause mitochondrial dysfunction may promote pathogenesis. Due to the lack of an animal model and the unobtainable of human samples, however, the relationship between FARS2 deficiency and nervous system development and maintenance is unknown; research into FARS2 deficiency therefore remains in its infancy. Functional studies of FARS2 in Saccharomyces cerevisiae, have provided invaluable insights into studying Fars2 function. Nevertheless, when considering phenotypic heterogeneity of human disease or translational relevance at the level of organs or tissues, it is clear that a single-cell microorganism is not an ideal model system in which to study human neurodegenerative diseases .
Therefore, to mimic the pathophysiology of FARS2 deficiency, we established mutant p.D142Y and global Fars2-knockout-mouse models. Embryos genotyped as homozygous Fars2-knockout or expressing the p.D142Y mutation, stop developing during early gestation, just before the neural plate is formed. Furthermore, conditional Fars2-deletion in mouse embryos at embryonic day (E) 11 causes significant neural cell apoptosis and a lethal phenotype at late-gestation. Thinner cortex and enlarged ventricle were observed which is consist with the MRI results revealing cortical atrophy and reduced cerebral white matter volume in human patients. In vitro Fars2-knockdown mouse neurons showed delayed and disrupt neurite outgrowth. Both in vivo and in vitro experiments revealed disrupted mitochondrial protein synthesis, impaired OXPHOS biogenesis, and injured mitochondria. The further generated fars2-knockdown zebrafish models showed impaired motor axon growth and reduced locomotor capacity, features believed to be consistent with the manifestation of HSP patients.
To our knowledge, the present study is the first to demonstrate a relationship between FARS2 deficiency and brain function using cell culture, and animal (mice and zebrafish) models. These models help elucidate the pathophysiology that underlies FARS2-related neurodegenerative diseases.
Review of FARS2 variations and human disease
The human FARS2 gene consists of seven exons, of which exons 2–7 are responsible for coding mtPheRS. As the smallest aaRS, mtPheRS has only two structural domains: class II catalytic domain and anti-codon binding domain . Here, we summarize all the variations in FARS2 that have been reported to cause human disease to date (Table 1). These mutations are spread throughout the whole peptide of mtPheRS and include 30 missense mutations, 2 nonsense mutations, 6 inframe or outframe deletions and 1 splice-site mutation (Fig. 1A). Patients inherit these mutations in an autosomal recessive manner: compound heterozygous for two mutations or homozygous for one mutation. Conservation analysis across species showed that most of the missense mutations occur at highly conserved sites, with a ConSurf score of no less than 6. While some mutations occurred at less conserved sites, such as p.R223, p.D325, p.D364 and p.L371, can still be related to disease (Fig. 1B).
FARS2 deficiency comprises a spectrum of disease severities. The phenotypes can be divided into three types according to the age of onset, namely infantile-onset epileptic encephalopathy, later-onset spastic paraplegia, and juvenile onset refractory epilepsy . We observed that in addition to neuronal-muscle manifestations, global or neurological developmental delay is the most common phenotype of these patients. Mutants on which the aminoacylation activity measurement was carried out all showed varying degrees of disruption, the p.D325Y mutant relating to the infantile-onset epileptic encephalopathy phenotype and the p.D364G mutant relating to the later-onset spastic paraplegia even showed absolutely disappeared aminoacylation activity [12, 22]. Furthermore, in our previous study, the aminoacylation activity of HSP-associated FARS2 p.D142Y mutant was confirmed to be almost completely destroyed by purifying the recombinant human mtPheRS proteins in vitro , suggesting that loss of canonical FARS2 function could be a fundamental common mechanism of FARS2-related neuropathy in human. OXPHOS deficiency is another common phenotype of FARS2 mutant patients. However, OXPHO complex activity differs between patients, and even between cell types of the same patient. Decreased complex I (CI) and CIV activities were mostly detected, while increased CII and CIV levels were detected in a patient with p.V197M and an exon 2 microdeletion. The variabilities in clinical presentation and biochemistry results suggest that the effect of FARS2 deficiency can various among systems. Therefore, studying the effect of FARS2 deficiency on CNS specifically is a requisite for explaining the neuropathological mechanism of FARS2 loss of function disease.
Fars2 is expressed early during mouse embryonic development and expressed widely in multiple organs
Fars2 encodes mtPheRS which transfers phenylalanine to its cognate tRNA in mitochondria for protein translation, suggesting that it is required for nearly all eukaryotic cells and stages of eukaryotic development [30, 31]. To investigate the expression pattern of Fars2 in mouse embryos throughout their development, we used time-mating to obtain embryos and/or embryonic CNS samples at seven different developmental stages. Total mRNA was extracted and Fars2 expression levels were examined with quantitative RT-PCR (qRT-PCR). Fars2 was expressed as early as E 7.5. The level of Fars2 mRNA expression in the embryo and/or embryonic CNS remained stable until E 14, and then increased dramatically in the brain of E 17 embryos (Additional file 1: Fig.S1A). Protein lysates were collected from various organs from E 17 embryos and western blotting was used to determine the level of mtPheRS expression. MtPheRS is expressed and is particularly high in heart and liver, followed by kidney and the CNS (Additional file 1: Fig. S1B and C). The above results show that Fars2 is expressed in mouse embryos at a very early stage, and is widely expressed in multiple organs in late embryonic mice.
Sufficient Fars2 function is essential for embryonic neurogenesis in mice
Our previous work confirmed that the p.D142Y mutation disrupts the aminoacylation activity of mtPheRS, leaving only a trace of enzymatic activity. To learn more about the HSP mechanism caused by this missense mutation and the physiological roles of Fars2, we used the CRISPR/Cas 9 system to create heterozygous global mutant and global knockout (KO) Fars2 mice (Additional file 1: Fig. S2A). The distribution of Fars2 genotypes in the progeny of Fars2 null/+ and Fars2 c.424G>T/+ intercrossing was counted. Genotyping confirmed that the ratio of heterozygote to wildtype was 2:1 in both mutant and KO strains, while no homozygous mutant or KO mice were born (Table 2; Fig. 2A and B). Atrophic embryos were discovered in the uterus of pregnant mice at E 14.5 subsequently, indicating embryonic lethality (Additional file 1: Fig. S2B). We used time-mating to determine the exact developmental stage of this lethal effect. During E 7.0-7.5, no obvious differences in morphology were observed between littermates, whereas abnormal appearances were observed among E 7.5-8.0 embryos derived from Fars2 null/+ and Fars2 c.424G>T/+ intercrossing (Fig. 2C, Additional file 1: Fig. S2C). Hematoxylin and eosin staining of E 7.5-8.0 embryos revealed that embryos with normal appearances had developed posterior amniotic fold structures, characteristic of embryos at the late streak stage; embryo with abnormal appearances, however, had stopped at the early streak stage and the extraembryonic portion of the egg cylinder had begun to disintegrate (Fig. 2D). Genotyping the normal and abnormal embryos confirmed them to be wildtype and homozygous Fars2-deficient strains (Fig. 2E). We collected four groups of littermates extracted from Fars2 c.424G>T/+ intercrossing at E 7.0-7.5 and eight litters (four normal, and four with abnormal morphology) extracted at E 7.5-8.0 to confirm the stage at which Fars2-deficient embryos stop developing. Total mRNA was extracted from the embryos without any maternal components. Nepn  and Mesp2  were chosen as markers of the definitive endoderm and mesoderm, and were reported to express at E 7.5 and E 7.0 respectively. In addition, Pax6 starts to express in the neural stem/progenitor cells of ectoderm at E 8.0, the earliest stage of CNS development, and marks embryonic neurogenesis . As a result, at both E 7.0-7.5 and E 7.5-8.0, all of the littermates express Mesp2 and Nepn. Pax6 expression was found in embryos with normal morphology at E 7.5-8.0, but not in embryos with a smaller appearance. These findings suggest that embryos with a homozygous Fars2 KO or mutant genotype can develop definitive endodermal and mesodermal layers, but not the ectoderm (Fig. 2F and G). The above results indicate that Fars2 function is required for embryonic neurogenesis, and the p.D142Y is a loss of function mutant which produces a KO-like phenotype in mice.
Fars2 deficiency in the CNS results in a thinner cortex and an enlarged ventricle due to progressive cell apoptosis
To determine if Fars2 deletion affects brain development, we used CRISPR/Cas9 to create conditional-knockout Fars2 mice (Fars2 fl/fl). By mating Fars2 fl/fl mice with Nestin-Cre mice, which specifically express Cre recombinase in nervous tissue by E 11.0. Thus, neuron-specific Fars2 knockout (cKO) mouse with the genotype of Nes-Cre+; Fars2 fl/fl were generated (Additional file 1: Fig. S3A and B). Western blotting results of FARS2 in brain and liver confirmed the knockout efficiency and tissue specificity of Cre (Additional file 1: Fig. S3C). We discovered that cKO pups were born with a slight purple color and died shortly after birth (Fig. 3A). Gross examination of their cerebral cortex revealed red lesions. The total brain volume did not significantly differ between the cKO and control groups, but the weight of the brain was significantly reduced in the cKO mice (Fig. 3B-D). Frozen brain slices were cut in coronal sections to evaluate structural changes of Fars2-deficient embryos. At E 17.5, Nissl staining of the cortex, with and without the cranium, revealed an enlarged ventricle and reduced cortical thickness (Fig. 3E and F). Nissl staining and immunofluorescence staining for TUJ1 at E15.5 revealed that the thickness of all formed layers in the cortex were affected (Fig. 3G and H).
The TUNEL assay was then used to detect cells undergoing apoptosis. Several positive dots were scattered thinly in the dorsal cortex and ventral cortex of E 14.5 cKO mice; at E 17.5, however, the positive dots were distributed throughout these two regions (Fig. 3I). We then collected protein lysates from E 14.5 and E 17.5 embryonic cerebral cortex, and used Western blotting to determine Caspase3 and Cleaved-caspase 3 levels. Cleaved-caspase fragments of cKO were accumulated gradually (Fig. 3 J). These data suggest that the high rate of cellular apoptosis in Fars2-deficient mice may produce a thinner cortex, and that this process is a progressive one.
Fars2 deficiency affects neurite outgrowth and causes neuron apoptosis
To investigate the role of Fars2 in long-term neuron development and maturation, we infected primary cultured neurons with mcherry-shFars2 lentivirus and obtained neurons in which Fars2 expression was efficiently knocked down (Additional file 1: Fig. S4A and B). Immunocytochemistry for MAP2 facilitated the tracking of neurite morphogenesis. The length of the longest primary neurite of neurons with Fars2 knocked-down, was relatively normal following four days of culture after infection; but with a slighter figure and fewer secondary branches than in the control group. While on the eighth day, the mean length of the longest primary neurite was significantly shorter than that of control neurons, but similar to control neurons on the fourth day, indicating a development delay (Fig. 4A and B). Sholl’s analysis of concentric circles revealed that neurite arborization was significantly lower in neurons with Fars2 knocked-down than in controls (Fig. 4C and D). Images taken with a low-power lens on the eighth day showed that connections between Fars2-deficient neurons were severely disrupted due to axon degeneration (Fig. 4E). In addition to morphological changes, we discovered that after long-term in vitro cultivation, very few neurons in the Fars2-knockdown group survived. TUNEL assays and Western blotting were used to detect apoptosis at 8 and 12 days after infection, respectively. An increased number of cell death was detected in Fars2 knock-down group compared to the control group (Fig. 4F). And the level of Cleaved-caspase 3/Caspase 3 was significantly higher in Fars2 knock-down neurons (Fig. 4G).
The above findings suggest that the outgrowth of neurites was severely affected by Fars2 deficiency, and progressive neuron apoptosis was thus triggered. As a result of delayed neurite development and devastating early apoptosis, the structural connection between Fars2-deficient neurons was disrupt.
Mitochondrial protein biosynthesis is disrupted in Fars2 -deficient organisms
The mitochondrial translation machinery that mtPheRS participate in includes the synthesis of thirteen mitochondrial respiratory chain complex subunits. The proportions of phenylalanine among the five mitochondrial complexes from humans, rats, mice, and zebrafish, are shown in Additional file 1: Table S5. With the exception of CII, which lacks mitochondrial encoded subunits, phenylalanine is present in all the other four complexes. To better understand the effect of Fars2 deficiency on protein translation, we looked at the total levels of five fully assembled respiratory chain complexes in cortex and primary cultured neurons. In E 14.5 embryos from cKO mice, Western blotting revealed significantly reduced levels of CI and CIV, as well as slightly reduced levels of CIII. At E 17.5, these three complexes were further reduced. While CII and CV levels were slightly higher in Fars2-deficient embryos (Fig. 5A-C). To determine whether the reduced levels were due to reduced translation of the relevant subunits, we examined the levels of the mitochondrial proteins ND1, CO2, and CYTB, which belong to the three affected complexes. Western blotting data showed consistent variation of ND1, CO2 and CYTB levels with their corresponding complexes, indicating that disruption in subunits translation is responsible for deficient mitochondrial respiratory chain complexes (Fig. 5D-F). Changes within cultured neurons after Fars2 knockdown, however, followed different patterns. CI was almost disappeared in Fars2 knockdown neurons after 16 days in culture, whereas CII and CV were slightly increased following 4 and 16 days of infection respectively (Fig. 5G and H); and the ND1 level showed significantly reduction in Fars2 knockdown neurons at 16th day but can still be synthesized, which indicate that fars2 knocking down affect both ND1 translation process and the assembly of mitochondrial CI in neurons (Fig. 5I and J). We also measured the number of mitochondria by calculating the amount of TOMM20 by Western blotting (Fig. 5A, B and G). The Fars2-deficient mouse cortical and cultured neurons both showed unchanged mitochondrial mass which indicated that the down-regulated complex levels cannot be attributed to the number change of mitochondria (Fig. 5K and L). Furthermore, in Fars2-deficient tissue and cells, transcriptional levels of mitochondrial encoded respiratory chain complex subunits were generally upregulated (Fig. 5M and N), which further demonstrates that the decreased complex levels in Fars2-deficient samples occurs at the post-transcriptional level.
Fars2 deficiency leads to impaired OXPHOS biogenesis and damaged mitochondria
The mitochondrial electron transfer chains are responsible for the production of almost 80% of cellular energy in the form of adenosine triphosphate (ATP), and also help regulate reactive oxygen species (ROS) . We assessed the effects of reduced mitochondrial respiratory chain complexes on ATP and ROS in E 17.5 cKO cortex embryo, primary cultured neurons, and the PC12 cell line. Our findings show that Fars2-deficient tissue and cell lines have lower ATP concentrations and higher ROS levels, implying that mitochondrial electron transfer is dysfunctional (Fig. 6A and B; Additional file 1: Fig. S5A and B). ATP concentration in cultured neurons, however, did not change over the course of 12 days (Fig. 6C). While ROS levels were slightly lower than in controls during the first four days of Fars2 knockdown, they began to rise steadily after the eighth day (Fig. 6D).
Increased ROS production often accompanies mitochondrial dysfunction, and reduction of the mitochondrial membrane potential (Δψm) is an associated hallmark [36, 37]. We monitored Δψm in the Fars2-deficient cortex and PC12 cells with JC-1 staining. Increased green fluorescence and decreased red fluorescence were detected, suggesting that mitochondria depolarization reduced Δψm (Fig. 6E; Additional file 1: Fig. S5C).
Nicotinamide adenine dinucleotide (NAD) participates in a variety of biological processes, including mitochondrial functions, oxidative stress generation, cell apoptosis, and axonal degeneration . And the NAD+/NADH dinucleotide pair drives a wide range of reduction–oxidation reactions in cells. According to our results, the total pool of NAD level was significantly lower in the Fars2-deficient cortex and in cultured neurons, and the reduction is largely due to the low NAD in the mitochondria pool, but not the cytosolic pool (Fig. 6F and H). And the reduction of total NAD is largely due to the reduced NAD+ level in the mitochondria pool in Fars2-deficient mouse cortex (Fig. 6G). Interestingly, the NAD+/NADH ratio of cultured neurons undergo Fars2 knockdown decreased at the 4th and 8th day of culture but seemed to be slightly rescued between the two time points (Fig. 6I).
We further used a Seahorse XF24 extracellular flux analyzer to measure mitochondrial aerobic respiration. Compared with the control group, maximal respiration and ATP production were both reduced (Additional file 1: Fig. S5D and E). This suggests that the current and potential functions of mitochondria in Fars2-deficient cells were disrupted. To further validate the effect on mitochondria, mitochondrial ultrastructure were directly observed using transmission electron microscopy in mouse cortex, cultured neurons and PC12 cells. In Fars2-deficient organisms, a generalized phenomenon was observed that mitochondrial cristae was linearized and various geometrical shapes were formed, and the phenotype deteriorated over time(Fig. 6J–M; Additional file 1: Fig. S5F and G). Images of a whole cell from 0.5 d postnatal mouse cortex even revealed a disintegration fate of Fars2 deficiency (Fig. 6J). These morphological changes are believed to be associated with alterations in protein composition within mitochondrial protein complexes that influence inner mitochondrial membrane folding and curvature . What’s more, the fate of Fars2-knockdown PC12 cell was measured by Annexin V/ PI staining, the results showed the number of early and late apoptosis cells were both increased in comparison to the control cells (Additional file 1: Fig. S5H).
The above findings, together, indicate that mitochondrial function is severely impaired in Fars2-deficient cells and animals under the condition of impaired OXPHO complexes, and that the accumulation of damaged mitochondria may eventually lead to cells apoptosis.
Aberrant fars2 function causes reduced locomotor capacity in zebrafish
Zebrafish model was made to obtain behavioral data that correlated with spastic paraplegia . We investigated fars2 expression during embryonic development in zebrafish by qRT-PCR. fars2 is expressed early in the zebrafish embryo, and its expression increases between 14-hpf and 24-hpf (Additional file 1: Fig. S6A). fars2-knockdown zebrafish were obtained by microinjecting specific morpholino (MO) antisense oligonucleotides into fertilized one-cell stage embryos, and hb9:eGFP transgenic lines were used to study motor neurons and axons. qRT-PCR was used to confirm the effectiveness of fars2 knockdown (Additional file 1: Fig. S6B and C). At 54 h post fertilization (hpf), both strains of fars2-knockdown zebrafish present curved bodies and U-shaped somites (Fig. 7A and B). Motor axon lengths were significantly decreased in fars2 morphants, indicating impaired motor axon growth due to fars2 deficiency (Fig. 7C and D). What’s more, early fars2 deficiency during the embryonic period caused lethality. We calculated the survival rate of fars2-deficient zebrafish and discovered a significant decrease within 24-hpf, leaving 20–50% of surviving zebrafish (Fig. 7E).
Locomotor behavioral tests were then performed on 54-hpf larvae. In fars2 morphants, the stereotypic escape response, which is normal in control larvae, was greatly reduced or absent (Fig. 7F; Additional file 2: movie S1). The digital tracks and corresponding heat-maps were recorded to further evaluate the motor capacity of the fish (Fig. 7G and H). The total movement distance, velocity, mobility, and maximum acceleration in the fars2-MO group were significantly lower than those of the control-MO injected groups, suggesting a phenotype similar to that of patients with HSP (Fig. 7I-L).
Potential downstream pathways during neuronal development in response to Fars2 deficiency
To identify the molecular mechanisms and the dynamic changes underlying the neurodegenerative phenotypes caused by Fars2 deficiency, we performed RNA-seq analysis at 1,3,5,7, and 10 days after infection using mouse primary cultured neurons. The Pearson correlation value between each sample was calculated based on the fragments per kilobase of exon model per million mapped fragments (FPKM) value of all the genes. The results showed that the correlation coefficients of samples within each group were all close to 1, indicating good biological repetition effect of samples within groups. The correlation coefficients between groups at different time points showed that the Pearson value of 1 d shFars2 vs. 3 d shFars2 were slightly higher than that of 1 d shCtrl vs. 3 d shCtrl. Similarly, the value of 3 d shFars2 vs. 5 d shFars2, 5 d shFars2 vs. 7 d shFars2 and 7 d shFars2 vs. 10 d shFars2 was also slightly higher than that of 3 d shCtrl vs. 5 d shCtrl. 5 d shCtrl vs. 7 d shCtrl and 7 d shCtrl vs. 10 d shCtrl (Fig. 8A). These results suggest that the overall developmental process of shFars2 neurons is slower than that of shCtrl neurons, which is consistent with our previously observation in primary cultured neurons and the developmental delay manifestation in human patients.
In order to explore the interaction between shFars2 and shCtrl group differentially expressed gene sets at different developmental stages of neurons, we drew Venn plots for DEGs at each time point. The results showed that there were specific DEGs at each time point, while overlapped DEGs also existed. In addition to the Fars2 gene, Dctn3 and Bcl2l13 have different expression pattern in Fars2-deficient samples at each neuronal development period (Fig. 8B). qRT-PCR validated that Dctn3 decreased immediately after Fars2 knockdown both in vitro and in vivo, and kept transcribing at lower level in comparison to control neurons during the whole developmental process (Fig. 8C and D).
To further explore the signal transduction alteration within Fars2-knockdown neurons at different developmental stages. We calculated the number of DEGs of shFars2 vs. shCtrl group of 3 , 5 , 7 and 10 days spost-infection. The number increased steadily along with cultured time, and peaked on the seventh day after infection; 1441 up-regulated transcripts and 836 down-regulated transcripts met the statistical significance threshold (p < 0.05, fold change > 1.5 or 0.67), and then the change began to converge (Fig. 8E). Volcano plot displayed that the expression of some cell cycle regulation related genes such as Cdc family and Cdkn family were up-regulated in shFars2 neurons at 3 d post-infection. At the stage of neurite extension on day-5, genes related to microtubule formation and axon development, such as Tuba8 and Kif19a, as well as genes related to cell adhesion, such as Mag and Cldn11, were significantly up-regulated in shFars2 group. While the expression level of cilia movement related gene, Rsph4a, decreased significantly, which may affect the growth of neuronal processes. On day-7, the expressions of some apoptosis-related genes including Naip1 and Peg10 began to decrease, the inhibition of Peg10 was reported to activate apoptosis , which was consistent with the increase of apoptosis level of shFars2 neurons on day-8. According to our previous results, the function of mitochondria in shFars2 group neurons were severely damaged at 10 days post-infection. Transcriptome results showed that Atp12a, a gene related to the maintenance of membrane electrochemical gradient, was up-regulated in shFars2 group neurons, and the expression level of a lipid metabolism related gene, Alox8, and a carbon dioxide transfer related gene, Car14, both up-regulated, which might contribute to regulating intracellular pH value (Fig. 8F). GO enrichment analysis showed the DEGs of each time points were enriched in different basic biological processes. On day 3, DEGs were mainly enriched in the pathways related to cell cycle regulation; On day 5, the enriched pathways were mainly related to cell morphology and development. On day 7, in addition to development-related pathways, pathways related to intercellular signal transduction were also significantly enriched. On day 10, several cell death regulation pathways were enriched (Fig. 8G). GSEA of primary cultured neuron transcriptomes revealed that genes involved in axon ensheathment are upregulated in Fars2 knocked-down group at 7th day after infection, and genes involved in NF-κB signaling pathway are upregulated in Fars2 knocked-down neurons at 10th day after infection (Fig. 8H).
Mutations in genes that encode mitochondrial aminoacyl-tRNA synthetase are increasingly linked to human neurodegenerative diseases. In a previous study, we discovered p.D142Y, a novel FARS2 mutation that causes pure type HSP. In the present study, we reviewed all FARS2 mutations associated with human disease and discovered a high degree of heterogeneity in the onset of symptoms and clinical outcomes in individuals with different mutation sites and types. What’s more, a general loss of enzymatic function mechanism was implied in human cases. Establishing reliable in vitro and in vivo models with FARS2 deficiency will allow for the development of formal diagnostic criteria for FARS2 deficiency, and the promotion of further research into its associated pathogenic mechanism. And in this study, we mainly discuss the effect of great loss of FARS2 canonical function in nerous systems.
Magnetic resonance imaging (MRI) in patients with FARS2 mutations reveals various degrees of reduced cerebral white matter volumes, a common feature within brain structures . White matter is composed of growing axon and dendrites and serves an important role in providing structural connectivity between gray matter regions via the formation and organization of neural networks, and further facilitates neurobehavioral operations . The process of neurite outgrowth is energy-demanding. Mitochondrial dysfunction may therefore produce abnormalities in these neural processes, resulting in the onset and/or progression of neurodegenerative diseases such as epilepsy and Alzheimer’s disease [43, 44]. In the present study, mouse neurons deficient for Fars2 exhibited highly-impaired neurite outgrowth, preventing neuron network formation. This is consistent with white matter loss in the brain of patients with FARS2 mutations. The strong degeneration of motor neurons in zebrafish, and excessive neuronal death within the motor cortex in Fars2-deficient mouse models, clarified the spastic paraplegia phenotype observed in human patients with FARS2 mutations. This is further supported by the observed reduction in motor ability in the surviving zebrafish.
In early-onset epileptic encephalopathy patients with FARS2 mutations, global brain atrophy, and cortical atrophy in particular, is a general MRI change in the later course of the disease. The present data show that Fars2-deficient mouse embryos have a thinner cortex and an enlarged ventricle; these correspond to the neural atrophy observed in human brain. Subsequent experiments suggested that progressive neuronal apoptosis is a response to cortical tissue thinning that eventually leads to brain atrophy. This biological response is a common occurrence in many late-stage neurometabolic disorders.
Apart from changes in MRI, most FARS2-mutant patients exhibited elevated lactate levels; this may signal anaerobic metabolism caused by mitochondrial dysfunction. In the present study, mitochondrial function was comprehensively examined in animal models and in vitro. Fars2-deficient mouse embryos had a lower mitochondrial inner membrane potential, indicating lower mitochondrial activity. Furthermore, the strong reduction in ATP levels suggested that mitochondrial bioenergetic functions were reduced. ROS levels are a sign of oxidant-antioxidant homeostasis in organisms because they are a byproduct of mitochondrial oxidative metabolism . Low expression of Fars2 in vitro and in vivo leads to elevated levels of ROS. Excessive ROS availability contributes to neuronal cell death via stimulation of those biochemical pathways associated with oxidative stress and cellular apoptosis [46, 47]. The NAD+/NADH dinucleotide pair maintains a proper NAD+/NADH ratio under physiological conditions by driving a wide range of reduction–oxidation (redox) reactions and inhibiting excessive ROS generation . Total NAD of the mitochondria pool, however, were reduced in Fars2-deficient tissues and cells; this suggests an imbalance between oxidant and antioxidant levels in organisms deficient for Fars2.
Given the classic aminoacylation function of mtPheRS, disorganized mitochondria translation machinery is a prime suspect in Fars2 deficiency-induced mitochondrial dysfunction. The present data reveal that Fars2 deficiency affects the expression levels of OXPHOS-subunit protein and subsequently the stable levels of OXPHO complexes. In the mouse brain, Fars2-KO reduced mitochondrial CI, CIII, and CIV; Fars2-knockdown in primary neurons in culture, however, only affected CI of the electron transport chain. This phenotypic heterogeneity was also observed in patients with FARS2 mutations, and could be attributed to varying degrees of aminoacylation function loss in different organisms.
Results in this study indicate that CI is more vulnerable to Fars2 deficiency. It is reported that CI deficiency is the most common OXPHOS disorder and may be due to the complicated and multifaceted assembly process required by CI [49, 50]. As the first enzyme in OXPHOS, CI oxidizes NADH into NAD+ and drives proton translocation across the mitochondrial inner membrane together with CIII and CIV; this forms the mitochondrial membrane potential that then promotes ATP generation by CV . The present data show that expression of CI level was severely reduced in both in vitro and in vivo Fars2-deficient environments, leading to the reduced conversion of NADH to NAD+ and limited proton translocation. Reduced electron transfer produces mitochondrial depolarization, and the release of ROS ultimately diminishes ATP production.
With the exception of decreased CI, CIII, and CIV, the slight enhancement of CII and CV and the general upregulation of subunit mRNA levels highlight a compensatory mechanism against dysfunctional mitochondrial translation. Furthermore, transcriptome sequencing analysis revealed that the Fars2-defect induced neuronal dysplasia may closely related to the activation of axon sheathing process and NF- κB signaling pathway during neuron development. The myelin sheath is critical for increasing neural processing speed and efficiency , whose activation might be a compensatory rescue pathway in respond to Fars2 knockdown in neurons. NF-κB pathway activation is reported to be required for the initiation of apoptosis, and also be a key mediator of neuroinflammation in various neurodegeneration disease including Alzheimer’s Disease and Parkinson’s Disease [53, 54].
Although several researches have suggested non-canonical function, such as post-transcriptional regulation of mRNA expression, in some aaRSs . No evidence for FARS2 function other than mitochondrial translation activity has been shown yet. While in this study, transcriptome analysis reveals a potential downstream molecule of Fars2 deficiency, Dctn3. Dctn3 is a subunit of Dynactin, which is a multisubunit protein complex required for activating dynein and regulating retrogradely transports cargo along microtubules . Dysregulation of retrograde axon transport is considered to be a pivotal pathogeny of motor neuron degeneration disease . These evidence suggests that in addition to mitochondrial dysfunction, disruption of retrograde axon transport induced by Dctn3 reduction might be another possible pathogenesis of Fars2-related neuropathy.
A limitation of this study is that we only focus on Fars2-deficient neurons and their fate of apoptosis. However, apoptotic cells were observed both in dorsal and ventral cortex region of E 14.5 cKO mouse embryos during cortical neurogenesis, which suggests that neural progenitors might also be affected. Whether and how Fars2 deficiency affect the proliferation and differentiation of neural progenitors would be an important aspect to study in the future.
To sum up, we have successfully established mouse and zebrafish models of Fars2 deficiency that mimic the neuropathy and metabolism changes seen in human patients. We found that dysfunction caused by disruption to mitochondrial translation leads to neuronal apoptosis and possibly neurodegenerative diseases. Taken together, our study emphasizes the critical role of Fars2 in mitochondrial function and its pathological consequences for neuronal development and maintenance (Fig. 9).
Materials and methods
Animal models were established under technical and facility support of Shanghai Model Organisms Center. All experiments were conducted with ethics approval from the Air Force Medical University under regulations of the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health.
Global Fars2 -mutant and Fars2 -KO mice
Mice with whole-body p.D142Y mutant and KO were generated by using CRISPR/Cas 9 technology. In brief, Cas9 mRNA, guide RNA were obtained by in vitro transcription. Oligo donor DNA with mutant point was synthesized. The F0 p.D142Y mice was generated by microinjecting Cas9 mRNA, guide RNA (GGGACAACTATTACTTGAAT) and donor DNA (GACCAGCTTCCTCCAGTGGTCACCACCTGGCAGAACTTTGATAGCCTGCTAATCCCAGCTGACCACCCCAGCAGGAAAAAGGGGTACAACTATTACTTGAATCGCGCACACATGCTGAGAGCACACACATCAGCGCATCAGTGGGACTTGCTGCATGCGGGACTTAATGCCTTCCTTGTG) into the oosperm of C57BL/6J mice. The F0 KO mice was generated by microinjecting Cas9 mRNA, guideRNA (GGGACAACTATTACTTGAAT) into the oosperm of C57BL/6J mice. The genotypes of offspring mice were determined by PCR and Sanger sequencing.
Conditional Fars2 -KO mice
LoxP were inserted into Fars2 of C57BL/6J mice by injecting Cas9 mRNA, guide RNA (gRNA#1: AGCAAGCTCTGAGCTACCCAGGG; gRNA#2: GGACAAGATGCTTCACAATATGG) and donor vector into the oosperm to obtain Fars2 fl/fl mice. CNS-specific Fars2-deficient mice were generated by mating Fars2 fl/fl with transgenic mice containing Nestin driven Cre recombinase . The Nestin-Cre mouse was purchased from Shanghai Model Organisms (China). The genotypes of offspring mice were determined by PCR and Nucleic acid gel electrophoresis.
Adult wild-type AB strain zebrafish pairs were mated under 28.5 ℃ on a 14 h light / 10 h dark cycle in fish water (0.2% Instant Ocean Salt in deionized water). An average of 200–300 embryos were generated. The embryos were washed and staged according to the previous study . The hb9:eGFP transgenic line was established according to the previous described . fars2 translation-blocking morpholino (ATG-MO), splice-blocking morpholino (E3I3-MO) and standard control morpholino (MO) were designed and produced by Gene Tools, LLC (http://www.gene-tools.com/). The MO sequences were listed in Additional file 1: Table S1. MOs were injected into fertilized one-cell stage embryos according to standard protocols . The amount of the MOs used for injection was as follows: 4ng Control-MO and ATG-MO, 8ng E3I3-MO were used for injection per embryo. RT-PCR was performed to confirm the efficacy of the E3I3-MO. ef1α was used as the internal control. The sequences of the primers were listed in Additional file 1: Table S2. The zebrafish facility at Shanghai Model Organisms Center is accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International.
Zebrafish spinal motor neurons studies
To evaluate spinal motor neurons formation in zebrafish, fertilized one-cell hb9:eGFP transgenic lines embryos were injected with 4ng control-MO, 4ng fars2-ATG-MO or 8ng fars2-E3I3-MO. At 54-hpf, embryos were dechorionated, anesthetized with 0.016% tricaine methanesulfonate (Sigma-Aldrich). Zebrafish were then oriented on lateral side or dorsal side, and mounted with 3% methylcellulose in a depression slide for observation by fluorescence microscopy. The phenotypes of spinal motor neurons were analyzed.
Stereotypic escape response assays
54-hour-old living zebrafish larvae were dechorionated manually at least 3 h before the experiment. To evaluate the escape response, fish were touched with the tip of a fine needle for at least 2 times at the dorsal tip of the tail or trunk. An escape response in which the fish did not move a distance of at least 3 times its own body length was considered as reduced.
At 5-dpf, the larvae were divided into three groups: control-MO, fars2-ATG-MO and fars2-E3I3-MO. Larvae from each group were collected, cleaned and placed in 96-well plates. Each well contained 0.2 ml of fish water and one larva, and ten larvae were in each group. Behavioral tests were performed as following: the larvae were allowed to acclimate for 15 min before locomotion monitoring . Next, the larvae were allowed to freely explore the aquarium for 30 min. A camera positioned above the plate was used for movement tracking. All digital tracks were analyzed by Ethovision XT software (Noldus Information Technology, Wageningen, Netherlands), and a minimum movement distance of 0.2 mm filtered out system noise. Four parameters, including the total movement distance, velocity, mobility and maximum acceleration were analyzed. Embryos and larvae were analyzed with Nikon SMZ18 Fluorescence microscope and subsequently photographed with digital cameras. A subset of images was adjusted for levels, brightness, contrast, hue and saturation with Adobe Photoshop 7.0 software (Adobe, San Jose, California) to optimally visualize the expression patterns. Quantitative image analyses processed using image based morphometric analysis (NIS-Elements D4.6, Japan) and ImageJ software (U.S. National Institutes of Health, Bethesda, MD, USA).
Brain separation and section preparation
Pregnant mice of proper days were deeply anesthetized with 10% chloral hydrate and the embryos were extracted. Fresh brain of each embryo were isolated in precooled PBS and then fixed in 4% paraformaldehyde overnight. For paraffin sections, the tissue was dehydrated in Gradient alcohol and embedded in paraffin. Coronal sections were made at a 3 μm thickness with a paraffin slicing machine (Leica, RM2125RTS). For cryostat sections, brain was placed in 30% sucrose until sink to the bottom and and embedded in Tissue Freezing Medium (Leica, 03819110). sectioned at a 12 μm thickness with a freezing microtome (Leica, CM1950).
Primary neuron culturing and lentivirus infection
Cultures of cortical neurons were prepared from embryonic day 14–16 mouse brains as described previously with some modifications. In brief, embryo cortex was isolated in pre-cooled HBSS (Gibco, 14,175,095). After removing the meninges, tissue was cut into pieces and transferred into Neurobasal Medium (Gibco, A3582901) supplemented with 2% B27 (Gibco, 17,504,044), 1% L-Glutamine (Gibco, A2916801). Blowed the tissue pieces into suspension using a pipettor. Filtrate the suspension by using a 70 μm Cell Strainers (FALCON, 352,350) to obtain single-cell suspension and then seed into dishes pre-coated by Poly-L-Lysine (Sigma, P1399). After 1 days in vitro, rLV-U6-shFars2-CMV-mcherry-WPRE and rLV-U6-shNC(scramble)-CMV-mcherry-WPRE (Brainvta Company, China) were added into the dishes. shRNA sequences were listed in Additional file 1: Table S3. The culture medium was changed after 12 hours’ infection. Half amount of medium was changed every 2 days in the following cultivation.
Primary cultured neurons were seeded on round glass coverslip in 24-well plates at a density of 2 × 105 per well. After infection, cells were fixed and proceed to regular immunofluorescence at indicated days. The following antibodies were used: MAP2 (Abcam, ab11267, 1:1000), Neuron-specific β- Tubulin (TUJ1) antibody (R&D, MAB1195-SP, 1:500). Secondary Antibodies was purchased from Jackson ImmunoResearch: Alexa Fluor® 488-conjugated AffiniPure Goat Anti-mouse IgG (H + L) (115-545-003).
Hematoxylin and eosin staining
Embryos at E 8.0 were isolated from the pregnant mice after cross-mating and fixed with 4% paraformaldehyde, dehydration in 30% sucrose and embedded in Tissue Freezing Medium. Cryostat sections with 12 μm thickness were made by a freezing microtome and stained with Hematoxylin and Eosin Staining Kit (Beyotime, C0105S). The images were detected under a light microscopy.
Paraffin sections of embryo brain were prepared and incubated for 45 s in Nissl solution (Beyotime, C0117) followed by dehydration with different ethanol concentrations as described in the protocol of the instructions.
Apoptotic cells were detected by using Fluorescein (FITC) Tunel Cell Apoptosis Detection Kit (Servicebio, G1501) according to the instruction. In brief, cells coverslips or mouse brain sections were fixed with 4% paraformaldehyde, rinsed with PBS, and then permeabilized by 0.1% Triton X-100. The TUNEL reaction mixture was used for FITC end-labeling the fragmented DNA of the apoptotic cells. The cell nuclei were stained with 2 mg/ml DAPI (Invitrogen, D1306). The FITC-labeled TUNEL-positive cells were imaged under a fluorescence microscope.
Detection of cellular ATP levels
Cellular ATP levels were measured using a firefly luciferasebased ATP assay kit (Beyotime, S0026) according to the manufacturer’s instructions. Briefly, cortical tissue or harvested cultured cells were lysed with a lysis buffer, and centrifuged at 12,000 × g for 5 min. 20 µl of each supernatant or standard substance was mixed with 100 µl luciferase reagent. Luminance (RLU) was measured by a multimode microplate reader (Tecan Spark). Standard curves were generated and the protein concentration of each sample was measured by BCA protein assay. Total ATP levels were expressed as pmol/ng protein.
Measurement of reactive oxygen species
In brief, adherent neurons or cell suspension from cortical tissue were incubated with 6-Carboxy-2′, 7′-dichlorodihydrofluorescein diacetate (DCFH-DA) (Beyotime, S0033S) at a final concentration of 10 mM for 20 min and washed 3 times with PBS. And then ROS generation was measured by the fluorescence intensity with excitation and emission settings at 488 and 525 nm.
Measurement of NAD level
NADtotal levels and NAD+/NADH were determined using NAD+/NADH assay kit with WST-8 (Beyotime, S0175) according to the manufacturer’s instructions. In brief, adherent neurons or cortical tissues were lysed with 400 µl of lysis buffer, and centrifuged at 12,000 ×g for 10 min. 90 µl of alcohol dehydrogenase was added to a 96-well plate. NADtotal levels were obtained by adding 20 µl of the supernatant or standard substance. And NADH levels were obtained by adding 20 µl of the suspension or standard substance after incubating at 60 °C for 30 min and was added to a 96-well plate. Subsequently, 10 µl of chromogenic solution was added to the plate and the mixture was incubated at 37 °C for 30 min. The absorbance values were measured at 450 nm and analyzed on a multimode microplate reader (Tecan Spark). Standard curve was generated and the protein concentration of each sample was measured by BCA protein assay. The amount of NAD+ was derived by subtracting NADH from NADtotal.
Transmission electron microscopy detection
Primary neurons and PC12 Cells were respectively digested using Neuronal Isolation Enzyme (with papain) (Thermo 88,285) and 0.25% Trypsin-EDTA (Gibco, 25,200,072), and then centrifuged at 800 rpm for 10 min to obtain cell pellet. Cell pellets and cortex blocks were fixed in 2.5% glutaraldehyde for 24 h and then post-fixed in 2% osmium tetroxide for 1 h. After dehydration, the specimens were embedded in epoxide resin. Ultrathin sections were made and stained with uranyl acetate and lead citrate, images were captured at a magnification of 20,000 × and 300,000 × on a Zeiss electron microscope.
Measurement of mitochondrial membrane potential
JC-1 kit (Beyotime, C2006) was employed to estimate mitochondrial depolarization in cortical tissue and PC12 cell. Briefly, cell suspension from cortical tissue or adherent cells after indicated treatments were incubated with an equal volume of JC-1 staining solution (5 µg/ml) at 37 ℃ for 20 min and rinsed twice with PBS. Mitochondrial membrane potentials were monitored by determining the relative amounts of dual emissions from mitochondrial JC-1 monomers or aggregates indicated by the green/red fluorescence intensity ratio using FAC Scalibur flow cytometer (BD).
Preparation of mitochondrial and cytosolic fractions
Mitochondrial and cytosolic fractions were isolated using a commercially available cytosol/mitochondria fractionation kit according to the manufacturer’s protocol (Beyotime, C3606).
Cortical and neuron cell were lysed with the RIPA buffer (Beyotime, P0013B). Protein extracts were analyzed by 10% SDS-PAGE and blotted onto PVDF membrane. The following primary antibodies were used at indicated concentrations: Total OXPHOS rodent antibody (Abcam, ab110413, 1:250); ND1 antibody (Abcam, ab181848, 1:10000); CO2 antibody (Abcam, ab198286, 1:1000); CYTB antibody (LifeSpan BioSciences, 197,737, 1:500); TOMM20 antibody (Abcam, ab186735, 1:5000); Caspase-3 antibody (Cell Signaling Technology, 9662, 1:1000); Cleaved Caspase-3 antibody (Cell Signaling Technology, 9661, 1:1000); β-actin antibody (Sigma, A1978, 1:3000); FARS2 antibody (Invitrogen, PA5-53738, 1:1000). All secondary antibodies were purchased from ZhuangzhiBio (anti-mouse, 1: 8000; anti-rabbit, 1:8000) Western blotting quantification was performed using the ImageJ software.
Quantitative real-time PCR
Total RNA was isolated from tissue or cultured cells using Multisource Total RNA Miniprep Kit (Axygen, 365), and then used for cDNA synthesis (PrimeScript™RT Master Mix, Takara, RR036Q) and amplification by real-time PCR according to the manufacturer’s instructions (SYBR Premix Ex Taq™ II, Takara, RR820A). Relative gene expression quantification was based on the comparative threshold cycle method (2−ΔΔCt) using β-actin as endogenous control gene. The primer sequences were provided in Additional file 1: Table S2.
RNA extraction, library construction and transcriptomic sequencing
Total RNA from primary cultured neurons after 1,3,5,7,10 days of infection was extracted using Multisource Total RNA Miniprep Kit (Axygen, 365). RNA quality was performed using Agilent 2100 Bioanalyzer (Agilent Technologies, USA) and checked using RNase free agarose gel electrophoresis. After qualification, total mRNA was enriched by Oligo (dT) beads (Epicentre, USA). RNA was fragmented into short fragments and reverse transcript into cDNA with random primers. Second-strand cDNA were synthesized and the cDNA fragments were purified with QiaQuick PCR extraction kit (Qiagen, Netherlands), end repaired, poly A added, and ligated to Illumina sequencing adapters. Selecting ligation products with proper size by PCR and agarose gel electrophoresis, and then sequencing was performed by Gene Denovo Biotechnology Co. (Guangzhou, China) on Illumina HiSeq2500.
The obtained reads were filtered by fastp (version 0.18.0) to get high quality clean reads without adapters or low-quality bases. An index of the reference genome was built, and paired-end clean reads were aligned to the mouse (mm10) genomes from UCSC using HISAT2 (v2.1.0). For RNAs differential expression analysis, reads were pseudo aligned to mouse (GENCODE GRCm38 release M16) transcriptomes using DESeq2 software between two different groups. Transcripts with log2 |fold change| > 1 and false discovery rate (FDR) < 0.05 were considered differentially expressed. Gene set enrichment analysis (GSEA), venn diagram, volcano Plot and gene ontology (GO) enrichment analysis were performed at Omicsmart online platform (https://www.omicsmart.com) based on the screened out differentially expressed genes (DEGs).
Data were analyzed using GraphPad Prism 8 software. Statistical significance was calculated using student’s t test or ANOVA as appropriate. Data are presented as mean ± SD or SEM. Asterisks indicate the level of statistical significance, *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.
The datasets used and analysed during the current study are available from the corresponding author on reasonable request.
Cytochrome c Oxidase Subunit 2
Complex I-Complex V
Differentially expressed genes
- E3I3- MO:
Fragments Per Kilobase of exon model per Million mapped fragments
Green Fluorescent Protein
Gene Set Enrichment Analysis
hours post fertilization
Hereditary Spastic Paraplegia
Kyoto Encyclopedia of Genes and Genomes
Microtubule Associated Protein 2
Magnetic resonance imaging
mitochondrial aminoacyl-tRNA synthetases
mitochondrial phenylalanyl-tRNA synthetase
phenylalanine mitochondria tRNA
Nicotinamide Adenine Dinucleotide
NADH Dehydrogenase Subunit 1
Reactive Oxygen Species
scramble shRNA as a negative control
- shFars2 :
Translocase of Outer Mitochondrial Membrane 20
Mitochondrial Membrane Potential
Chihade J. Mitochondrial aminoacyl-tRNA synthetases. Enzymes. 2020;48:175–206.
Sissler M, Gonzalez-Serrano LE, Westhof E. Recent advances in mitochondrial aminoacyl-tRNA synthetases and disease. Trends Mol Med. 2017;23(8):693–708.
Scheper GC, van der Klok T, van Andel RJ, van Berkel CG, Sissler M, Smet J, Muravina TI, Serkov SV, Uziel G, Bugiani M, et al. Mitochondrial aspartyl-tRNA synthetase deficiency causes leukoencephalopathy with brain stem and spinal cord involvement and lactate elevation. Nat Genet. 2007;39(4):534–9.
Steenweg ME, Ghezzi D, Haack T, Abbink TE, Martinelli D, van Berkel CG, Bley A, Diogo L, Grillo E, Te WNJ, et al. Leukoencephalopathy with thalamus and brainstem involvement and high lactate “LTBL” caused by EARS2 mutations. Brain. 2012;135(Pt 5):1387–94.
Elo JM, Yadavalli SS, Euro L, Isohanni P, Gotz A, Carroll CJ, Valanne L, Alkuraya FS, Uusimaa J, Paetau A, et al. Mitochondrial phenylalanyl-tRNA synthetase mutations underlie fatal infantile Alpers encephalopathy. Hum Mol Genet. 2012;21(20):4521–9.
Bayat V, Thiffault I, Jaiswal M, Tetreault M, Donti T, Sasarman F, Bernard G, Demers-Lamarche J, Dicaire MJ, Mathieu J, et al. Mutations in the mitochondrial methionyl-tRNA synthetase cause a neurodegenerative phenotype in flies and a recessive ataxia (ARSAL) in humans. PLoS Biol. 2012;10(3): e1001288.
Edvardson S, Shaag A, Kolesnikova O, Gomori JM, Tarassov I, Einbinder T, Saada A, Elpeleg O. Deleterious mutation in the mitochondrial arginyl-transfer RNA synthetase gene is associated with pontocerebellar hypoplasia. Am J Hum Genet. 2007;81(4):857–62.
Konovalova S, Tyynismaa H. Mitochondrial aminoacyl-tRNA synthetases in human disease. Mol Genet Metab. 2013;108(4):206–11.
Fine AS, Nemeth CL, Kaufman ML, Fatemi A. Mitochondrial aminoacyl-tRNA synthetase disorders: an emerging group of developmental disorders of myelination. J Neurodev Disord. 2019;11(1):29.
Hotait M, Nasreddine W, El-Khoury R, Dirani M, Nawfal O, Beydoun A. FARS2 mutations: more than two phenotypes? A case report. Front Genet. 2020;11:787.
Yang Y, Liu W, Fang Z, Shi J, Che F, He C, Yao L, Wang E, Wu Y. A newly identified missense mutation in FARS2 causes autosomal-recessive spastic paraplegia. Hum Mutat. 2016;37(2):165–9.
Peretz M, Tworowski D, Kartvelishvili E, Livingston J, Chrzanowska-Lightowlers Z, Safro M. Breaking a single hydrogen bond in the mitochondrial tRNA(Phe) -PheRS complex leads to phenotypic pleiotropy of human disease. FEBS J. 2020;287(17):3814–26.
Roy H, Ling J, Alfonzo J, Ibba M. Loss of editing activity during the evolution of mitochondrial phenylalanyl-tRNA synthetase. J Biol Chem. 2005;280(46):38186–92.
Vantroys E, Larson A, Friederich M, Knight K, Swanson MA, Powell CA, Smet J, Vergult S, De Paepe B, Seneca S, et al. New insights into the phenotype of FARS2 deficiency. Mol Genet Metab. 2017;122(4):172–81.
Sun L, Wei N, Kuhle B, Blocquel D, Novick S, Matuszek Z, Zhou H, He W, Zhang J, Weber T, et al. CMT2N-causing aminoacylation domain mutants enable Nrp1 interaction with AlaRS. Proc Natl Acad Sci USA. 2021;118(13):e2012898118.
Klipcan L, Moor N, Finarov I, Kessler N, Sukhanova M, Safro MG. Crystal structure of human mitochondrial PheRS complexed with tRNA(Phe) in the active “open” state. J Mol Biol. 2012;415(3):527–37.
Almalki A, Alston CL, Parker A, Simonic I, Mehta SG, He L, Reza M, Oliveira JM, Lightowlers RN, McFarland R, et al. Mutation of the human mitochondrial phenylalanine-tRNA synthetase causes infantile-onset epilepsy and cytochrome c oxidase deficiency. Biochim Biophys Acta. 2014;1842(1):56–64.
Barcia G, Rio M, Assouline Z, Zangarelli C, Roux CJ, de Lonlay P, Steffann J, Desguerre I, Munnich A, Bonnefont JP, et al. Novel FARS2 variants in patients with early onset encephalopathy with or without epilepsy associated with long survival. Eur J Hum Genet. 2021;29(3):533–8.
Vernon HJ, McClellan R, Batista DA, Naidu S. Mutations in FARS2 and non-fatal mitochondrial dysfunction in two siblings. Am J Med Genet A. 2015;167A(5):1147–51.
Almannai M, Wang J, Dai H, El-Hattab AW, Faqeih EA, Saleh MA, Al AA, Alwadei AH, Aljadhai YI, AlHashem A, et al. FARS2 deficiency; new cases, review of clinical, biochemical, and molecular spectra, and variants interpretation based on structural, functional, and evolutionary significance. Mol Genet Metab. 2018;125(3):281–91.
Ville D, Lesca G, Labalme A, Portes VD, Arzimanoglou A, de Bellescize J. Early-onset epileptic encephalopathy with migrating focal seizures associated with a FARS2 homozygous nonsense variant. Epileptic Disord. 2020;22(3):327–35.
Kim SY, Jang SS, Kim H, Hwang H, Choi JE, Chae JH, Kim KJ, Lim BC. Genetic diagnosis of infantile-onset epilepsy in the clinic: application of whole-exome sequencing following epilepsy gene panel testing. Clin Genet. 2021;99(3):418–24.
Raviglione F, Conte G, Ghezzi D, Parazzini C, Righini A, Vergaro R, Legati A, Spaccini L, Gasperini S, Garavaglia B, et al. Clinical findings in a patient with FARS2 mutations and early-infantile-encephalopathy with epilepsy. Am J Med Genet A. 2016;170(11):3004–7.
Forman EB, Gorman KM, Ennis S, King MD. FARS2 causing complex hereditary spastic paraplegia with dysphonia: expanding the disease specstrum. J Child Neurol. 2019;34(10):621.
Meszarosova AU, Seeman P, Jencik J, Drabova J, Cibochova R, Stellmachova J, Safka BD. Two types of recessive hereditary spastic paraplegia in Roma patients in compound heterozygous state; no ethnically prevalent variant found. Neurosci Lett. 2020;721: 134800.
Sahai SK, Steiner RE, Au MG, Graham JM, Salamon N, Ibba M, Pierson TM. FARS2 mutations presenting with pure spastic paraplegia and lesions of the dentate nuclei. Ann Clin Transl Neurol. 2018;5(9):1128–33.
Walker MA, Mohler KP, Hopkins KW, Oakley DH, Sweetser DA, Ibba M, Frosch MP, Thibert RL. Novel compound heterozygous mutations expand the recognized phenotypes of FARS2-linked disease. J Child Neurol. 2016;31(9):1127–37.
Chen Z, Zhang Y. A patient with juvenile-onset refractory status epilepticus caused by two novel compound heterozygous mutations in FARS2 gene. Int J Neurosci. 2019;129(11):1094–7.
Roux CJ, Barcia G, Schiff M, Sissler M, Levy R, Dangouloff-Ros V, Desguerre I, Edvardson S, Elpeleg O, Rotig A, et al. Phenotypic diversity of brain MRI patterns in mitochondrial aminoacyl-tRNA synthetase Mutations. Mol Genet Metab. 2021;133(2):222–9.
Harvey AJ. Mitochondria in early development: linking the microenvironment, metabolism and the epigenome. Reproduction. 2019;157(5):R159–79.
Wei PZ, Szeto CC. Mitochondrial dysfunction in diabetic kidney disease. Clin Chim Acta. 2019;496:108–16.
Addeo M, Buonaiuto S, Guerriero I, Amendola E, Visconte F, Marino A, De Angelis MT, Russo F, Roberto L, Marotta P, et al. Insight into nephrocan function in mouse endoderm patterning. Int J Mol Sci. 2019;21(1):8.
Saga Y, Hata N, Koseki H, Taketo MM. Mesp2: a novel mouse gene expressed in the presegmented mesoderm and essential for segmentation initiation. Genes Dev. 1997;11(14):1827–39.
Osumi N, Shinohara H, Numayama-Tsuruta K, Maekawa M. Concise review: Pax6 transcription factor contributes to both embryonic and adult neurogenesis as a multifunctional regulator. Stem Cells. 2008;26(7):1663–72.
Papa S, Martino PL, Capitanio G, Gaballo A, De Rasmo D, Signorile A, Petruzzella V. The oxidative phosphorylation system in mammalian Mitochondria. Adv Exp Med Biol. 2012;942:3–37.
Dogan SA, Pujol C, Maiti P, Kukat A, Wang S, Hermans S, Senft K, Wibom R, Rugarli EI, Trifunovic A. Tissue-specific loss of DARS2 activates stress responses independently of respiratory chain deficiency in the heart. Cell Metab. 2014;19(3):458–69.
Liu S, Liu S, He B, Li L, Li L, Wang J, Cai T, Chen S, Jiang H. OXPHOS deficiency activates global adaptation pathways to maintain mitochondrial membrane potential. Embo Rep. 2021;22(4): e51606.
Ying W. NAD+/NADH and NADP+/NADPH in cellular functions and cell death: regulation and biological consequences. Antioxid Redox Signal. 2008;10(2):179–206.
Vincent AE, Ng YS, White K, Davey T, Mannella C, Falkous G, Feeney C, Schaefer AM, McFarland R, Gorman GS, et al. The spectrum of mitochondrial ultrastructural defects in mitochondrial myopathy. Sci Rep. 2016;6:30610.
Babin PJ, Goizet C, Raldua D. Zebrafish models of human motor neuron diseases: advantages and limitations. Prog Neurobiol. 2014;118:36–58.
Lu X, Yang S, Jie M, Wang S, Sun C, Wu L, Chang S, Pei P, Wang S, Zhang T, et al. Folate deficiency disturbs PEG10 methylation modifications in human spina bifida. Pediatr Res. 2021. https://doi.org/10.1038/s41390-021-01908-6.
Filley CM, Fields RD. White matter and cognition: making the connection. J Neurophysiol. 2016;116(5):2093–104.
Dulla CG, Coulter DA, Ziburkus J. From molecular circuit dysfunction to disease: case studies in epilepsy, traumatic brain injury, and Alzheimer’s disease. Neuroscientist. 2016;22(3):295–312.
Klein GT, Van Hugte E, Frega M, Guardia GS, Foreman K, Panneman D, Mossink B, Linda K, Keller JM, Schubert D, et al. m.3243A > G-induced mitochondrial dysfunction impairs human neuronal development and reduces neuronal network activity and synchronicity. Cell Rep. 2020;31(3):107538.
Singh A, Kukreti R, Saso L, Kukreti S. Oxidative stress: a key modulator in neurodegenerative diseases. MOLECULES. 2019;24(8):1583.
Mariani E, Polidori MC, Cherubini A, Mecocci P. Oxidative stress in brain aging, neurodegenerative and vascular diseases: an overview. J Chromatogr B Analyt Technol Biomed Life Sci. 2005;827(1):65–75.
Sen T, Sen N, Jana S, Khan FH, Chatterjee U, Chakrabarti S. Depolarization and cardiolipin depletion in aged rat brain mitochondria: relationship with oxidative stress and electron transport chain activity. Neurochem Int. 2007;50(5):719–25.
Anderson KA, Madsen AS, Olsen CA, Hirschey MD. Metabolic control by sirtuins and other enzymes that sense NAD(+), NADH, or their ratio. Biochim Biophys Acta Bioenerg. 2017;1858(12):991–8.
Guerrero-Castillo S, Baertling F, Kownatzki D, Wessels HJ, Arnold S, Brandt U, Nijtmans L. The assembly pathway of mitochondrial respiratory chain complex I. Cell Metab. 2017;25(1):128–39.
Skladal D, Halliday J, Thorburn DR. Minimum birth prevalence of mitochondrial respiratory chain disorders in children. Brain. 2003;126(Pt 8):1905–12.
Nouws J, Nijtmans LG, Smeitink JA, Vogel RO. Assembly factors as a new class of disease genes for mitochondrial complex I deficiency: cause, pathology and treatment options. Brain. 2012;135(Pt 1):12–22.
Djannatian M, Timmler S, Arends M, Luckner M, Weil MT, Alexopoulos I, Snaidero N, Schmid B, Misgeld T, Mobius W, et al. Two adhesive systems cooperatively regulate axon ensheathment and myelin growth in the CNS. NAT COMMUN. 2019;10(1):4794.
Ju HC, Choi DY, Park MH, Hong JT. NF-kappaB as a key mediator of brain inflammation in Alzheimer’s disease. CNS Neurol Disord Drug Targets. 2019;18(1):3–10.
Singh SS, Rai SN, Birla H, Zahra W, Rathore AS, Singh SP. NF-kappaB-mediated neuroinflammation in Parkinson’s disease and potential therapeutic effect of polyphenols. Neurotox Res. 2020;37(3):491–507.
Garin S, Levi O, Forrest ME, Antonellis A, Arava YS. Comprehensive characterization of mRNAs associated with yeast cytosolic aminoacyl-tRNA synthetases. RNA Biol. 2021;18(12):2605–16.
Shen C, Honda H, Suzuki SO, Maeda N, Shijo M, Hamasaki H, Sasagasako N, Fujii N, Iwaki T. Dynactin is involved in Lewy body pathology. Neuropathology. 2018;38(6):583–90.
Eschbach J, Dupuis L. Cytoplasmic dynein in neurodegeneration. Pharmacol Ther. 2011;130(3):348–63.
Tronche F, Kellendonk C, Kretz O, Gass P, Anlag K, Orban PC, Bock R, Klein R, Schutz G. Disruption of the glucocorticoid receptor gene in the nervous system results in reduced anxiety. Nat Genet. 1999;23(1):99–103.
Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. Stages of embryonic development of the zebrafish. Dev Dyn. 1995;203(3):253–310.
Kanungo J, Lantz S, Paule MG. In vivo imaging and quantitative analysis of changes in axon length using transgenic zebrafish embryos. Neurotoxicol Teratol. 2011;33(6):618–23.
Nasevicius A, Ekker SC. Effective targeted gene “knockdown” in zebrafish. Nat Genet. 2000;26(2):216–20.
Zhao T, Zondervan-van DLH, Severijnen LA, Oostra BA, Willemsen R, Bonifati V. Dopaminergic neuronal loss and dopamine-dependent locomotor defects in Fbxo7-deficient zebrafish. PLoS ONE. 2012;7(11): e48911.
Hilgenberg LG, Smith MA. Preparation of dissociated mouse cortical neuron cultures. J Vis Exp. 2007;10:562.
All staff from Shaanxi Provincial Key Laboratory of Clinic Genetics deserve our profuse thanks for their indefatigable and meticulous work as study coordinator. And also, I (KC) want to thank my newborn son (Chufan Chen) for his great support.
This study was supported by the Key Research and Development Plan in Shaanxi, Grant/Award Number: 2019SF-059 and 2020SF-204; the Key Innovative Project in Shaanxi, Grant/Award Number: 2021ZDLSF02-02; National Natural Science Foundation of China, Grant/Award Number: 31570906.
Ethics approval and consent to participate
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Fars2 expression pattern during mouse embryonic development and in multiple organs of E 17 mouse embryos. Figure S2. Homozygous Fars2 knockout mouse cannot survive to born. Figure S3. Establishment of conditional neural-specific Fars2 knockout-mouse model. Figure S4. Establishment of Fars2-knockdown neurons in vitro using shFars2 lentivirus. Figure S5. Mitochondrial dysfunction was confirmed in PC12 cell line. Figure S6. Effectiveness of fars2 knockdown was confirmed by RT-PCR. Table S1. Table of Morpholino sequences. Table S2. Table of primer sequences in qPCR. Table S3. Table of shRNA sequences. Table S4. Table of siRNA sequences. and Table S5. Proportion of phenylalanine in 5 mitochondrial complexes in human, mouse, zebrafish and rat.
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Chen, X., Liu, F., Li, B. et al. Neuropathy-associated Fars2 deficiency affects neuronal development and potentiates neuronal apoptosis by impairing mitochondrial function. Cell Biosci 12, 103 (2022). https://doi.org/10.1186/s13578-022-00838-y
- Mitochondrial phenylalanyl-tRNA synthetase
- Neurite outgrowth
- Mitochondrial dysfunction
- Oxidative phosphorylation complexes
- Zebrafish model