A loss-of-function mutation in human Oxidation Resistance 1 disrupts the spatial–temporal regulation of histone arginine methylation in neurodevelopment
Genome Biology volume 24, Article number: 216 (2023)
Oxidation Resistance 1 (OXR1) gene is a highly conserved gene of the TLDc domain-containing family. OXR1 is involved in fundamental biological and cellular processes, including DNA damage response, antioxidant pathways, cell cycle, neuronal protection, and arginine methylation. In 2019, five patients from three families carrying four biallelic loss-of-function variants in OXR1 were reported to be associated with cerebellar atrophy. However, the impact of OXR1 on cellular functions and molecular mechanisms in the human brain is largely unknown. Notably, no human disease models are available to explore the pathological impact of OXR1 deficiency.
We report a novel loss-of-function mutation in the TLDc domain of the human OXR1 gene, resulting in early-onset epilepsy, developmental delay, cognitive disabilities, and cerebellar atrophy. Patient lymphoblasts show impaired cell survival, proliferation, and hypersensitivity to oxidative stress. These phenotypes are rescued by TLDc domain replacement. We generate patient-derived induced pluripotent stem cells (iPSCs) revealing impaired neural differentiation along with dysregulation of genes essential for neurodevelopment. We identify that OXR1 influences histone arginine methylation by activating protein arginine methyltransferases (PRMTs), suggesting OXR1-dependent mechanisms regulating gene expression during neurodevelopment. We model the function of OXR1 in early human brain development using patient-derived brain organoids revealing that OXR1 contributes to the spatial–temporal regulation of histone arginine methylation in specific brain regions.
This study provides new insights into pathological features and molecular underpinnings associated with OXR1 deficiency in patients.
Oxidation Resistance 1 (OXR1, HGNC: 15,822, alias: TLDC3) is a highly conserved gene of the TLDC domain-containing family [1,2,3,4,5,6], first reported in Escherichia coli in 2000  and then confirmed in almost all eukaryotes . OXR1 plays an essential role in oxidative stress resistance [3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18,19,20], but not directly in reactive oxygen species (ROS) scavenging  nor possessing catalase or superoxide dismutase activity . It is involved in fundamental cellular and biological processes including DNA damage response [7, 8, 14], antioxidant pathways [14, 21], cell cycle [14, 15, 22], mitochondrial functions [8, 14, 16], glucose metabolism , S-nitrosylation , V-ATPase regulation , lysosomal functions , immune defense [25, 26], inflammatory response [18, 27], neuronal protection [3,4,5, 12, 17,18,19,20, 28], and life-span [13, 29]. It is associated with a wide range of diseases, such as amyotrophic lateral sclerosis(ALS) [3, 17,18,19], Parkinson's disease , retinopathy [30, 31], and Lupus nephritis , pulmonary arterial hypertension , etc. [21, 32, 33].
Due to the importance of ROS as signaling molecules in the brain, the high metabolic rate, and a relatively low antioxidant capacity, the brain is assumed to be particularly susceptible to an imbalance in ROS levels . Oxr1-depleted mice display progressive cerebellar ataxia, shortened lifespan, and confers neuronal sensitivity to oxidative stress [5, 12]. Overexpression of Oxr1 in variant ALS mouse models delayed the onset of neuromuscular phenotypes and neuroinflammatory responses and improved motor neuron functions [17,18,19]. A systemic analysis of mammal TLDc domain-containing proteins showed Oxr1 was highly expressed in the central nervous system of mice, and disruptions of the TLDc domain in Oxr1 caused neurodegeneration . In addition to the C-terminal TLDc domain, Oxr1 long isoforms also contain the evolutionarily conserved Tre2/Bub2/Cdc16 (TBC) domain and Lysin Motif (LysM) . However, the structural or functional roles of these domains remain elusive.
In 2019, it was reported that five patients from three families who carried four biallelic loss-of-function OXR1 variants were associated with cerebellar atrophy . The impact of OXR1 deficiency was further assessed in Drosophila models. Complete deletion of the mustard (mtd) gene, the fly homolog of human OXR1 and NCOA7, was lethal, which was rescued by the replacement of human OXR1 cDNA (TLDC domain only). In addition, neuron-specific depletion of mtd in flies led to an accumulation of aberrant lysosomal structures, massive neuronal loss, and early death . However, no human disease models are available to explore the pathological impact of OXR1 deficiency in the brain. The role of OXR1 in cellular functions and molecular mechanisms remains largely unknown.
Here, we characterized a novel and deleterious OXR1 mutation in patients with severe neurological features. We produced patient-derived lymphoblasts, fibroblasts, iPSCs, and used the latter to generate monolayer neuronal cell cultures and various brain organoid models (e.g., hippocampus, hypothalamus, midbrain, and forebrain) to systematically analyze key aspects of the pathological features and molecular mechanisms associated with OXR1 depletion in human. We found that the pathological phenotypes in patient lymphoblasts, such as impaired cell survival, proliferation, and hypersensitivity to oxidative stress, were rescued by the replacement of TLDc domain. In the patient derived iPSCs, impaired neural differentiation along with dysregulation of genes essential for neurodevelopment was identified. Further, we demonstrated that OXR1 influenced histone arginine methylation by activating protein arginine methyltransferases and revealed that OXR1 contributed to the spatial–temporal regulation of histone arginine methylation during human brain development.
Characterization of OXR1 deficiency and description of clinical phenotypes
Three sisters whose parents were first cousins (Fig. 1a) presented delay or loss of developmental milestones after birth or at 8–10 months of age accompanied with focal myoclonic movement, hypoactivity, cognition problems, and frequent onset of epilepsy. Upon examination, patients showed non-communicative, generalized hypotonia, hyporeflexia/areflexia, ataxia and lack of spontaneous movements (Table 1). Electroencephalogram examination revealed generalized frank epileptic activity. Magnetic resonance imaging (MRI) disclosed atrophy particularly in vermis and cerebellum, and thin corpus callosum (Fig. 1b). To identify the genetic lesion responsible for these clinical features, we first performed exome sequencing of patient II-6. Exome analyses revealed that the homozygous variant Chr8:107,758,095 G > C at the donor splice site located at exon 18 of the OXR1 gene (Hg19 chr8:g.107758095G > C, XM_005250993.1:c.2552 + 5G > C) segregated with the disease (Fig. 1c). This mutation was not present in any of the nearly 60,000 exomes deposited by the Exome Aggregation Consortium, Cambridge, MA (URL: http://exac.broadinstitute.org, accessed August 2016). The human OXR1 gene consists of 19 exons (ENSG00000164830, GRCh38), resulting in at least 4 major isoforms, OXR1A-D, with different tissue distributions. For instance, OXR1A and OXR1D are expressed in human brain , whereas most other cell types express isoforms B and D. All isoforms contain the C-terminal TLDc domain. Sequencing cDNA of patient II-4 fibroblasts, the flanking OXR1 exons 17–19 showed a shorter fragment (185 bp) compared to control cells (259 bp), indicative of homozygous skipping of exon 18 in OXR1 (Fig. 1d, Additional file 1: Fig. S1a,b), predicted to cause a frameshift leading to loss of the highly conserved C-terminal 69 amino acids of the TLDc domain of all OXR1 isoforms. Western blot of control cells showed a strong band corresponding to the OXR1B protein that was absent in the patient cells (Fig. 1e), indicating that lack of exon 18 leads to protein instability. Therefore, it appears that the OXR1 splicing mutation results in OXR1 depletion.
OXR1 deficiency compromises cell growth, survival, and response to oxidative stress in patient-derived lymphoblasts
To explore the functional impact of OXR1 deficiency in patients, we established lymphoblast cell lines (LB) from the patient (LB-OXR1ΔEx18) and one unaffected sibling as control (LB-CTRL). The OXR1 transcripts that were specific to the patients’ exon skipping sequence (Exon 18) were largely reduced in LB-OXR1ΔEx18 whereas the OXR1B transcripts increased about 1.7 fold (Additional file 1: Fig. S1c). The LB-OXR1ΔEx18 showed reduced proliferation (Fig. 2a), higher sensitivity to H2O2 with increased apoptosis and necrosis (Fig. 2b, Additional file 1: Fig. S2a). H2O2 treatment induced higher ROS levels in LB-OXR1ΔEx18 compared to control (Additional file 1: Fig. S2b). To determine the levels of oxidative DNA damage, we measured accumulation of 8-oxoguanine (8-oxoG) in genomic DNA . Basal levels of 8-oxoG were similar in patient and controls, but increased 8-oxoG levels were observed in LB-OXR1ΔEx18 upon H2O2 exposure (Fig. 2c). Furthermore, we determined the rate of mitochondrial DNA (mtDNA) mutagenesis , showing elevated mutation frequencies in the D-loop and 12S regions of mtDNA in LB-OXR1ΔEx18 (Additional file 1: Fig. S2c). We have previously shown that OXR1 upregulates the expression of antioxidant genes via the p21 signaling pathway in Hela cells . In this study, we observed a reduction of both p21 and heme oxygenase (HO-1) proteins in LB-OXR1ΔEx18 (Fig. 2d) and upregulation of stress response genes, including DUSP1, and the transcription factors (TF) FOS and JUN at the mRNA level (Fig. 2e, Additional file 1: Fig. S2d). ROS induced activation of caspases is pivotal in apoptosis. Caspase-9 mRNA levels (Fig. 2f), as well as the activated forms of Caspase-9 (35 and 37 kDa) were significantly increased in LB-OXR1ΔEx18 (Fig. 2g). Taken together, these results demonstrate that OXR1 deficiency reduces cell proliferation and increases intracellular oxidative stress, oxidative DNA damage and apoptosis.
The TLDc domain is essential for OXR1’s anti-oxidation function . OXR1D is predominantly composed of the TLDc domain and is expressed in all types of tissues in human. To validate whether the observed cellular phenotypes were due to the mutation in the TLDc domain, we performed rescue experiments using LB-OXR1ΔEx18 cells stably expressing OXR1D-GFP. Strikingly, expression of OXR1D-GFP was sufficient to increase the proliferation capacity of LB-OXR1ΔEx18 (Fig. 2h) and reduced apoptosis and necrosis to levels comparable to control LBs (Fig. 2i). These results suggest that the splicing site mutation in TLDc domain leads to depletion of OXR1 isoforms with loss-of-function.
Loss of OXR1 causes morphological and developmental defects in neuronal cells
To overcome the challenge due to the lack of access to patient brain tissues for understanding the pathological mechanisms, we generated iPSCs from fibroblasts of patient II-4 and a healthy control (AG05836). Loss of exon18 was conserved, and the OXR1 protein was absent in the patient iPSCs (Additional file 1: Fig. S3a,b). All control and OXR1ΔEx18 clones showed normal karyotypes, similar expression levels of pluripotency markers, and in vitro confirmation of pluripotency via directed differentiation to the three germ layers (Additional file 1: Fig. S3c-f, Additional file 2: Video S1). However, further differentiation of iPSCs into neurons revealed irregularities in the morphology of OXR1ΔEx18 neural aggregates (NAs) (Additional file 1: Fig. S3g,h). Moreover, OXR1-depletion reduced neurite growth (Fig. 3a), as well as the number of neurons, concomitant with reduced expression of neuronal markers, e.g., microtubule-associated protein 2 (MAP2) and class III beta-tubulin (Tuj1) (Additional file 1: Fig. S3i, j). Similar to patient LBs (Fig. 2b), OXR1-depleted neuronal cells displayed enhanced apoptosis (Additional file 1: Fig. S3k). Notably, OXR1 mRNA and protein levels increased during neural differentiation in control cells (Additional file 1: Fig. S3l). Taken together, these results demonstrate an essential role of OXR1 in neural differentiation.
OXR1 impacts transcription during neuronal differentiation
Previous work in our lab has demonstrated a role for OXR1 in transcriptional regulation [1, 15]. To elucidate OXR1-dependent regulation of transcriptional networks during neural differentiation, the transcriptome analysis was performed at three distinct stages during neuronal differentiation (iPSCs, NAs and neurons) of control and OXR1ΔEx18 iPSCs (Additional file 1: Fig. S3i, m-t). Transcriptomes of three clones of control iPSCs, NAs and neurons showed distinct clustering, suggesting a well-defined, stage-specific neural differentiation process (Fig. 3b). In contrast, no prominent clustering was observed between the three groups of OXR1ΔEx18 cells. Significantly, the differentially expressed gene (DEG) population in OXR1ΔEx18 cells (Padj < 0.05) had a steep increase of the median Log2FC (fold change) from the iPSC stage (median Log2FC = 0.8276, 95% CI 1.119–1.158), to the NA stage (median Log2FC = 1.069, 95% CI 1.541–1.607), and to the neuron stage (median Log2FC = 1.342, 95% CI 1.735–1.792) (Fig. 3c). With a threshold of Log2FC > ǀ1.5ǀ, almost 3.5-fold more DEGs were identified at the neuron stage (3793 DEGs, representing 12.8% of total genes) than at the initial iPSC stage (1089 DEGs, representing 3.5% of total genes) (Additional file 1: Fig. S3m). These results indicate that the impact of OXR1 on transcription is more pronounced at the neuron than the iPSC stage. Further, by searching published databases, we found that DEGs in patient-derived OXR1ΔEx18 neurons overlapped 39%, 41%, and 31% with disease-associated genes or high-confident risk genes of cerebellar atrophy, autism spectrum disorder, and schizophrenia, respectively (Table 2).
From iPSCs to neurons via NAs, lack of OXR1 lead to progressive failure in neuron-fate commitment (Additional file 1: Fig. S3n), suggesting impaired neuronal differentiation trajectories upon OXR1 depletion. Pathway enrichment by Gene Set Enrichment Analysis (GSEA) showed that the core pathways involved in neural development, such as Hedgehog, WNT, and TGF-beta, were activated in control, but not in OXR1ΔEx18 (Fig. 3d). Gene Ontology by GSEA revealed underrepresentation of biological processes required for the nervous system development, including hindbrain development (Fig. 3e, Additional file 1: Fig. S3p-q). Some overrepresented biological processes such as synaptic signaling and neurotransmitter transport were also identified (Additional file 1: Fig. S3o), indicating a potential role of OXR1 in synaptic regulation. Genes essential for neurogenesis and neurodifferentiation as well as axon guidance and cell migration were significantly down-regulated in Neu-OXR1ΔEx18 (Fig. 3f-g). In addition, we found that several pro-apoptotic genes were upregulated while anti-apoptotic genes were down-regulated in Neu-OXR1ΔEx18 (Additional file 1: Fig. S3r), potentially contributing to the increased death of neurons (Additional file 1:Fig. S3k). Several oxidative stress response genes were upregulated (Additional file 1: Fig. S3s), supporting the oxidative distress in OXR1-depleted neurons, as observed in LB-OXR1ΔEx18. According to a topologic biological network analysis of the dysregulated transcriptome of Neu-OXR1ΔEx18, we found that OXR1 orchestrated the expression of many key upstream regulators crucial for regulating transcription and substantial pathways for neural differentiation (Additional file 1: Fig. S3t). Further biological module clustering directly revealed OXR1-depletion impaired axon guidance at multiple levels, from axon genesis to neurite extension and synapse formation . Among them, the dysregulated Reelin pathway, cell adhesion molecules (ie. NRCAM), Class-3 semaphorin, and Slit/Robo pathways were highlighted, which correlated to the formation of a thinner corpus callosum [38,39,40].
OXR1 modulates PRMT-mediated histone modifications
Next, we explored molecular mechanisms underlying OXR1-dependent regulation of gene expression. Previous mass spectrometry analysis revealed the interaction of overexpressed OXR1-GFP in U2OS cells with protein arginine methyltransferase 5 (PRMT5) , an epigenetic regulator of gene expression. Here, we confirmed the interaction of endogenous OXR1 with PRMT5 as well as PRMT1 in U2OS cells by immunoprecipitation (IP) (Additional file 1: Fig. S4a), and in human fibroblasts (control vs OXR1ΔEx18) by an in-situ proximity ligation assay (Fig. 4a). PRMTs can activate or repress gene expression by catalyzing histone (H) arginine (R) methylations on the N-terminus of histones H3 and H4, e.g., H3R2, H3R17, and H4R3 . Crosstalk between histone arginine methylation and lysine acetylation/methylation has also been reported . We examined histone modifications during the differentiation of iPSCs to neural stem cells (NSCs) using an antibody-based screening and revealed increased histone methylations on H3R2/R17, H4R3, H3K4/K9/K27 in the control NSCs (Fig. 4b). In contrast, OXR1-depleted NSCs showed significantly impaired histone arginine/lysine methylations, e.g., H3R2me2a, H3R17me2s/2a, H4R3me2s/2a, H3K4me3, H3K9me3, and H3K27me3. However, we did not detect any changes in the expression of PRMTs in either OXR1ΔEx18 iPSCs or NSCs as compared to control cells (Additional file 1: Fig. S4b). Thus, our results suggest that OXR1 may mainly affect the catalytic activity of PRMT-induced epigenetic histone modifications during neurogenesis.
To gain more insight into how OXR1 regulates histone marks, we integrated our RNA-sequencing data with published ChIP-sequencing datasets using the standard quality control available from http://cistrome.org/db. As shown in Fig. 4c, genes enriched with the repressive mark H4R3me2s overlapped with 71% of the down-regulated genes during neural differentiation in control cells (1319 out of 1869 genes), of which 63% of the genes were differentially expressed (up-regulated) in OXR1ΔEx18 cells (802 out of 1266 genes). Furthermore, genes enriched with H3K4me3, H3K9me3, H3K27me3, and H3K27ac overlapped with 16%-27% of the down- or up-regulated DEGs in OXR1ΔEx18 cells. These data may suggest a role of OXR1 in the regulation of histone modifications, particularly H4R3me2s.
To further explore a functional role of OXR1 in PRMT-catalyzed methylation of H4R3, we assessed the activity of the PRMT5/MEP50 methyltransferase on the Histone 4 N-terminal tail (aa1–21) in a methyltransferase assay in vitro by adding to the reactions full-length hOXR1A or mutant proteins with partial or complete deletion of TLDc (Fig. 4d). Interestingly, full-length hOXR1A and the mutants with partially truncated TLDc (OXR1A_∆TLDc_1) stimulated PRMT5-catalyzed H4 methylation by threefold. In contrast, the OXR1A mutant lacking TLDc (OXR1A_ΔTLDc_2) failed to stimulate PRMT5 activity, suggesting the TLDc domain of OXR1 may have a role in the regulation of PRMT-catalyzed H4R3 methylation.
Impaired development of OXR1 deficient human brain organoids
To unravel pathological processes at molecular, cellular, and structural levels, we modeled OXR1 deficiency in the developing human brain organoids. We established cerebral organoids , and various brain region-specific organoids [44, 45]derived from patient iPSCs. Cerebral organoids often contain heterogeneous tissues resembling various brain regions, which relies on the intrinsic differentiation signaling and self-organization capacities of iPSCs (Fig. 5a). Significantly, OXR1ΔEx18 cerebral organoids were smaller than the healthy controls (CTRL) with slower growth rate and suboptimal morphology of neuroepithelial buds (Fig. 5a). After about 30 days, the cortical regions in healthy controls displayed dense and radially organized ventricular /subventricular zone (VZ/SVZ) like structures harboring the neural progenitor cells (NPCs). The neuronal layer composed of TBR1+ and CTIP2+ deep-layer neurons was highly reminiscent of the formation of cortical plate (CP) (Additional file 1: Fig. S5a). New-born neurons continued to migrate radially outward, became mature (MAP2+) and obtained rudimentary separations during the lamination process (Fig. 5b). However, in OXR1ΔEx18 organoids, the two layers still intermingled at day 75 (Fig. 5b; Additional file 1: Fig. S5b), indicating either delayed neurogenesis or impaired neuron migration, or a combination. In addition, we observed a thick Reelin-positive (RELN+) layer along the basal surface reminiscent of the marginal zone with long and extended RELN+ plexus at the inner layer in control organoids at day75 (Fig. 5c; Additional file 1: Fig. S5c). It is known that RELN expressed at the marginal zone is essential for correct cortical and cerebellar neuronal lamination , and RELN localization along dendrites suggests a role in synaptic remodelling and neuronal maturation . However, thinner marginal zone with reduced RELN deposition, shorten axons and lack of RELN+ plexus was observed in patient iPSC-derived organoids. These results suggest that OXR1 deficiency leads to delayed neuronal layering in human cerebra.
OXR1ΔEx18 patients showed a global neurodevelopmental delay characterized by the dysfunctions of cognition, fine motor skills, learning and language, suggesting impaired functions not only in cerebellum, but also in cortical (i.e., prefrontal) areas and deep structures (i.e., hippocampus, basal ganglia). To assess the importance of OXR1 in distinct brain regions, we guided neural progenitor cells (NPCs) to form brain region-specific identities by supplement of highly tailored external patterning factors at the early stage (Fig. 5d-h, 6a-c, 7a-d) [44, 48, 49]. From control iPSCs, we successfully generated hypothalamus, hippocampus, forebrain (cortical) and midbrain organoids with large VZ-like structures containing the region-specific NPCs. Molecular markers for cell identities were analyzed by IHC of different region-specific organoids. OXR1 is highly expressed in both POMC+ peptidergic neurons and OTP+ hypothalamic neurons of hypothalamus organoids at day 50 (Additional file 1: Fig. S5d). Strikingly, OXR1-deficient clones from patient iPSCs lost the ability to form hypothalamus organoids (Fig. 5e). In addition, iPSC-OXR1ΔEx18 generated hippocampal organoids from only one clone (Fig. 5f) with a reduced size of SOX2-positive progenitor zone, a less organized layer of DCX-positive newborn neurons, and fewer NeuN-positive mature neurons (Fig. 5g-h; Additional file 1: Fig. S5e-f). Taken together, these results suggest a crucial role of OXR1 in early hypothalamus and hippocampal development.
OXR1 impacts early cortical development by shaping histone arginine methylations
We successfully generated cortical (forebrain) organoids from control and OXR1ΔEx18 iPSCs (Fig. 6a, Additional file 1: Fig. S6a). Control cortical organoids contained the large and radially organized VZ/SVZ like structures marked by SOX2 (Fig. 6b; Additional file 1: Fig. S6b), where cortical NPCs underwent active cell division (marked by Ki-67, in Fig. 6c). The newly generated layer of CTIP2+ early born neurons extended the formation of CP resembling human early corticogenesis (Fig. 6b). Interestingly, we identified a specific distribution of the histone arginine methylation mark H4R3me2s, which showed low levels in VZ/SVZ (including outer SVZ) like structures and increased levels in developing neurons outside the progenitor zone following the radial migration (Fig. 6d). This finding suggests a regulatory role of H4R3me2s during neurogenesis and the establishment of the cortical plate. However, the specific H4R3me2s distribution was affected in OXR1ΔEx18 cortical organoids, showing an overall reduction of the H4R3me2s level (Fig. 6d). In addition, OXR1ΔEx18 cortical organoids showed a thin and poorly organized VZ/SVZ with markedly reduced number of Ki67+ proliferating NPCs, and a barely developed CTIP2+ cortical plate (Fig. 6b, c). H3R2me2s, another target of PRMT5, displayed an even distribution pattern across the normal laminar cortical structures of control and OXR1ΔEx18. However, an overall reduction of the H3R2me2s level was detected in OXR1ΔEx18 cortical organoids (Fig. 6e). These results suggest that OXR1 impacts the early human corticogenesis by shaping histone arginine methylations.
OXR1 impacts early midbrain development by shaping histone arginine methylations
In addition, we generated midbrain organoids from control and OXR1ΔEx18 iPSCs (Fig. 7a). Midbrain neurogenesis starts at VZ of the midbrain floor plate (mFP) where midbrain progenitors are differentiated into various types of tyrosine hydroxylase (TH+) midbrain dopaminergic (mDA) cells (Additional file 1: Fig. S6c). We guided midbrain NPCs towards the identity of substantia nigra pars compacta (SNc) mDA neurons via the Shh-FOXA1/2-SOX6 pathway by using SHH, CHIR99021 and purmorphamine as patterning factors [37, 44, 50,51,52,53,54,55,56]. As expected, FOXA2+ progenitors were broadly detected at day 20, migrated through the intermediate zone (IZ) and became TH+ mDA when reaching mantle zone (MZ) at day 50 (Fig. 7b). In OXR1ΔEx18 midbrain organoids, FOXA2+ progenitors were sparsely detected at day 20 (Fig. 7b) with fewer Ki67+ proliferating midbrain NPCs (Fig. 7c), although the structural organization of VZ remained largely intact. In contrast, we observed an abnormal increase of TH+ mDA generated at day 50 (Fig. 7b), suggesting dysregulation of midbrain NPCs specification derived from the activation of FOXA2. It is known that upregulation of EN2 by FGF8 in the midbrain-hindbrain boundary is essential for further cerebellar development [57,58,59]. Similarly, EN2 was induced in VZ in control organoids, but absent in OXR1ΔEx18 (Fig. 7d), suggesting a potential mechanism underlying the cerebellar defects in patients. Notably, the distribution pattern of H4R3me2s in the control midbrain organoids was distinct from the cortical organoids by showing a high level in the VZ progenitor zone (Fig. 7e). Nevertheless, the level of H4R3me2s was dramatically decreased in OXR1ΔEx18 organoids. We did not detect any significant changes of H3R2me2a or H3R2me2s levels in OXR1ΔEx18 midbrain organoids (Additional file 1: Fig. S6d,e). But the distribution pattern and the expression level of histone lysine mark H3K4me3 were severely dysregulated in OXR1ΔEx18 midbrain organoids (Additional file 1: Fig. S6f). Taken together, these results suggest that OXR1 modulates distinct histone marks during early midbrain development, accounting for the clinical phenotypes in OXR1ΔEx18 patients.
In this work, advanced human cell culture systems modeling OXR1 depletion provide new insight into the molecular and cellular pathology of rare diseases associated with inherited mutations of the TLDc containing family. We reported a de novo mutation within the critical TLDc domain of OXR1 leading to exon skipping, protein depletion (OXR1ΔEx18) and a severe early-onset syndrome characterized by epilepsy, global developmental delay, cognitive disabilities, and cerebellar atrophy. Using 2D neural differentiation and various 3D brain organoid models derived from patient iPSCs, we identified specific neurodevelopmental defects due to OXR1 depletion, including impairment of NPC proliferation, fate-specification, neurogenesis, neurite outgrowth, and neuronal migration. We revealed a role of OXR1 in epigenetic regulation of early human brain development by modulating PRMT-catalyzed histone arginine methylation.
The highly conserved TLDc domain is common for six different proteins (e.g., TBC1D24 and NCOA7), which all appear to display neuroprotective roles according to recent studies [3, 60]. A mutation in TBC1D24 was the first genetic cause of human epilepsy found . Later, more than 50 mutations in TBC1D24 have been associated with various types of drug-resistant epilepsy, intellectual disability, and DOORS (deafness, onychodystrophy, osteodystrophy, mental retardation, and seizures) syndrome [62, 63]. Most pathogenic mutations of TBC1D24 are in the TBC domain and around 12 mutations are located in TLDc domain. Depletion of TBC1D24 by the acute silencing or mutation leads to defects in axon formation and maturation, and neuronal radial migration [64, 65], consistent with our finding that the OXR1 deficiency impaired axon development (Figs. 3g, S3j, 5b, c). Interestingly, acute silencing of TBC1D24 in vivo showed reduced corpus callosum projections in rats , which was one of MRI signs identified in our OXR1 deficient patients.
Previous studies show that loss of OXR1 in various animal models and cell lines leads to neurodegeneration [3,4,5, 12, 17,18,19,20, 28]. Most of these studies demonstrate that the TLDc domain is essential for protection of neurons against oxidative and nitrosative stress-induced cell death, e.g. in mice cerebellum [3, 5, 12, 19, 23]. Other studies suggest that OXR1 could regulate glucose metabolism in mice cerebellum  or lysosomal structure and function in Drosophila neurons . Here, we found elevated oxidative stress and increased cell apoptosis in human OXR1ΔEx18 patient derived lymphoblasts and 2D differentiated neurons (Additional file 1: Fig. S3i, k), which are the first human cell based models with minimal artificial intervention showing phenotypes due to OXR1 deficiency. In addition, we performed the deeper module analysis of enriched biological processes and identified the impaired mitochondrial function, glycolysis and autophagy in OXR1ΔEx18 neurons (Additional file 1: Fig. S3t). Oxidative eustress, also known as redox homeostasis is crucial for brain development . We identified several key modulators of redox homeostasis that were significantly affected in Neu-OXR1ΔEx18, including H2O2 scavengers, key redox-sensitive targets and redox hubs (Additional file 1: Fig. S3s,t), suggesting an important role of OXR1 not only in maintaining the balance between oxidative eustress and distress, but also in the proper redox signaling required for neuron development [66,67,68,69].
According to BrainSpan Atlas of the developing human brain , OXR1 expression increases largely from the early second trimester, reaching a plateau at two years of age until brain development is completed. OXR1 is highly expressed in neuronal cell types and is also detected in oligodendrocyte progenitor cells and microglia across human adult cortex (cited in http://celltypes.brain-map.org/rnaseq). Similarly, we detected an increased level of OXR1 during neural differentiation (Additional file 1: Fig. S3l). Both OXR1 deficient patients  and OXR1 knockout mouse models [5, 12] showed cerebella related symptoms after birth. In lack of the access to viable human brain tissues, human brain organoids are the only models enabling advanced studies of brain development and patho-mechanisms. Importantly, brain organoids exhibit more complex cytoarchitecture over time including the organization of progenitor cells and sequential generation lamination of neurons. By using the 2D neural differentiation model, we found that OXR1-deficiency reduced neurite growth (Fig. 3a) as well as the expression of neuronal markers, e.g., MAP2 and Tuj1 (Additional file 1: Fig. S3i, j), suggesting an important role of OXR1 in neurodifferentiation. Taking the advantage of brain organoids in recapitulating the complexity of early corticogenesis, we revealed that OXR1 deficiency resulted in delay of specific cellular programs, including NPC proliferation, CP formation, cortical layering, and axon/dendrites growth. Cortical organoids at around day 50 coincide with the post-conception week (PCW) 14–16 in humans . Impaired region-specific NPC proliferation and neurogenesis have been closely linked to primary microcephaly and vermis/cerebellar hypoplasia [72, 73]. We tried the protocol of cerebellum organoids described by the Sasai group . Unfortunately, those organoids dissociated at an early stage, which limited our study of OXR1´s roles in the cerebellar progenitor cell development and the maturation processes of cerebellar neuron subtypes. Efficient neural induction, especially the application of region-specific patterning factors, restricts the generation of non-neuroectodermal cells (e.g., microglial, endothelial, and meningeal cells) that are critical to the development and function of the human brain. Recently, there have been significant advancements and attempts to improve organoid longevity and maturity, enhance glial cell production, reconstitute microglia and vasculatures, and fuse multiple region-specific organoids. We believe that future improvements will allow organoids to faithfully recapitulate late gestational and early post-natal development. In addition, the current study is limited due to the use of unrelated healthy iPS cells as control. Although we particularly chose a female control to minimize gender affection, these control cells share some differences in age, genetic background, and other factors with the patient iPS cells. In order to precisely generate human rare disease models, illustrate translational relevance, and develop new therapeutic approaches, an isogenic control with the same genetic background is very important in iPS-derived disease modeling. For example, direct correction of OXR1 genetic mutation in patient iPS cells, using genome editing technology, is warranted in future studies.
Genes essential for neurogenesis, neurodifferentiation, axon guidance, and cell migration were significantly down-regulated in Neu-OXR1ΔEx18 (Fig. 3f-g). Consistently, most altered factors in the OXR1-deficient organoids showed a similar trend of differential expression in OXR1-deficient neurons of the 2D model. For example, down-regulation of MAP2 (Log2FC = -3.00) and RELN (Log2FC = -5.36) was identified in OXR1ΔEx18 neurons (Padj < 0.05) as detected in OXR1ΔEx18 cerebral organoids (Fig. 5b, c). Reduced expression of TBR1 was observed in OXR1ΔEx18 cerebral organoids (Fig. 5b), but the expression of TBR1 was barely detected in the 2D neural differentiation model. Further, the expression level of FOXA2 (Log2FC = -7.32) and EN2 (Log2FC = -7.80) was strikingly decreased in OXR1ΔEx18 neurons, which is consistent with the sparse detection of FOXA2-positive progenitors and failed induction of EN2 in OXR1ΔEx18 midbrain organoids (Fig. 7b, d). Although the level of tyrosine hydroxylase (TH) showed a significant decrease (Log2FC = -2.83) in OXR1ΔEx18 neurons (the 2D neural differentiation model), the guided OXR1ΔEx18 midbrain dopaminergic (mDA) cells showed an abnormal increase of TH at day 50. Of note, the 2D neural differentiation model and the guided 3D brain organoid model represent distinct processes of neurodevelopment. Our work highlights the important role of OXR1 in shaping transcription during early neurodevelopment.
Arginine methylation, a widespread post-translational modification catalyzed by PRMTs, plays key roles in epigenetic regulation and signal transduction influencing gene transcription, RNA metabolism and DNA damage response. Dysregulation of arginine methylations has been linked to pathomechanisms of several diseases, including neurodegeneration (i.e., PRMT1-ALS, PRMT5-PD), and muscular disorders [17, 41, 75, 76]. PRMT5-mediated H4R3me2 is involved in regulating proliferative neuroepithelium/neural progenitors in mouse cortex . Conditional depletion of PRMT5 in the mouse CNS (from E10.5) results in early postnatal lethality with severe developmental defects, e.g., depletion of the external granular layer of the cerebellum, reduced thickness of the cortex with decreased neuronal cells in CP and SOX2/Ki67-positive proliferating NPCs in VZ/SVZ . In this study, we used various human cell culture systems and revealed the regulation of histone arginine methylation by the interaction of OXR1 and PRMT5 in human brain development. We provided direct evidence that OXR1 promoted PRMT5-mediated histone arginine (e.g., H4R3) methylations (Fig. 4d). In 2D derived NSC-OXR1ΔEx18, OXR1 deficiency led to dysregulation of multiple histone arginine methylations (e.g., H4R3me2s/2a, H3R2me2a, H3R17me2s/2a, H3K4me3, H3K9me3, and H3K27me3) (Fig. 4b), without affecting the levels of different PRMTs (Additional file 1: Fig. S4b). Strikingly, the H4R3me2s associated genes overlapped with 71% of the down-regulated genes in the normal neural differentiation, of which 63% of the genes were differentially expressed (up-regulated) in NSC-OXR1∆Ex18 (Fig. 4c). Interestingly, H4R3me2s showed a specific distribution in the cortical brain organoids with a low level in VZ/SVZ like structures and increased levels in developing neurons outside the progenitor zone (Fig. 6d), suggesting a regulatory role of H4R3me2s during neurogenesis and the establishment of the cortical plate. However, the distribution of H4R3me2s in the midbrain organoids was distinct from the cortical brain organoids, showing a high level in the VZ progenitor zone (Fig. 7e). Notably, dysregulation of H4R3me2s was identified in OXR1ΔEx18 cortical and midbrain organoids, (Figs. 6d and 7e). It has been demonstrated that brain organoids possess comparable transcriptomic profiles to specific stages of fetal brains, as well as similar regulation of epigenetic programs, e.g., histone modifications [79, 80]. Our work reveals that OXR1 impacts histone arginine modifications in the early stage of human cortical and midbrain development, suggesting an important molecular patho-mechanism underlying the neurodevelopmental defects in OXR1ΔEx18 patients.
Altogether, we presented the first evidence of molecular neuropathology underlying OXR1 deficiency associated cerebellar atrophy, neurodevelopmental impairment, and increased susceptibility to oxidative stress. We provided new insights into the roles of OXR1 in cell survival and neural differentiation. We used brain organoids to recapitulate the early brain developmental defects of OXR1ΔEx18 patients in the context of brain region-specific cyto-architectures, including impaired NPC proliferation, neuronal fate specification, axon formation and neuronal migration. We revealed an important role of OXR1 in the epigenetic regulation of early human brain development by modulating PRMT-catalyzed histone arginine methylations. This study provides new insights into pathological features and molecular underpinnings associated with OXR1 deficiency in patients.
Whole exome analysis
Exonic sequences were enriched in the DNA sample of patient II-6 using SureSelect Human All Exon 50 Mb Kit (Agilent Technologies, Santa Clara, California, USA). Sequences were determined by HiSeq2000 (Illumina, San Diego, California, USA) as 100-bp paired-end runs. Data analysis including read alignment and variant calling was performed by DNAnexus software (Palo Alto, California, USA) using the default parameters with the human genome assembly hg19 (GRCh37) as reference.
Immortalized lymphoblast cell (LB) lines were generated from patient II-4’s and healthy control’s peripheral blood mononuclear cells transformed with Epstein-Barr virus (EBV) as described by Bernacki SH et al. , in the National Laboratory for the Genetics of Israeli Populations (NLGIP). The cells were cultured at 37 °C with 5% CO2 in growth medium: RPMI 1640 medium with GlutaMAX™ (Gibco, Cat.61870–010), supplemented with 20% fetal bovine serum (FBS), 100 IU/mL penicillin and 100 μg/mL streptomycin. Fibroblast cells and U2OS were cultured at 37 °C with 5% CO2 in growth medium: DMEM medium (Sigma-Aldrich, D6429) supplemented with 10% FBS. The trypan blue (0.4%, Invitrogen™) exclusion method is used to determine the number of viable cells present in a cell suspension. Cell counting was employed on the Countess® automated cell counter (Invitrogen™).
Plasmids and electroporation and cell sorting
The human cDNA of OXR1A ORF (NM_001198533) and the truncated variants of OXR1A, OXR1A ∆TLDc_1 (lacking 69 aa from the C-terminal) and OXR1A ∆TLDc_2 (lacking the TLDc domain) were synthesized by GeneScript. They were inserted into plasmid pET28b and fused with the Hisx6 Tag between the Nde I and BamH I sites to obtain pET28b-OXR1A and pET28b-OXR1A ∆TLDc plasmids. All the plasmids were verified by sequencing. The plasmids of pET28b-OXR1A, OXR1A_∆TLDc_1 and OXR1A_∆TLDc_2 were transformed into E. coli and further used for the purification of recombinant hOXR1 proteins by Ni–NTA and gel filtration columns.
Full length cDNA (NM_001198534) representing human OXR1D, synthesized by RT-PCR with mRNA from human HeLa cells, was clone in vector pEGFP-N1 and pEGFP-C1 with the GFP Tag between the Xho I and BamH I sites, to obtain pEGFP-N-OXR1D plasmids and pEGFP-C-OXR1D, respectively.
The pEGFP-C-OXR1D (pE-OXR1D) and The pEGFP-N2 (Vector) were transfected into patients’ LB (OXR1ΔEx18) by electroporation using a method modified from the patent (CN 103981218 A) (Wenting Hou, et al. Department of Biochemistry and Molecular Biology Peking University Health Science Center). LB-OXR1ΔEx18 in log phase growth were counted and harvested by centrifugation at 800 × g for 10 min at room temperature. After two washes with RPMI 1640 containing 10% FBS, the cells were resuspended in 37 °C RPMI 1640 (with 10% FBS) at 1 × 107 cells/mL. 250 μL of cell suspension and 15 μg of pEGFP-c-OXR1D or 12.5 μg of pEGFP-N2 (Vector) were combined in a BTX 0.4 cm electroporation cuvette. Electroporation was done with ECM-830 (BTX®) at 210 V/pulse length = 30 ms/pulse = 1. Electroporated cells were transferred to 2 mL of pre-warmed culture medium (with 10% FBS) at 37 °C. The electroporated cells were allowed to grow in 5% CO2 at 37 °C for 48 h(h) before cell sorting. Fluorescence-activated cell sorting (FACS)of EGFP-labeled LB were accomplished by FacsAria IIu high speed sorter at Flow Cytometry Core Facility, Oslo university hospital. The sorted cells were cultured at 37 °C with 5% CO2 in growth medium for two weeks. The expression level of OXR1 was tested by real time qPCR.
RNA isolation, cDNA sequence and gene expression analysis by quantitative real-time polymerase chain reaction
Total RNA was isolated from patient and control samples using the RNeasy Kit (Qiagen) following the manufacture’s protocol. The quality of purified RNA was estimated on an Epoch Microplate Spectrophotometer (Bio-Tek). Sequencing of OXR1 cDNA from lymphocytes cDNA was prepared from fresh lymphocytes of patient II-4 and control using Maxima™ 1st strand cDNA synthesis kit (Thermo Scientific) and amplified by PCR using Phusion High-Fidelity PCR Master Mix (Thermo Scientific). The PCR products were separated using 2% (w/v) agarose gel electrophoresis and their sequence determined by Sanger sequencing. Two-steps qPCR method was used to determine mRNA levels. First, the cDNA was generated from total RNA samples using the High-capacity cDNA reverse transcription kit (Applied Biosystem). Real-time PCR reactions were prepared using Power SYBR Green Master mix including about 3 ng cDNA in each reaction, and qPCR was performed in the StepOnePlusTM Real-Time PCR System (Applied Biosystem) with the standard cycle conditions: 95 oC for 10 min(min); 40 cycles at 94 °C for 15 s(s) and 65 °C for 30 s. Samples were measured in triplicates. ACTIN-ß and GAPDH were used as internal standard, as indicated. Quantification was done using the comparative cycle threshold (CT) method of relative quantification (RQ), where RQ is the normalized signal level in a sample relative to the normalized signal level in the corresponding calibrator. The CT for the target gene was normalized to the CT for GAPDH or ACTIN-ß in the same sample, giving ΔCT sample. The average ΔCT from cells’ cDNA was used as the ΔCT calibrator for the calculation of ΔΔCT. RQ is defined as 2 − ΔΔCT, where ΔΔCT = ΔCT sample – ΔCT calibrator. Primers used to amplify target genes by qPCR are as follows:
HO-1 F (5’- AACTTTCAGAAGGGCCAGGT-3’), R (5’- CTGGGCTCTCCTTGTTGC-3’); NRF2 F (5’- ACACGGTCCACAGCTCATC-3’), R (5’-TGTCAATCAAATCCATGTCCTG-3’);
P21 F (5’- GGCGGGCTGCATCCA -3’), R (5’- AGTGGTGTCTCGGTGACAAAGTC-3’);
OXR1 F (5’-TTATGGTACTGGAGAGACCTTTGTTTT-3’),
R (5’- AAAACATATTATCTCCTGTCCACTTAAAGAC -3’);
FOS F (5’- GGGGCAAGGTGGAACAGTTA-3’), R (5’- GTCTGTCTCCGCTTGGAGTG -3’);
c-JUN F (5’-GTGCCGAAAAAGGAAGCTGG-3’), R (5’- CTGCGTTAGCATGAGTTGGC-3’);
DUSP1 F (5’- GGATACGAAGCGTTTTCGGC-3’), R (5’ -GGCCACCCTGATCGTAGAGT-3’);
CASP9 F (5’- TCTACGGCACAGATGGATGC -3’), R (5’- CATGGTCTTTCTGCTCCCCA -3’);
NANOG F (5’- AATACCTCAGCCTCCAGCAGATG -3’),
R (5’- TGCGTCACACCATTGCTATTCTTC -3’);
OCT4 F (5’- GTACTCCTCGGTCCCTTTCC -3’), R (5’- CAAAAACCCTGGCACAAACT -3’);
SOX2 F (5’- GGATAAGTACACGCTGCCCG -3’), R (5’- ATGTGCGCGTAACTGTCCAT -3’);
ACTIN F (5’- GTTACAGGAAGTCCCTTGCCATCC -3’),
R (5’- CACCTCCCCTGTGTGGACTTGGG -3’);
GAPDH F (5’- CCACATCGCTCAGACACCAT -3’), R (5’- GCGCCCAATACGACCAAAT -3’).
Cell pellets were prepared using RIPA lysis buffer (150 mM NaCl, 50 mM Tris pH 7.5, 1 mM EDTA, 0.1% SDS, 0.5% Sodium deoxycholate, and 1% Triton X-100, 1 mM DTT and 1 × Protease Inhibitor Cocktail). The soluble protein extracts were separated by SDS-PAGE (4–15% mini-protein TGX precast gel, 12 or 15 well, BioRad) and transferred to a PVDF membrane (Trans-blot Turbo mini-Format 0.2 μM PVDF, cat.170–4156 by BIO-RAD). The membrane was blocked with 5% milk in PBS-T buffer (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2PO4, 1.4 mM KH2PO4, 0.1% Tween-20) for 4 h at room temperature following incubation with the indicated primary antibodies, overnight at 4 °C. For detection, membranes were incubated with secondary antibodies diluted in PBS-T buffer, except anti-GAPDH-HRP 1:5000 (Abcam Cat# ab9482, RRID:AB_307272). After washing 3 times with PBS-T buffer, the membrane was developed with ECL substrate (Super Signal West Fem to Maximum Sensitivity Substrate, by Thermo Scientific). The signals were detected by a BIO-RAD Imager and analyzed by Image Lab software. A list of antibodies used is provided in Additional file 3: Table S1.
Cell viability and apoptosis assay
The LB cells were plated in 6-well plates at 1 × 106 cells/mL overnight. Then the cells were exposed to the indicated concentrations of H2O2 in fresh medium for 1 h and recovered for 24 h. After treatment, the cells were stained with propidium iodide (PI) and annexin V using Alexa Fluor 488 annexin V/Dead cell apoptosis kit (Invitrogen, Cat.V13245) according to the manufacture’s instruction. Analysis was performed via flow cytometry with BD Accuri C6 Sample software. At least 10,000 cells were analyzed for each sample.
Apoptotic cell death was determined by In Situ cell death detection kit (Merck, Cat#11,684,795,910) according to the manufacturer’s instructions. Briefly, cells were fixed with 4% paraformaldehyde and permeabilized with 0.1% Triton X-100 and 0.1% sodium citrate for 15 min. After washing, 100 μL of TUNEL reaction mixture was added and incubated for 1 h at 37 °C. The cells were counterstained with 1 μg/mL DAPI (4′,6′-diamidino-2-phenylindole). The positive cells were imaged using an EVOS fluorescence microscope and analyzed using ImageJ software.
Cellular general ROS measurement
The LBs in 6-well plates at 1 × 106 cells/mL were incubated with 10 μM ROS probe CM-H2DCFDA (chloromethyl dichlorodihydrofluorescein diacetate, acetyl ester, Invitrogen, cat. 6827) in phosphate buffered saline (PBS) at 37°C for 30 min, washed once by PBS and recovered in phenol red free medium for 20 min. Next, the cells were exposed to 0.75 mM H2O2 in phenol red free medium for 1 h. Then, the cells were spin down and washed once with PBS, re-suspended in 0.5 mL cold PBS and kept on ice in dark. The fluorescence was measured using a BD Accuri™ C6 Cytometer.
HPLC–MS/MS analysis of 8-oxo(dG) in genomic DNA
LBs were plated in 6-well plates at 1 × 106 cells/mL overnight, and then exposed to the indicated concentrations of H2O2 in fresh medium and harvested after 1 h, 4 h and 6 h treatment. Total DNA was isolated from LB using the DNeasy Kit (Qiagen) according to the manufacturer’s instructions. This assay consists of two independent experiments with technical triplicates (the first assay) or sextuplicates (the second assay).
For mass spectrometry analysis, genomic DNA was enzymatically hydrolyzed to deoxyribonucleosides by incubation for 40 min at 40 °C in a mixture of benzonase (Santa Cruz Biotechnology, sc-391121B), nuclease P1 from Penicillium citrinum (Sigma, N8630), and alkaline phosphatase from E. coli (Sigma Aldrich, P5931) in 10 mM ammonium acetate buffer pH 6.0 with 1 mM magnesium chloride. Proteineous contaminants were precipitated by addition of three volume equivalents of ice-cold acetonitrile and centrifugation at 16,000 g at 4°C for 40 min. Supernatants were collected and vacuum-dried at room temperature. The resulting residues were dissolved in water for HPLC–MS/MS quantification.
Chromatographic separation and mass spectrometry detection of 8-oxo(dG) was performed at room temperature using an Agilent 1290 Infinity II UHPLC system coupled with an Agilent 6495 Triple Quad (Agilent Technologies, Germany) and an Eclipse Plus C18 1.8 µm 150 × 2.1 mm i.d. column equipped with a guard column. The mobile phases used were: A (0.1% formic acid in water), and B (0.1% formic acid in methanol). Chromatographic separation started with 95% A and 5% B for 2.5 min; followed by linear gradients of: 0.5 min 5–13% B, 2.5 min 13–17% B, 1.5 min 45–65% B, 1.5 min 65–70% B, 1 min 70–5% B; 4 min re-equilibration with 5% B.
Chromatographic separation of unmodified nucleosides was performed using a Shimadzu Prominence LC-20AD HPLC system with an Ascentis Express C18 2.7 µm 150 × 2.1 mm i.d. column equipped with an Ascentis Express Cartridge Guard Column (Supelco Analytical, Bellefonte, PA, USA) with EXP Titanium Hybrid Ferrule (Optimize Technologies Inc.). Chromatographic separation was performed with isocratic flow of 25% methanol in water with 0.1% formic acid, 0.16 ml/min at 40 °C. Online mass spectrometry detection of unmodified nucleosides was performed using an Applied Biosystems/MDS Sciex API5000 Triple quadrupole mass spectrometer (ABsciex, Toronto, Canada).
The deoxyribonucleosides were monitored by multiple reaction monitoring in positive electrospray ionization mode, using following mass transitions: 252.1 → 136.1 (dA), 228.1/112.1 (dC), 268.1/152.1 (dG), 243.1/127.0 (dT), 284.1/168.0 (8-oxodG).
mtDNA mutation frequency measurement
Total DNA containing nuclear DNA and mtDNA were isolated from LB using the DNeasy Kit (Qiagen) according to the manufacturer’s instructions. Double stranded mtDNA mutation frequency was estimated as described previously . Total DNA was digested with S1 nuclease (10U, Qiagen) for 15 min at 37 °C and subsequently digested with TaqI restriction enzyme (100U, New England Biolabs) at 65 °C for 1 h to remove single strands, damaged DNA and non-mutated DNA. The remaining mutated TaqI restriction fragments were quantified in a qPCR reaction. To ensure complete digestion of non-mutated TaqI restriction sites, TaqI was added to qPCR reaction mixture (100 U) and an additional step of 65 °C for 15 min is included prior to the standard qPCR program. Mutation frequency is calculated as (2exp(CT_TaqI – CT_NT)*4) − 1 per nt. The primer pairs for each gene were 12S F (5’- AAACTGCTCGCCAGAACACT -3’) and R (5’- CATGGGCTACACCTTGACCT -3’); D loop F (5’- CCCGGAGCGAGGAGAGTAG -3’) and R (5’- CACCATCCTCCGTGAAATCAA -3’).
Generation of human induced pluripotent stem cells (hiPSCs)
Fibroblasts from skin biopsy of patient II-2 and healthy control AG05836 (Coriell Institute) were cultured at 37 °C with 5% CO2 in growth medium: DMEM medium, supplemented with 10% fetal calf serum, 100 IU/mL penicillin and 100 μg/mL streptomycin. iPSCs were generated with CytoTune®-iPS 2.0 Sendai Reprogramming Kit (ThermoFisher Scientific, Cat#A16517) using a multiplicity of infection of 5:5:3 (KOS:c-Myc:Klf4). Individual colonies with typical morphology of human embryonic stem cells were manually picked around 3–4 weeks after infection. The iPSCs were continuously cultured and expanded on Geltrex coated 6-well plates with Essential 8 medium (ThermoFisher Scientific, Cat#A1517001). Human iPSC cell lines used in this manuscript: AG05836 clone 1, 10, 15, 27 as healthy controls and OXR1 clone 2, 10, and 11.
Neural differentiation of hiPSCs
Differentiation of hiPSC towards neural lineage was performed as previously reported with modifications . Briefly, iPSCs were pretreated with 50 μM Y-27632 (Stemcell Technologies) for 1 h. Then the cells were dissociated with Accutase (Stemcell Technologies) and plated onto low attachment V-type 96-well plate at 20 000 cells per well or AggreWell800 plate (Stemcell Technologies) to form aggregates. The culture was maintained in neural induction medium consisting of DMEM/F12 supplemented with 2% B27 supplement without vitamin A, 1% N2 supplement, 200 nM Noggin (R&D Systems) and 10 μM Y-27632 for 48 h. Formed aggregates were then transferred onto low attachment 6-well plates and continued culture for another 4 days. To induce neural rosette formation, the aggregates were transferred onto Geltrex coated 6-well plate and cultured for 4 days in neural induction medium supplemented with 200 ng/mL Noggin, 200 ng/mL DKK1 (R&D Systems), and 20 ng/mL bFGF (Life Technologies). Neural rosettes were manually picked and transferred onto low attachment 6-well plate in neural induction medium supplemented with 10 ng/mL bFGF and 10 ng/mL EGF to allow neuroepithelial sphere formation. Five days later, two procedures were performed to directly neural differentiation. First, the neural aggregates were placed onto Geltrex coated plates and treated with neural differentiation medium consisting of Neurobasal A medium supplemented with 10 ng/mL BDNF, 10 ng/mL GDNF, 200 μM ascorbic acid, 500 μM cyclic AMP. Second, the neural aggregates were dissociated into single cells with Accutase and replated onto poly-D-lysine/laminin coated 24 or 48-well plates for further differentiation with neural differentiation medium supplemented with 10 ng/mL BDNF, 10 ng/mL GDNF, 200 μM ascorbic acid, 500 μM cyclic AMP.
Generation of cerebral organoids
Generation of cerebral organoids from iPSCs was performed based on an unguided approach previously described  with minor modifications to improve reproducibility, especially batch variations. Briefly, the neuroectodermal in the form of neuronal rosette were generated from multipotent iPSC aggregates (EBs) in suspension using minimalistic media, coupled with Matrigel embedding, and followed by the expansion of the elaborate architecture of the germinal zone of neural stem cells or progenitor cells with the differentiated neurons migration outwards containing the neural tissues of diverse identities or the potential to developing into multiple neural structures. Neuroepithelial buds developed further in a bioreactor. A major modification was using pipetting technic to remove all debris accumulated around the EBs by setting a multichannel pipette at 30 μL and gently pipetting up and down 5 times just before half medium replacement, which, we found, could largely improve EB’s quality. Another modification was to maintain the organoids in 96-well U-bottom ultra-low attachment plates until Matrigel embedding with half medium changing daily, aiming for individual organoid tracing and quality control.
Generation of brain region-specific organoids
Generation of brain region-specific organoids from iPSCs was performed mainly based on a previous report  with modifications. The major modifications include: 1) culturing iPSCs with feeder free methods and generating EBs with Aggrewell™ 800 plates; 2) pattering since Day1; and 3) three-dimensional suspension culturing on an orbital shaker (Thermo fisher) with 120 rpm since Day2. Briefly, on Day 0, iPSC colonies were detached from Matrigel (Corning, cat. no. 354230) and disassociated by ReLeSR™ (Stemcell Technologies). After washed with fresh stem cell medium with 10 μM Y-27632, 10,000 cells per microwell of Aggrewell™800 (3.0 × 106 per well of 24-well in 2 mL stem cell medium with 10 μM Y-27632) were seeded by centrifuging. From Day 1, half of the medium was replaced following protocols diverge for the procedures to differentiate forebrain, midbrain and hypothalamus organoids developed by Qian et al. . The protocol of hippocampus organoids was developed by Fadi et al. . Briefly on Day 2, around 30 organoids were transfer to each well of a 6-well plate placed on an orbital shaker rotating at 120 rpm in an incubator. For hippocampus organoids, on Days 1-7, half medium was replaced with EB medium consisting of DMEM/F12 (Thermo Fisher Scientific), 1XGlutaMAX™ Supplement(Thermo Fisher Scientific), 20% (vol/vol) KnockOut™ Serum Replacement (Thermo Fisher Scientific), 1X Antibiotic–Antimycotic (Thermo Fisher Scientific), 1X 2-Mercaptoenhanol (Thermo Fisher Scientific), 1X Non-essential Amino Acids(NEAA, Thermo Fisher Scientific), 1 IU/mL Heparin (StemCell Technologies), 100 nM LDN193189 (StemCell Technologies), and 5 µM SB431542 (StemCell Technologies). On Days 8-14, medium was exchanged to hippocampus patterning medium consisting of DMEM/F12, 1X GlutaMAX™ Supplement, 1XNEAA, 1X N2 (Thermo Fisher Scientific), 1 IU/mL Heparin, 2 µM CHIR99021 (StemCell Technologies), and 10 ng/mL Recombinant Human BMP-7(Peprotech). On Days15-44, hippocampus neurogenesis medium was applied, which consists of 1:1 DMEM/F12: Neurobasal Medium (Thermo Fisher Scientific), 1X GlutaMAX™ Supplement, 1X NEAA, 1X N2, 1 IU/mL Heparin, 1X B-27 Supplement Minus Vitamin A, and 2.5 mg/mL human insulin (Sigma-Aldrich). After Day 45, neurogenesis medium was exchanged with maturation medium consisting of 1:1 DMEM/F12: Neurobasal Medium, 1X GlutaMAX™ Supplement, 1X NEAA, 1X N2, 1 IU/mL Heparin, 1X B-27 Supplement Minus Vitamin A, 2.5 mg/mL human insulin, 20 ng/mL BDNF(Peprotech), 20 ng/mL recombinant Human GDNF (Peprotech), 200 µM Ascorbic Acid (StemCell Technologies), and 500 µM Dibutyryl-cAMP (StemCell Technologies).
Immunofluorescent staining of neural cells were performed as previously described . The cells were fixed with 4% paraformaldehyde (PFA) for 15 min, permeabilized with 0.1% Triton-X100 in PBS for 15 min and blocked with 5% bovine serum albumin(BSA), 5% normal goat serum, and 0.1% Triton X-100 in PBS at room temperature for 30 min, then incubated with primary antibodies overnight at 4 °C. After washing with 0.1% Tween-20/PBS, the samples were incubated with secondary antibodies for 1 h and counterstained with DAPI nuclear staining. Quantitative results were obtained with ImageJ software counting 10 random fields of each experiment.
Brain organoids were fixed with 4% PFA for 30 min at room temperature. Organoids were washed 3 times with PBS and immersed in a sucrose solution gradient, 10% for 30 min, 20% for 1 h, then in 30% sucrose solution overnight. Organoids were embedded in tissue freezing medium (Tissue-Tek® O.C.T. Compound) and sectioned with a cryostat (Thermo) at 16 μm thickness. For immunostaining, cryosection slides were laid at room temperate for at least 20 min, washed 3 times with PBS before being blocked with blocking medium consisting of 5% goat serum, 5% bovine serum albumin, 0.5% Triton-X in PBS for 1 h followed by incubation with primary antibodies diluted in PBS buffer containing 0.5% goat serum, 0.5% BSA, 0.1% Tween 20 and 0.05% Triton-X at 4 oC overnight. For OXR1 staining, sections were then incubated at 90 °C in citrate antigen-retrieval buffer (pH 6.0) for 25 min and rinse once with PBS before blocking. After three washes in PBS containing 0.1% Tween 20, staining with secondary antibodies and DAPI was performed for 1 h at room temperature in the dark moisture chamber. After three washes in PBS containing 0.1% Tween 20 (10 min each), once quickly rinsing in PBS, and once quickly rinsing in pure water, the slides were mounted with ProLong™ Glass Antifade Mountant (Fisher Scientific, Waltham, Massachusetts). Cover glass thickness was 0.17 mm. The primary and secondary antibodies used, as well as their dilution are provided in Additional file 3: Table S1.
Analysis of immunolabelling
Images were captured on a Leica SP8 confocal microscope, using a 40 × oil-immersion objective. Image analysis was performed in Fiji (ImageJ, NIH). Prior to analysis, Z-stacks were flattened with maximum or average z-projection. The same capture conditions and analysis settings were applied on both control and patient organoid sections. For quantification of progenitor cell proliferation and apoptosis, organoids were stained for KI67 and cleaved Caspase3(cCas3), respectively. Random fan-shaped regions from the apical surfaces of the ventricular zone (VZ) to the boundary between the outer subventricular zone (oSVZ)and cortical plate (CP) or between VZ and intermediate zone (IZ) in midbrain organoids were cropped for analyses. KI67+ nuclei and cCas3+ were counted and divided by the total number of nuclei stained by DAPI in the region respectively. To quantify the distribution of histone modifications in the developing cortical and midbrain organoids, radial structures were chosen randomly. From the pial surface to the basal surface, radial columns were cropped and evenly divided into 10–11 bins for analyses. The relative vertical position coordinates, y-coordinates on the image were recorded and normalized to the full thickness to measure their relative laminar positions in the differentiation and migration processes of midbrain progenitor cells or cortical laminar positions. The positions of each modification positive nuclei were separately marked using the Cell Counter plugin in Fuji(version 2.3.0). The positive cell proportion of the relative vertical positions in 10–11 evenly divided bins for each marker were calculated by (positive cell counts within each bin/total positive cell counts) and plotted in Prism (GraphPad 8.0). The mean intensity per cell were used to analysis the expression level of each marker.
RNA sequencing and analysis
We conducted RNA-sequencing with three individual clones of each genotype at three stages of neuronal differentiation: iPSC (iPS_CTRL and iPS_OXR1ΔEx18), neural aggregate (NA) (NA_ CTRL and NA_ OXR1ΔEx18), and neuron (Neu) (Neu_CTRL and Neu_OXR1ΔEx18). in total, 18 samples were sequenced. Library construction and sequencing were performed through a commercially available service provided by BGI TECH SOLUTIONS (HONGKONG) CO. (Agreement No. F20FTSEUHT0295).
The quality control of fastq files was performed with FastQC v0.11.9 . Alignment to reference genome (GRCh38) was accomplished with hisat2 v2.1.0 , while annotation and count matrix were computed with featureCounts v.2.0.0 . We accomplished downstream DEGs analysis in R v3.6.1 with DESeq2 v1.24.0 . Gene Set Enrichment Analysis(GSEA)  was performed to identify enriched pathways (using KEGG  and Reactome  databases) and gene ontology (GO) terms. Heatmaps were generated in R v3.6.1 with heatmap3 v1.1.7 . For the topologic biological network analysis, 1978 leading-edge genes determined by GSEA, which contributed the most to the top enriched GO terms in nervous system development, cell adhesion, extracellular matrix organization, neuron death, stress response, mitochondrial genome maintenance, glucose metabolism, lysosomal structure, and TFs from DEGs with |Log2FC|> 0.5 were recruited. Next, we constructed the protein–protein interaction (PPI) network for differential expression gene by searching STRING protein interaction database , with string score > 700 as the low limit value of confidence, FDR < 5%, and then performed network modules analysis by ReactomeFIViz . A topologic biological network composed of the PPIs of 1539 DEGs clustering into 44 modules were established to help us infer the central transcriptome events. Each module consists of a set of genes that are both connected in the protein functional interaction network and highly-correlated in biological databases . We run cytoHubba , TRRUST  and iRegulon(v1.3)  to identifiy hub genes, the drivers of a module and highly connected, and the essential upstream regulators. This network data were imported into and visualized by Cytoscape software v3.8.1 .
Proximity ligation assay
Proximity ligation assays (PLAs) were performed on fibroblast cells derived from the OXR1ΔEx18 patient and a healthy control. Cells were grown on glass coverslips, fixed with 4% PFA (added 1 M glycine in the second wash step after fixation), and permeabilized with 0.1% Triton-X/1X PBS for 20 min. The PLA was conducted according to the manufacturer’s instructions using the Duolink In Situ Red Starter Kit Mouse/Rabbit (Cat. No. DUO92101, SIGMA). After blocking for 30 min at 37 °C, the cells were incubated with anti-OXR1 (1:200) and PRMT5 antibody (1:200) or anti-OXR1 and anti-PRMT1 antibody (1:200) overnight at 4 °C. PLA Anti-Rabbit PLUS and Anti-Mouse MINUS probes containing the secondary antibodies conjugated with oligonucleotides, were added, and incubated at 37 °C for ligation 30 min. Then the fluorescently labeled oligonucleotides resulting in red fluorescence signals were added to the reaction and incubated for amplification at 37 °C for 100 min in the dark. The slides were mounted with Duolink In Situ Mounting Medium with DAPI (Cat. No. DUO82040, SIGMA). Microscopy was carried out using a Leica SP8 confocal microscope equipped with a 40X oil-immersion lens. The same input parameters were used throughout all experiments. Sample images were prepared in FUJI (ImageJ, NIH).
The U2OS cell pellet was lysed with NETN buffer (20 mM Tris–HCl pH 8.0, 100 mM NaCl, 0.5% NP-40, 0.5 mM DTT, 1:100 protease inhibitor cocktail, 400 U/mL). The lysate was pre-cleared by Dynabeads™ Protein A (Thermofisher Scientific, Cat#10001D) beads for 30 min. 1 mg pre-clear cell lysate was pre-incubated with 1 μg of purified rabbit anti IgG antibody (Sigma-Aldrich Cat# I5006, RRID:AB_1163659) or 3 μg OXR1 antibody (Bethyl Cat# A302-036A, RRID:AB_1576565) for 1 h at 4ºC rotator, and then incubated with 20 μl of protein A beads overnight at 4 °C with gentle agitation. The beads were pelleted and washed 3 times in 1 × TBS buffer (50 mM Tris HCl pH 7.4, 150 mM NaCl, add 1 mM PMSF before used). The precipitated proteins were denatured in SDS loading buffer at 95 °C for 10 min and separated in 10% SDS-PAGE gels.
Methylation assays with H4 N-terminal tail (aa 1–21) peptides
In vitro methylation assays were performed as previously described . In a 30μL reaction, 7.35 nM PRMT5/MEP50 complex (Active motif, Cat#31,356), 0.4 mM N-terminal H4 peptides, H4(1–21) WT and H4(1-21_R3K), with and without OXR1A or truncated OXR1A_ΔTLDC_1 or OXR1A_ΔTLDC_2, in a concentration gradient 1.83 nM( +), 7.35 nM(+ +) and 29.4 nM(+ + +) respectively, were carried out in 50 mM HEPES (pH 8.0), 10 mM NaCl, and 1 mM DTT containing [3H] SAM (25:1 molar ratio of SAM(NEB, Cat#B9003S) to [methyl-3H]-SAM (PerkinElmer, Cat#NET155001MC). Reactions were incubated for 1 h at 37°C, and then quenched with 0.5 μL of 100% trifluoroacetic acid. Each reaction was run in quadruplicate. Twenty-five microliters of each reaction were transferred to Merck Millipore MultiScreenHTS 96-Well Filter Plates with Negatively Charged Membrane (Cat#10,245,703) and air-dried for 30 min. The papers were subsequently washed in 50 mM NaHCO3 at pH 9.0 for 45 min, then air dried. Radioactivity was counted using 1450 MicroBeta Instrument (Wallac at Department of Immunology and Transfusion Medicine, Oslo University Hospital, Oslo, Norway).
Quantification and statistical analysis
Each assay consists of at least two independent experiments with technical triplicates as described specifically. Data are presented as mean ± S.E.M., or mean ± S.D., as indicated in the figure legends. Organoid samples were randomly taken from the culture for experiments and analyses. Data analyses comparing control and OXR1 patient derived organoids were not performed blindly because the visual difference between the two groups is striking, and blinding was not possible to an informed researcher. Briefly, statistical analyses were performed using the t test in Excel or Graphpad Prism 8. A confidence level of 95% (*p < 0.05; ** p < 0.01, *** p < 0.001) was considered statistically significant. Detailed information was included in each figure legend.
Availability of data and materials
Datasets supporting the conclusions are available in the Gene Expression Omnibus repository (GEO) at accession number (GSE:206,614) .
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We thank the family of the described patients for participating in our studies.
The review history is available as Additional file 5.
Peer review information
Tim Sands was the primary editor of this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.
Open access funding provided by University of Oslo (incl Oslo University Hospital). Funding was provided by the Research Council of Norway, South-East Norway Regional Health Authority and Central Region Norway Health Authority. The Proteomics and Metabolomics Core Facility PROMEC is funded by NTNU and the Central Norway Regional Health Authority and is a member of the National Network of Advanced Proteomics Infrastructure (NAPI), which is funded by the RCN INFRASTRUKTUR-program (295910).
Ethics approval and consent to participate
The study was performed with the approvals of the ethical committees of Hadassah Medical Center and the Ministry of Health (0393–17-HMO), and South-East Health Authorities of Norway (2014/658).
The authors declare no conflicts of interest.
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Supplementary Video 1, The cardiomyocyte differentiation of control (Video S1a) and OXR1ΔEx18 (Video S1b) iPSCs.
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Lin, X., Wang, W., Yang, M. et al. A loss-of-function mutation in human Oxidation Resistance 1 disrupts the spatial–temporal regulation of histone arginine methylation in neurodevelopment. Genome Biol 24, 216 (2023). https://doi.org/10.1186/s13059-023-03037-1