DiGeorge syndrome critical region 8 (DGCR8) is a critical component of the canonical microprocessor complex for microRNA biogenesis. However, the non-canonical functions of DGCR8 have not been studied. Here, we demonstrate that DGCR8 plays an important role in maintaining heterochromatin organization and attenuating aging. An N-terminal-truncated version of DGCR8 (DR8dex2) accelerated senescence in human mesenchymal stem cells (hMSCs) independent of its microRNA-processing activity. Further studies revealed that DGCR8 maintained heterochromatin organization by interacting with the nuclear envelope protein Lamin B1, and heterochromatin-associated proteins, KAP1 and HP1γ. Overexpression of any of these proteins, including DGCR8, reversed premature senescent phenotypes in DR8dex2 hMSCs. Finally, DGCR8 was downregulated in pathologically and naturally aged hMSCs, whereas DGCR8 overexpression alleviated hMSC aging and mouse osteoarthritis. Taken together, these analyses uncovered a novel, microRNA processing-independent role in maintaining heterochromatin organization and attenuating senescence by DGCR8, thus representing a new therapeutic target for alleviating human aging-related disorders.
Aging is a crucial driving force in human degenerative disorders. Possible mechanisms for organismal aging include loss of heterochromatin1, free radicals, programmed senescence, telomere shortening, genomic instability2, nutritional intake, and growth signaling3. Stem cell aging is newly recognized as an important culprit in organismal aging3,4. For example, aging of mesenchymal stem cells (MSCs) has been shown to drive aging-associated tissue degeneration5. MSCs, which have the potential to differentiate into mesodermal lineages like osteoblasts, chondrocytes and adipocytes, can be isolated from various tissues including bone marrow, cord blood, adipose tissue and dental pulp6. Premature depletion of MSCs is observed in patients with Hutchinson-Gilford progeria syndrome (HGPS) and Werner syndrome (WS), two premature aging diseases that are associated with accelerated atherosclerosis, osteoporosis, and osteoarthritis7,8. Despite numerous studies showing that MSCs play pivotal roles in tissue rejuvenation, regeneration, and repair by differentiating into various somatic cell types6, little is known about the key regulators of MSC aging.
Aging-associated declines in stem cell functionality are often accompanied by epigenetic changes, such as changes in genomic DNA methylation, histone modifications, and chromatin remodeling enzymes3,4,9,10. Heterochromatin domains are structurally inaccessible and usually transcriptionally inactive. These domains are established during early stages of embryogenesis and are gradually lost with aging, resulting in the de-repression of normally silenced genes1. Heterochromatin is associated with specific proteins, such as heterochromatin protein 1 (HP1), and specific histone modifications, like H3K9me3 (trimethylated histone H3 at lysine-9)11, both of which decrease along with heterochromatin loss. In MSCs derived from patients with HGPS or WS, reduction of heterochromatin is observed, along with decreased levels of HP1, H3K9me3, and H3K27me34,7,9,12. Whereas heterochromatin loss drives human MSC (hMSC) aging9,13, the re-establishment of heterochromatin alleviates premature aging and promotes longevity in Drosophila and human cells14,15, suggesting that the maintenance of heterochromatin organization could be an effective therapeutic intervention against aging.
DGCR8, which is known as Pasha in flies and worms, is a 773-amino-acid (a.a.) protein critical for microRNA (miRNA) biogenesis via the formation of a microprocessor complex with Drosha. This complex processes primary miRNA in the nucleus16. The C-terminal tail (a.a. 685-773) of DGCR8 binds to and stabilizes Drosha; the dsRNA-binding domain (a.a. 510-684) interacts with pri-miRNAs to increase binding affinity; the RNA-binding heme domain (a.a. 276-498) ensures processing accuracy and enhances efficiency. In contrast, little is known about the N-terminal region (a.a. 1-270), except that it contains a nuclear localization signal (NLS)17,18. During the past decades, studies on DGCR8 have primarily focused on its miRNA-processing function. DGCR8-knockout mouse embryonic stem cells (mESCs) have defects in proliferation and differentiation due to the global loss of miRNAs19. DGCR8 deletion in murine B-cell progenitors impairs the transition from pro- to early pre-B cell stage, recapitulating the phenotypes observed in miR17-92 cluster knockout mice20. However, it was recently reported that UV-induced phosphorylation of DGCR8 at S153 mediates transcription-coupled nucleotide excision repair of UV-induced DNA lesions, independent of its miRNA-processing activity21, suggesting that DGCR8 may have non-canonical functions.
In this study, we show that DGCR8 interacted with heterochromatin and nuclear lamina proteins to maintain heterochromatin organization. Expressing a mutant version of DGCR8 lacking the N-terminal region in hMSCs resulted in heterochromatin destabilization and accelerated cellular senescence in a miRNA processing-independent way. Lentiviral-mediated expression of DGCR8, or DGCR8-interacting proteins such as KRAB-associated protein 1 (KAP1), Lamin B1, or HP1γ, rescued senescent phenotypes in DGCR8-deficient hMSCs. Moreover, a decrease in DGCR8 level was detected in hMSCs from older individuals, and restoring level of wildtype DGCR8 (or a miRNA-irrelevant version of DGCR8) attenuated hMSC senescence, as well as post-traumatic articular aging and osteoarthritis in mice. These results demonstrate a non-canonical role for DGCR8 in maintaining nuclear and heterochromatin organization, and protecting against cellular senescence in human MSCs. Thus, DGCR8 represents a new therapeutic target for combating age-associated disorders.
Generation of DGCR8-defective human ESCs
To study the role of DGCR8, we purposed to generate DGCR8-defective human embryonic stem cells (hESCs) by CRISPR/Cas9-mediated gene editing22. Initially, we tried to generate DGCR8-deficient hESCs by targeting exon 3 of DGCR8 using Cas9 and sgRNAs according to the previous study performed in mice19, resulting in DR8ex3 hESCs (Supplementary Fig. 1a, b). However, DR8ex3 hESCs exhibited abnormal karyotypes, with an inter-chromosomal translocation between chromosomes 19 and 22 (Supplementary Fig. 1c). Therefore, we next sought to target exon 2 of DGCR8 which contains the start codon ATG (Supplementary Fig. 1d). Complete ablation of full-length DGCR8 protein was confirmed by immunoblotting and immunofluorescence analyses (Supplementary Fig. 1e, p). DR8ex2 hESCs were karyotypically normal (Supplementary Fig. 1g), but exhibited poor maintenance of self-renewal capacity (Supplementary Fig. 1f, h–j) and high levels of spontaneous differentiation (Supplementary Fig. 1k). We next generated hESCs with an N-terminal-truncated version of DGCR8 (Deletion of a.a. 1-240) by specifically removing the coding sequence in exon 2 using a CRISPR/Cas9-mediated homologous recombination strategy23 (referred to as DR8dex2 hESCs) (Fig. 1a, b and Supplementary Fig. 1l, m, p). The resultant DR8dex2 hESCs were cultured for more than 50 passages without the emergence of abnormal karyotypes (Fig. 1c), or the loss of pluripotency markers (Fig. 1d and Supplementary Fig. 1n). DR8dex2 hESCs were also capable of forming teratomas composed of the three germ layers in vivo, confirming normal pluripotency (Fig. 1e and Supplementary Fig. 1o). Cell cycle kinetics and clonal expansion ability were also normal in DR8dex2 hESCs (Fig. 1f, g).
DGCR8dex2 accelerates senescence of hMSCs
To characterize the effects of DR8dex2 on the function of post-pluripotent cells, we differentiated DR8dex2 hESCs into human mesenchymal stem cells (referred to as DR8dex2 hMSCs)9,10,24. DR8dex2 hMSCs were positive for typical hMSC markers, such as CD73, CD90, and CD105, and negative for hMSC-irrelevant antigens, including CD34, CD43, and CD45 (Fig. 2a and Supplementary Fig. 2a). Western blotting verified the lack of full length DGCR8 in DR8dex2 hMSCs using an anti-DGCR8 antibody (Fig. 2b). Next, we analyzed the growth rate of DR8dex2 hMSCs through serial passaging. Compared with wildtype (WT) hMSCs that showed normal aging kinetics, DR8dex2 hMSCs senesced at as early as passage 3, and completely arrested growth at passage 6 (Fig. 2c). In addition to premature senescence, DR8dex2 hMSCs exhibited decreased proliferative potential (Fig. 2d and Supplementary Fig. 2b), increased senescence-associated (SA)-β-gal activity (Fig. 2e), upregulation of senescence markers P16 and P21 proteins (Fig. 2f), activation of senescence-associated secretory phenotype (SASP) (Fig. 2g), and enhancement of the DNA damage response (DDR) (Supplementary Fig. 2c, d). Despite increased DDR, genome-wide copy number variation (CNV) analysis indicated that genomic integrity was maintained in DR8dex2 hMSCs (Fig. 2h). In line with premature cellular senescence, chondrogenesis and osteogenesis were impaired in DR8dex2 hMSCs (Supplementary Fig. 2e, f). Finally, implantation of hMSCs into the tibialis anterior muscles of nude mice (Fig. 2i) revealed accelerated decay of DR8dex2 hMSCs compared to WT hMSCs. Taken together, these data suggest that the N-terminal truncation of DGCR8 promotes premature senescence and functional attrition of hMSCs.
DGCR8dex2 exhibits no global effects on miRNA processing
To determine whether the phenotypes of DR8dex2 hMSCs resulted from loss or gain of DGCR8 function, WT hMSCs were transduced with lentiviral vectors encoding: 1) Cas9 and a sgRNA targeting exon 5 of DGCR8, or 2) a DGCR8-specific shRNA. Both effectively reduced DGCR8 protein levels (Supplementary Fig. 2g, j) and induced premature senescence with phenotypes such as impaired cell proliferation (Supplementary Fig. 2h, k) and early onset of SA-β-gal activity (Supplementary Fig. 2i, l). By contrast, restoring WT DGCR8 (DR8-WT) in DR8dex2 hMSCs by lentivirus-mediated cDNA transfer alleviated cellular senescence (Fig. 2j, k and Supplementary Fig. 2m). These observations suggest that accelerated senescence of DR8dex2 hMSCs results from loss of DGCR8 activity.
We next investigated whether defective DGCR8 caused accelerated senescence in DR8dex2 hMSCs due to loss of its well-known miRNA-processing activity. Consistent with the fact that the C-terminal domains of DGCR8 are necessary for pri-miRNA processing25, the interaction between DR8dex2 and Drosha was preserved in DR8dex2 hMSCs, and there were no global differences in miRNA expression profiles between DR8dex2 and WT hMSCs (Supplementary Fig. 3a–c). Further, based on previous studies showing that the simultaneous mutation of amino acids 568, 569, 676, and 677 of DGCR8 abrogate its miRNA-processing ability25, we generated an expression vector for DGCR8-AA568-569KK/AS676-677KK (mutant of double-strand RNA-binding domain; mtDRBD), which lacked miRNA-processing function but preserved the interaction with Drosha (Supplementary Fig. 3d, e). Expression of DGCR8 mtDRBD (DR8-mtDRBD) rescued premature aging defects as effectively as DR8-WT, as evidenced by enhanced cell proliferation (Supplementary Fig. 3f and Fig. 3a, b), downregulation of cellular senescence-associated markers (Fig. 3c), reduced SA-β-gal activity (Fig. 3d), decreased DDR (Fig. 3e) in hMSCs-DR8dex2. Also, miRNA profiles were not globally changed in DR8dex2 hMSCs expressing DR8-WT or DR8-mtDRBD (Supplementary Fig. 3g). This suggests that the miRNA-processing function of DGCR8 is irrelevant to its ability to regulate hMSC senescence.
DGCR8 is required for maintaining heterochromatin structure
To further explain why N-terminal truncation of DGCR8 induced accelerated MSC senescence, we sought to identify new DGCR8-interacting proteins. We expressed Flag-tagged DGCR8 in HEK293T cells and performed immunoprecipitation with an antibody against Flag, followed by mass spectrometry (Supplementary Fig. 4a). In addition to well-known DGCR8-associated proteins like Drosha, we identified a component of heterochromatin, KAP1 (Supplementary Fig. 4b). Due to the limited resolution of mass spectrometry in recognizing low abundant and trypsin-insensitive proteins9, we performed co-immunoprecipitation (Co-IP) and identified more heterochromatin components that interacted with DGCR8, revealing DGCR8 as part of a complex containing major heterochromatin proteins KAP1, HP1α, HP1γ, as well as the nuclear lamina protein Lamin B1 (Fig. 4a, b and Supplementary Fig. 4c). In addition, chromatin immunoprecipitation-quantitative polymerase chain reaction (ChIP-qPCR) revealed that DGCR8 bound pericentromeric repetitive sequences, including α-Satellite (α-Sat) and Satellite 2 (Sat2) (Fig. 4c), which are primarily wrapped as architecturally condensed constitutive heterochromatin26,27,28. Furthermore, DGCR8 associated with heterochromatin-enriched genomic regions such as LINE1 and ribosomal DNA (rDNA)4,10,29,30,31 (Supplementary Fig. 4d). These data indicate that DGCR8 may be an important component of constitutive heterochromatin.
In DR8dex2 hMSCs, expression levels of KAP1, HP1α, HP1γ, and Lamin B1 were each decreased (Supplementary Fig. 4e). ChIP sequencing (ChIP-seq) analysis of DR8dex2 hMSCs revealed the downregulation of H3K9me3, a marker of constitutive heterochromatin, across the genome, especially within heterochromatin (HC) and lamina-associated domains (LADs) (Fig. 4d, e and Supplementary Fig. 5a–c). ChIP-qPCR further confirmed reduced H3K9me3 enrichment at genomic repetitive sequences, including α-Sat, Sat2, LINE1, and rDNA loci (Fig. 4f and Supplementary Fig. 4f), correlating with the upregulation of transcripts from these repetitive sequences (Fig. 4g and Supplementary Fig. 4g). In addition, we observed loss of “H3K9me3 mountains” in DR8dex2 hMSCs, which were mainly located at genomic repetitive sequences, such as pericentromeres (Fig. 4h and Supplementary Fig. 5d). Consistent with these changes in chromatin architecture, RNA-seq analysis revealed global changes in gene expression associated with heterochromatin decondensation in DR8dex2 hMSCs (Supplementary Fig. 6a–c). Finally, these heterochromatin alterations were reversed by the re-introduction of DR8-WT or DR8-mtDRBD, which was demonstrated by H3K9me3 ChIP-seq (Fig. 4i and Supplementary Fig. 6d). Taken together, these data indicate that DGCR8 is essential for maintaining heterochromatin architecture.
DGCR8 interacts with Lamin B1 and HC components
In human premature aging disorders, enlarged nuclei and loss of heterochromatin underneath the nuclear envelope are often observed9. Similar features were present in DR8dex2 hMSCs (Fig. 5a and Supplementary Fig. 7a, b, and i), suggesting a causative link between the inability of DR8dex2 to maintain heterochromatin organization and accelerated cellular senescence.
To map the domains of DGCR8 required for interacting with Lamin B1 and heterochromatin components, we generated a series of Flag-tagged DGCR8 fragments, including a.a. 1–300, a.a. 1–510, a.a. 300–773, a.a. 358–773, and a.a. 510–773 (Supplementary Fig. 7c–e). DGCR8 a.a. 300–773 was excluded from further investigations due to cytoplasmic mislocalization (Supplementary Fig. 7e). Both DGCR8 a.a. 1–300 and a.a. 510–773 contain domains that interacted with the heterochromatin components, KAP1 and HP1γ. For the a.a. 1–510 construct, a.a. 300–510 exerted an auto-inhibitory effect on a.a. 1–300 (Supplementary Fig. 7g). A docking site for Lamin B1 was deduced in DGCR8 a.a. 300–358 (Supplementary Fig. 7g, h). These studies suggest that DGCR8 acts as a scaffold protein to tether heterochromatin proteins to the nuclear lamina. Subsequently, we investigated whether such a scaffold-like activity of DGCR8 is required for regulating cellular senescence. We generated a DGCR8 mutant (DR8-Δ58, which lacks a.a. 300–358) that interacted with KAP1 and HP1γ, but not Lamin B1 (Supplementary Fig. 7f, h). Compared to DR8-WT, DR8-Δ58 was less able to rescue accelerated senescence in DR8dex2 hMSCs (Fig. 5b, c). On the other hand, overexpression of Lamin B1, KAP1, or HP1γ each alleviated senescent phenotypes of DR8dex2 hMSCs both in vitro and in vivo (Fig. 5d–h and Supplementary Fig. 8a, b). Taken together, these data suggest that DGCR8 regulates hMSC aging by maintaining heterochromatin organization via its interactions with Lamin B1 and heterochromatin components.
DGCR8 overexpression exerts a geroprotective role
We next investigated DGCR8 levels during physiological and pathological aging. In primary hMSCs derived from human donors at different ages, DGCR8 levels decreased with age (Fig. 6a and Supplementary Fig. 9a). DGCR8 was also downregulated in three stem cell models of human aging, including replicative senescent hMSCs (RS hMSCs), WRN-deficient hMSCs (WS-specific hMSCs) and LMNA (p.G608G)-mutated hMSCs (HGPS-specific hMSCs) (Fig. 6b and Supplementary Fig. 9b).
We then explored the possibility of targeting DGCR8 to alleviate human cellular aging. We transduced hMSCs with lentiviruses expressing DR8-WT or DR8-mtDRBD which could not participate in miRNA processing, and assessed aging phenotypes. Overexpression of either form of DGCR8 effectively rescued premature aging phenotypes in RS hMSCs (Fig. 6c and Supplementary Fig. 9c–e, j) and WS-specific hMSCs (Fig. 6d and Supplementary Fig. 9f, g, j). The senescence-associated phenotypes of hMSCs isolated from an 80-year-old individual were also alleviated by overexpressing either DR8-WT or DR8-mtDRBD (Fig. 6e, f and Supplementary Fig. 9h–j).
Lastly, we evaluated the in vivo beneficial effects of overexpressing DGCR8 on alleviating aging-related phenotypes. Joint is a critical tissue affected by the accumulation of senescent cells, which serves as a key inducement for the development of osteoarthritis32. Recently, the chemical clearance of senescent cells in mouse bone joint has been shown to alleviate the phenotypes of post-traumatic osteoarthritis and even to prevent age-associated bone loss33. Accordingly, we proposed that the ectopic expression of DGCR8 may attenuate the development of osteoarthritis by repressing cellular senescence. We utilized anterior cruciate ligament transection (ACLT) to induce articular aging and osteoarthritis in mice33 (Fig. 7a). One week after ACLT, lentiviruses encoding luciferase (Luc), DR8-WT or DR8-mtDRBD were injected into the articular cavities of these mice. Seven weeks post lentivirus injection, microcomputed tomography (micro-CT) analyses showed that in contrast to the Luc-injected ACLT mice that exhibited obvious bone damage, the bone quality of the ACLT mice with DR8-WT or DR8-mtDRBD was well maintained as those of the mice without ACLT (Fig. 7b). The subsequent histological assessment showed that ACLT-induced articular cartilage degeneration as the classical phenotype of osteoarthritis was rescued by injecting lentiviruses encoding DR8-WT or DR8-mtDRBD (Fig. 7c). Cellular senescence marker P16 was decreased whereas cellular proliferation marker Ki67 was increased in the articular cartilage of ACLT mice treated with lentiviral vectors expressing DR8-WT or DR8-mtDRBD as indicated by immunohistochemistry (Fig. 7d, e). In addition, the mRNA levels of typical aging markers (e.g., P16 and P21) and inflammation factors (e.g., IL6 and matrix metalloproteinase 13 (MMP13)) that are often upregulated in osteoarthritis were diminished in the articular cavities of the mice with DR8-WT or DR8-mtDRBD (Fig. 8a). Furthermore, RNA-seq analysis revealed that the ectopic expression of DR8-WT or DR8-mtDRBD reversed major gene expression alterations including increased inflammation and apoptosis, and also promoted osteogenesis and chondrogenesis in the articular cavities of the mice with ACLT-induced osteoarthritis (Fig. 8b and Supplementary Fig. 10a, b). Meanwhile, the rescue effect of DGCR8 on osteoarthritis was further evaluated in physiologically aged mice (Fig. 9a). Similar to ACLT mice, physiologically aged mice exhibited decreased bone density, upregulated aging markers and inflammation factors, and downregulated bone development-associated genes (Supplementary Fig. 10c–e). The bone quality of aged mice treated with lentiviral vectors expressing DR8-WT or DR8-mtDRBD was obviously improved compared to luc-expressed lentivirus-treated mice as control (Fig. 9b). In addition, the expression levels of factors involved in osteoarthritis were also rescued by DR8-WT or DR8-mtDRBD lentivirus-mediated treatment (Fig. 9c, d). Taken together, these results demonstrate that the inhibitory effects on cellular senescence and inflammatory response by DGCR8 may contribute to regeneration of bone and cartilage and alleviation of the symptoms of age-related osteoarthritis, suggesting the therapeutic potential of DGCR8 as a geroprotective factor.
In this study, we presented several lines of evidence supporting a non-canonical, miRNA processing-independent role of DGCR8 in regulating heterochromatin organization and protecting against cellular aging. First, DGCR8 was essential for preventing premature senescence by maintaining heterochromatin organization via interactions with both heterochromatin and lamina proteins. Second, DGCR8 levels were downregulated with age. Third, DGCR8 was geroprotective, both in vitro and in vivo. For the first time, we show that DGCR8 regulated hMSC aging through heterochromatin maintenance, thus identifying DGCR8 as an innovative intervention target for aging.
DGCR8 has been well studied as a miRNA-processing protein19,34,35,36. DGCR8-knockout mESCs arrest in the G1 phase of the cell cycle, a phenotype that can be rescued by members of the miR-290 family35. Disrupting DGCR8 in vascular smooth muscle cells reduces cell proliferation and promotes apoptosis via the downregulation of miR-17/92 and miR-143/145 clusters34. Conditional knockout of DGCR8 in skin results in the depletion of miRNAs and skin developmental defects36. In addition, DGCR8 regulates neural development in both miRNA-dependent and miRNA-independent ways37,38. A recent study has also reported a miRNA-independent function of DGCR8 in facilitating the splicing of Tcf711 that is necessary for mESCs to exit pluripotency and differentiate39. This suggests that DGCR8 has miRNA-independent functions. Here, we demonstrate that DGCR8 plays a distinct role in regulating hMSC senescence due to its function in maintaining heterochromatin architecture rather than miRNA processing.
We observed that hMSCs with loss of DGCR8 function exhibited abnormal nuclear and chromatin organization, with no marked changes in global miRNA profiles. Although 10.9% of the total miRNAs were differentially expressed in DR8dex2 hMSCs, these miRNAs may not be the major contributor to the heterochromatin changes in DR8dex2 hMSCs as evidenced by the minimal changes in miRNA profiles of DR8dex2 hMSCs expressing Luc, DR8-WT, or DR8-mtDRBD. To our knowledge, this is the first evidence that DGCR8 stabilizes heterochromatin by interacting with the nuclear lamina and heterochromatin proteins, independent of its miRNA-processing ability. We further propose a model in which DGCR8 functions as a scaffold protein to organize heterochromatin by linking heterochromatin with the nuclear lamina (Supplementary Fig. 11). Interestingly, deficiencies in Dicer, which processes pre-miRNAs into mature miRNAs, also induce heterochromatin disorganization and increase genomic instability in Drosophila40. In contrast to the protein-protein interaction between DGCR8 and heterochromatin components that we report in this study, deficiencies in Dicer disrupt localization of H3K9me2 by altering the specificity of siRNA-mediated targeting of H3K9me2. Despite utilizing different mechanisms, these two studies demonstrate the long-overlooked, non-canonical roles of these two miRNA-processing proteins in regulating nuclear and chromatin organization.
Previous studies on DGCR8 have primarily focused on its miRNA-processing function using DGCR8-knockout cell lines and animal models. DGCR8-deficient mouse fibroblasts and neural stem cells (NSCs) can be reprogrammed into iPSCs with lower efficiency due to the lack of canonical miRNAs41. Consistently, iPSCs derived from DGCR8-deficient fibroblasts and NSCs exhibit poor differentiation potentials, similar to DGCR8-deficient mESCs, which fail to downregulate pluripotency markers and therefore do not differentiate19. While all of these studies indicate that DGCR8 is required for the self-renewal and differentiation of mouse pluripotent stem cells in a miRNA-dependent manner, the potential for DGCR8 to regulate physiological functions in human stem cells remains unclear. In our multiple efforts to generate human ESC lines with DGCR8 defects for studying the novel functions of DGCR8, DR8ex2 hESCs with complete protein ablation failed to maintain pluripotency, and DR8ex3 hESCs, which expressed a 50-kDa truncated protein, exhibited genomic instability (primarily an intra-chromosomal translocation). However, DR8dex2 hESCs, which lacked the N-terminus of DGCR8, had normal self-renewal ability and pluripotency, but compromised proliferation upon differentiation into hMSCs. The diverse characteristics related to different DGCR8 defects suggest that the regulatory activities by DGCR8 are cell-type and developmental-stage dependent.
With previous studies having mainly utilized mouse cell lines, our study employed the CRISPR/Cas9-mediated gene editing techniques in human ESCs, followed by the directed differentiation of these ESCs to elucidate how DGCR8 regulates the homeostasis and senescence of adult stem/progenitor cells. Genetically modified DR8dex2 hESCs have the potential to self-renew, to differentiate into various adult stem cells and further into terminally differentiated cells. Previous studies have shown that the C-terminal domains of DGCR8 mainly participate in miRNA processing. We used DR8dex2 hMSCs to reveal that an intact DGCR8 N-terminus is required for heterochromatin organization and to protect against premature senescence. Strikingly, functional inactivation of DGCR8 in DR8dex2 hMSCs could be phenotypically rescued by expressing DR8-mtDRBD, which was deficient in miRNA processing. Moving forward, DR8dex2 hESCs provide an invaluable platform for investigating the cell type-specific consequences of DGCR8 in human aging-associated disorders, with implications in induced neural cells, cardiomyocytes, and vascular cells, etc., without the confounding effects of DGCR8-associated miRNA clusters.
We and others have recently provided evidence that disorganization of the nuclear envelope and heterochromatin is a driving force for hMSC aging, and that these aging processes can be targeted either by gene correction of disease-associated mutations, by gene editing-mediated cellular enhancement, or by treatment with geroprotective compounds4,7,9,15,24,42. In this study, we demonstrate that DGCR8 is a key regulator of human cellular aging via a heterochromatin-stabilizing mechanism. DGCR8 was downregulated in prematurely and physiologically aging hMSCs, and DGCR8 overexpression effectively alleviated senescent phenotypes. Notably, the accumulation of senescent cells and their secretion of SASP factors in many tissues are considered as risk factors for age-related pathologies, like osteoarthritis33,43. Due to the complexity and heterogeneity of osteoarthritis pathogenesis, the combined removal of local senescent cells and the transplantation of MSCs may be a good therapeutic option to date33,43,44,45. Here, we provide strong in vivo evidence that osteoarthritic mice that received gene therapy (using lentiviruses encoding DR8-WT or DR8-mtDRBD) exhibited reduced inflammation and increased osteogenesis- and chondrogenesis-related gene expression, suggesting enhanced regeneration in the articular cavity, and effective restoration of cartilage structure. Thus, this is a promising strategy for treating age- or injury-associated osteoarthritis. Given the fact that DR8-mtDRBD does not affect miRNA processing, this modified protein may represent a robust and safer gene therapeutic tool for treating osteoarthritis. Taken together, our study provides the first evidence that DGCR8 is a gatekeeper for maintaining heterochromatin organization, highlighting the fact that DGCR8-heterochromatin may be a novel target for understanding and hopefully treating aging-associated disorders in the future.
Wildtype (WT) human ESCs (hESCs, Line H9, WiCell Research Institute) and their genetic modified derivatives were maintained on feeder cells (mitomycin C-inactivated mouse embryonic fibroblast, MEF) in hESC culture medium containing DMEM/F12 (Thermo Fisher Scientific), 20% Knockout Serum Replacement (Thermo Fisher Scientific), 0.1 mM non-essential amino acids (NEAA, Thermo Fisher Scientific), 2 mM GlutaMAX (Thermo Fisher Scientific), 55 μM β-mercaptoethanol (Thermo Fisher Scientific), and 10 ng/ml bFGF (Joint Protein Central) or on Matrigel (BD Biosciences) in mTeSR medium (STEMCELL Technologies). Both Human MSCs (hMSCs) derived from hESCs and primary human MSCs were cultured in MEMα (Thermo Fisher Scientific) medium supplemented with 10% fetal bovine serum (FBS, Thermo Fisher Scientific (Cat:10099-141, Lot:1616964)), 0.1 mM non-essential amino acids (Thermo Fisher Scientific), 1% penicillin/streptomycin (Thermo Fisher Scientific), and 1 ng/ml bFGF (Joint Protein Central, JPC).
Antibodies used for western blotting included anti-DGCR8 (Ab191875, Abcam, 1:1000), anti-KAP1 (Ab22553, Abcam, 1:2000), anti-Lamin B1 (Ab16048, Abcam, 1:1000), anti-HP1α (#2616 S, Cell signaling, 1:1000), anti-HP1γ (#2619, Cell signaling, 1:3000), anti-LAP2 (611000, BD Biosciences, 1:1000), anti-P16 (550834, BD Biosciences, 1:500), anti-P21 (2947, Cell signaling, 1:1000), anti-GATA4 (sc-9053, SCBT, 1:1000), anti-HDAC1 (sc-7872, SCBT, 1:1000), anti-LBR (Ab32535, Abcam, 1:1000), anti-SUV39H1 (05615, Millipore, 1:1000), anti-Drosha (Ab12286, Abcam, 1:1000), anti-β-actin (sc-69879, SCBT, 1:5000), anti-GAPDH (sc-25778, SCBT, 1:3000). Antibodies used for immunostaining included anti-DGCR8 (Ab191875, Abcam, 1:1000), anti-Ki67 (VP-RM04, Vector, 1:800), anti-γH2AX (05636, Millipore, 1:400), anti-53BP1 (A300-273A, Bethyl Laboratories, 1:500), anti-FOXA2 (8186 S, Cell signaling, 1:100), anti-SMA (A5228, Sigma Aldrich, 1:100), anti-TuJ1 (T2220, Sigma Aldrich, 1:100), anti-H3K9me3 (Ab8898, Abcam, 1:800), anti-Lamin B (sc-6217, SCBT, 1:250), anti-LAP2 (611000, BD Biosciences, 1:500), anti-OCT3/4 (sc-5279, SCBT, 1:200), anti-SOX2 (sc-17320, SCBT, 1:100), anti-NANOG (Ab21624, Abcam, 1:200). Antibodies used for flow cytometry included anti-CD105 (17-1057-42, eBioscience, 1:100), anti-CD73 (550257, BD Biosciences, 1:100), anti-CD90 (555595, BD Biosciences, 1:100), anti-CD34 (555822, BD Biosciences, 1:100), anti-CD43 (580198, BD Biosciences, 1:100), anti-CD45 (555482, BD Biosciences, 1:100). Antibodies used for IHC included anti-P16 (Ab54210, Abcam, 1:1000), anti-Ki67 (GB13030-2, Servicebio, 1:500).
CRISPR/Cas9-mediated DGCR8 editing in hESCs
CRISPR/Cas9-mediated gene editing was performed as previously described with modifications22. In brief, guide RNAs (gRNAs) were designed at the website http://crispr.mit.edu. The coding sequences of gRNAs targeting exon 2 or exon 3 of DGCR8 were cloned into pCAG-mCherry-gRNA46. Upon treatment with ROCK inhibitor (Y-27632, TOCRIS) for 24 h, hESCs were electroporated by 4D-Nucleofector (Lonza) with both a gRNA vector and pCAG-1BPNLS-Cas9-1BPNLS-2AGFP46. Cells were then seeded on Matrigel-coated plates and cultured with mTeSR medium. After expansion for 48 h, EGFP/mCherry-dual-positive cells were collected by FACS and cultured on MEF feeders in hESC culture medium. Clones were manually picked and expanded for further characterization.
CRISPR/Cas9-mediated generation of DR8dex2 hESCs
CRISPR/Cas9-mediated gene editing was performed as previously described with modifications47. In brief, a donor plasmid was constructed with two 1.0–2.0-kb homology arms and a drug resistance cassette (neomycin). A gRNA targeting exon 2 of DGCR8 was cloned into a gRNA cloning vector (#41824, Addgene) (DGCR8-gRNA). Upon treatment with ROCK inhibitor (Y-27632, TOCRIS) for 24 hours, hESCs (5 × 106) were resuspended in 100 μl opti-MEM (Thermo Fisher Scientific) containing 7 μg hCas9 (#41815, Addgene), 7 μg DGCR8-gRNA and 7 μg donor plasmid and electroporated by 4D-Nucleofector (Lonza). Cells were then seeded on MEF feeder cells. G418 (100 μg/ml, Thermo Fisher Scientific) was added to initiate positive selection 2–4 days after electroporation. After G418 selection and clonal expansion for about 2 weeks, G418-resistant clones were picked and transferred to a 96-well plate for further characterization and expansion. The identification of gene-targeted clones was performed by genomic PCR and western blotting. Additionally, the pCAG-FLpo-2A-puro vector was used to remove the neomycin-resistance cassette in DR8 dex2 hESCs. Three days after the transfection, puromycin (1 μg/ml, Thermo Fisher Scientific) was used to enrich puro-resistant cells for 48 h. Ten days later, the emerging colonies were picked and expanded for future application.
Lentiviral CRISPR/Cas9-mediated DGCR8 or LMNB1 knockdown
Lentiviral CRISPR/Cas9-mediated gene editing was conducted as previously described48,49. Briefly, the sgRNA targeting DGCR8 or LMNB1 was cloned into lenti-CRISPRv2 (#52961, Addgene) with an hSpCas9 expression cassette (pLenti-CRISPRv2-DGCR8 or pLenti-CRISPRv2-LMNB1). Lentiviruses were then packaged with pLenti-CRISPRv2-DGCR8 or pLenti-CRISPRv2-LMNB1 and used to transduce WT hMSCs. Cells were treated with 1 μg/ml puromycin (Thermo Fisher Scientific) for screening at 72 h after the infection.
Generation and characterization of hMSCs
hMSCs were differentiated from hESCs as previously described24,50. Briefly, embryoid bodies were plated on Matrigel-coated plates in hMSC differentiation medium (MEMα (Thermo Fisher Scientific), 10% fetal bovine serum (FBS, Thermo Fisher Scientific (Cat:10099-141, Lot:1616964)), 1% penicillin/streptomycin (Thermo Fisher Scientific), 1 ng/mL bFGF (Joint Protein Central) and 5 ng/mL TGFβ (Humanzyme)) for 10 days. The confluent fibroblast-like cells were then maintained in hMSC culture medium and FACS-sorted (BD FACS Aria II) for the purification of CD73/CD90/CD105 tri-positive hMSCs. The functionality of hMSCs was verified by differentiation to osteoblasts, chondrocytes, and adipocytes of the sorted hMSCs24,50.
Isolation and culture of primary hMSCs
Primary hMSCs were isolated from the dental pulp of different individuals with the approval from the Ethics Committee of the 306 Hospital of PLA in Beijing44. The tissue block was cut using scissors in 1x TrypLE™ Express Enzyme plus Dispase IV and further digested at 37 °C for 30 min. The digestion mixture was passed through a 100 μm cell strainer and centrifuged at 500 g for 10 minutes at room temperature. The supernatant was carefully decanted, the pellet resuspended in MSC culture medium and plated onto gelatin-coated plates. After overnight incubation, the medium was changed every other day until the cells reached 80% confluency (4–5 days post-harvest).
Protein, DNA, and RNA analyses
For western blot, cells were lysed in RIPA buffer with protease inhibitor cocktail (Roche). Protein lysates were quantified using a BCA quantification kit (Thermo Fisher Scientific), subjected to SDS-PAGE and electrotransferred to PVDF membranes (Millipore). Then the membranes were blocked with 5% milk, and incubated with primary antibodies and horseradish peroxidase (HRP)-conjugated secondary antibodies. The results of western blotting were obtained by ChemiDoc XRS system (Bio-Rad). For RT-qPCR, total RNA was extracted in TRIzol (Thermo Fisher Scientific) and genomic DNA was removed using a DNA-free kit (Thermo Fisher Scientific). cDNA was generated by the GoScript Reverse Transcription System (Promega). RT-qPCR was performed using the qPCR Mix (TOYOBO) in a CFX384 Real-Time system (Bio-Rad). For genomic PCR, a DNA extraction kit (TIANGEN) was used to extract genomic DNA and PCR was carried out with PrimeSTAR (TAKARA).
For immunofluorescence, cells seeded on coverslips (Thermo Fisher Scientific) were fixed in 4% paraformaldehyde (PFA) for 20 min, permeabilized with 0.4% Triton X-100 in PBS for 1 hour, and blocked with 10% donkey serum in PBS (Jackson ImmunoResearch) for 1 h at room temperature. Cells were then incubated with primary antibodies in the blocking buffer at 4 °C overnight, washed with PBS three times, incubated with secondary antibodies at room temperature for 1 h, washed again with PBS three times, and counterstained with Hoechst 33342 (Thermo Fisher Scientific). Leica SP5 confocal system was used for immunofluorescence microscopy. DNA texture image in one nucleus was measured with Coefficient of Variation (C.V) as previously described9. The degree of variation of all pixel value in one nucleus was measured by C.V of DNA texture image with ImageJ (C.V = Standard deviation/Mean).
Transmission electron microscope (TEM)
One million cells of either WT or DR8dex2 hMSCs were harvested enzymatically by TrypLE (Thermo Fisher Scientific) and fixed with 4% PFA in PBS, pH 7.4, on ice for 2 h. Cells were subsequently dehydrated in a graded series of ethanol, infiltrated and embedded in Lowicryl resin HM20. Two hundred nanometer sections were obtained and imaged by a Spirit transmission electron microscope (FEI Company) operating at 100 kV.
Mutagenesis of DGCR8
The vector expressing DGCR8 (mtDRBD) was generated by introducing multipoint mutations at coding sequences of two DGCR8 dsRBDs as previously described25. Point mutagenesis was carried out using a Fast Multisite Mutagenesis System (TRANSGEN BIOTECH) according to the manufacturer’s instructions.
SA-β-gal staining assay
SA-β-gal staining of hMSCs was conducted as previously described51. Briefly, cells were fixed in 2% formaldehyde and 0.2% glutaraldehyde at room temperature for 4 min and stained with fresh staining solution at 37 °C overnight. SA-β-gal-positive cells were counted in randomly selected fields by Image J software.
Clonal expansion assay
Cells were seeded at 2000 cells per well in 12-well plates and cultured for around 10 days. Cells were fixed in 4% PFA for 30 minutes and stained with 0.2% crystal violet for 1 h at room temperature. Cell numbers were counted in randomly selected fields and staining areas were measured by Image J software.
AP staining analysis
hESCs were seeded at 2000 cells per well in 12-well plates and cultured for around 2 weeks. Cells were fixed and stained with Alkaline Phosphatase Stain Kit (Shanghai Yeasen Biotechnology Co. Ltd. China) according to the instructions. Stained areas were measured by Image J software (NIH).
For packaging of lentiviruses, HEK293T cells were co-transfected with lentiviral vectors, psPAX2 (#12260, Addgene) as well as pMD2.G (#12259, Addgene). Viral particles were collected by ultracentrifugation at 19,400 g at 4 °C for 2.5 h.
For exogenous Co-immunoprecipitation (Co-IP), HEK293T cells transfected with plasmid expressing Flag-DGCR8 were lysed in CHAPS lysis buffer (120 mM NaCl, 0.3% CHAPS, 1 mM EDTA, 40 mM HEPES, pH 7.5, and 1x complete protease inhibitor cocktail (Roche)). For endogenous Co-IP, hMSCs were lysed in CHAPS lysis buffer. HEK293T cells or hMSCs were lysed at 4 °C for 2 h, and then centrifuged at 12,000 g at 4 °C for 30 min. For endogenous Co-IP, lysates (1 mg protein) were pre-cleared with 20 μl of Protein A/G-PLUS Agarose beads (Santa Cruz) for 2–4 h, and then the supernatants were collected by centrifugation at 3 000 rpm at 4 °C for 3 min. The supernatants mixed with the indicated antibodies and beads were rotated overnight at 4 °C. The immunocomplexes were washed with CHAPS buffer three times and then eluted by boiling in 1 × SDS-loading buffer for 10 min.
The elution proteins of immunoprecipitation were separated by 12% SDS-PAGE gel and visualized by Pierce™ Silver Stain (Thermo Fisher Scientific). Following decoloration and dehydration of the gels, the proteins were digested with sequencing-grade trypsin (Worthington). Subsequently, the peptides were extracted from gel pieces with 0.1% formic acid and 50% acetonitrile, dried in a vacuum centrifuge (ThermoFisher, San Jose, CA), dissolved in 0.1% formic acid (FA) and separated by a C18 reverse phase column (75 μm×20 cm, 3 μm). The column was then eluted with a linear gradient of 5-30% acetonitrile in 0.2% formic acid at the rate of 300 nL/min for 100 minutes. The mass spectra were acquired by nanoLC-Q EXACTIVE (ThermoFisher, San Jose, CA) equipped with a nano-ES ion source (Proxeon Biosystems, Denmark). Full scan spectra (from m/z 300–1600) were acquired in the Orbitrap analyzer with a resolution of 60,000 at 400 m/z after the accumulation of 1,000,000 ions. The five most-intense ions per scan were selected for collision-induced dissociation (CID) fragmentation in the linear ion trap after the accumulation of 3000 ions. The maximal filling times were set at 500 ms for the full scans and 150 ms for the MS/MS scans. The dynamic exclusion list was restricted to a maximum of 500 entries with a maximum retention period of 60 s and a relative mass window of 10 ppm.
All raw files were processed with the MaxQuant software (Version 18.104.22.168). The generated peak list files were searched with Thermo Proteome Discoverer (22.214.171.1248) against the UniProt-proteome-human database (update-20160226). The search parameters were set as follows: enzyme was trypsin; up to two missed cleavages; alkylated cysteine as fixed modification; oxidation methionine as variable modifications. MS tolerance was 10 ppm while MS/MS tolerance was 0.02 Da. The required false discovery rate (FDR) was set to 1% at peptide and protein levels, and the minimum required peptide length was seven amino acids. At least one unique or razor peptide per protein group was required for protein identification.
ChIP-qPCR and ChIP-seq
ChIP was performed according to a previous protocol with slight modifications52. Briefly, 1 × 106 WT or DR8dex2 hMSCs were cross-linked in 1% vol/vol formaldehyde/PBS for 10 minutes at room temperature and then stopped by 125 mM Glycine for 5 min at room temperature. Samples were lysed on ice for 10 min. After sonicating by a Bioruptor® Plus sonication device (Diagenode), supernatants were incubated overnight at 4 °C with Protein G dynabeads (Thermo Fisher Scientific, 10004D) associated with 2.4 μg anti-DGCR8, H3K9me3 antibody or rabbit IgG. Subsequently, elution and reverse cross-link were performed at 68 °C for 2 h on a thermomixer. Then the DNA was isolated using phenol-chloroform-isoamylalcohol extraction and ethanol precipitation. The purified DNA was subjected to qPCR to evaluate DGCR8 or H3K9me3 occupation at satellite and LINE1 sites. The enriched fragments were constructed into libraries without the incorporation of spike-in controls via KAPA Hyper Prep Kits with PCR Library Amplification/Illumina series (KK8504) following the manufacturer’s instructions.
RNA-seq and miRNA-seq
Total cellular RNA was extracted using TRIzol (Thermo Fisher Scientific) from 1 × 106 cells per duplicate and genomic DNA was removed using a DNA-free kit (Thermo Fisher Scientific). Quality control and sequencing was done by Novogene Bioinformatics Technology Co. Ltd.
DNeasy Blood & Tissue Kit (Qiagen) was applied to extract genomic DNA from 1 × 106 cells per duplicate. Quality control and sequencing were done with standard protocols in Novogene Bioinformatics Technology Co. Ltd.
RNA-seq data analysis
Paired-end raw reads were trimmed with Trim Galore (v0.4.5, http://www.bioinformatics.babraham.ac.uk/projects/trim_galore/) and cleaned reads were mapped to UCSC hg19 human genome or UCSC mm10 mouse genome using hisat2 (v2.0.4)53 with default parameters. Aligned reads were then annotated and counted by HTSeq (v0.6.1)54. Differentially expressed genes (DEGs) were calculated using DESeq2 with the cutoff as follows: Benjamini–Hochberg adjust P value (p.adjust) < 0.05 and absolute log2 (fold change) > 0.5. The Pearson correlation coefficient (R) of RNA-seq replicates was computed based on DESeq2 regularized logarithm (rlog) normalized read count55. GO enrichment analysis was performed using ClusterProfiler package (v3.6.0)56 at Benjamini–Hochberg adjust P value (p.adjust) < 0.05.
ChIP-seq data analysis
The processing of ChIP-seq data followed standard pipeline without spike-in normalization9,57,58,59. Firstly, low-quality reads and adapters were trimmed by Trim Galore (v0.4.5). Then the cleaned reads were aligned to UCSC hg19 human reference genome using bowtie2 (v2.2.9)60 and PCR duplicates were removed by Samtools (v1.6)61. For H3K9me3 peak calling, MACS2 (v2.1.1)62 was used with default parameter.
The definition of “H3K9me3 mountains” was described previously9. Briefly, bam files of replicates were merged after alignment and called peaks by MACS2. Neighboring peaks were combined and “H3K9me3 mountains” were filtered at the cutoff 10 kb length. H3K9me3 mountains were then merged to union ones at the maximum distance between features 1 Mb.
Pearson correlation coefficient of replicates was computed based on DESeq2 regularized logarithm (rlog) normalized read count to evaluate the reproducibility.
For H3K9me3 ChIP-seq signal enrichment analysis at the regions of LADs and heterochromatin, we retained H3K9me3 signal enriched LAD and heterochromatin genomic regions. The average H3K9me3 signals were shown based on RPKM (Reads Per Kilobase Million) normalized read count on 10 bp bin size by deepTools263. To investigate H3K9me3 signals of the lost “H3K9me3 mountains” regions in DR8dex2 hMSCs transduced with Luc, DR8-WT, or DR8-mtDRBD, heatmap was shown based on CPM (Counts Per Million) normalized read count.
For genomic elements enrichment analysis, ChIPseeker (v1.14.1)64 was used to annotate H3K9me3 ChIP-seq peaks. Genomic features were divided into four categories, “Promoter”, “Exon”, “Intron”, and “Intergenic”. For heterochromatin and LAD annotation of H3K9me3 ChIP-seq signal, the genomic locations of heterochromatin regions were obtained from Roadmap Epigenomics Mapping Consortium65. The genomic locations of LADs were downloaded from the UCSC hg19 goldenPath.
microRNA-seq data analysis
The mirDeep2 pipeline was implemented for microRNA-seq data processing66. In brief, raw data was trimmed and then aligned to hg19 genome using bowtie. Known microRNAs were annotated by miRBase database67. Then the reads of each known microRNA were counted and the downstream analyzed by DESeq2. Normalized read count of each microRNA by DESeq2 size factor function was used for heatmap drawing.
Copy number variation data analysis
Copy number variation (CNV) analysis was performed according to previously described pipeline68. Briefly, paired-end reads were trimmed with Trim Galore (v0.4.5) and then mapped to UCSC hg19 human genome using bowtie2 (v2.2.9)60. The CNVs were estimated and corrected by HMMcopy (v1.25.0) in each 0.5 Mb window69. The CNV plot was generated by R package circlize (v0.4.5)70.
Teratoma formation assay
NOD/SCID mice were injected with 3 × 106 hESCs suspended in a Matrigel/mTeSR (1:4) solution. Teratomas were collected at around 8 weeks after injection for further analysis.
hMSC transplantation assay
In all, 1 × 106 WT and DR8dex2 hMSCs pretransduced with lentiviruses expressing luciferase were injected into tibialis anterior (TA) muscle of male nude mice. 1/3/5 days after transplantation, mice were imaged by IVIS spectrum imaging system (XENOGEN, Caliper) for luciferase activity detection.
ACLT-induced OA mouse model
An OA mouse model was generated as previously described33. Briefly, ACLT was performed on male C57BL/6 mice aged 8 weeks. For the ACLT surgery, after opening the joint capsule, the ACL was transected with microscissors under a surgical microscope. One week after surgery, lentiviruses expressing Luc, DR8-WT, or DR8-mtDRBD were injected into the articular cavity. At 7 weeks after lentivirus injection, the mice knee joints were examined by microCT analysis. The mice were then euthanized and the joints were collected for histological assessment of the medial tibial plateau joint and RNA-seq analysis. Total RNAs from three knee joints of each experimental group were mixed and then separated into three technical repeats for RNA-seq.
Histology and immunohistochemistry
Collected mouse joints were fixed in 4% PFA for 2 days, decalcified in 5% methanoic acid for 7 days and embedded in paraffin. Sections (5 μm) were cut from the paraffin blocks, stained with Fast Green FCF (0.02%) and safranin O (0.1%), and quantified using the Osteoarthritis Research Society International (OARSI) scoring system.
Immunohistochemical staining was performed using DAB staining method. Briefly, the slides were deparaffinized and rehydrated. Antigen retrieval was performed using 1 mg/ml Trypsin in PBS (Sigma-Aldrich) digestion for 10 min. Slides were then incubated with endogenous peroxidase inhibitor (Beijing Zhongshan Jinqiao Biological Technology Co., Ltd.) at room temperature for 10 minutes to block endogenous peroxidase activity. Slides were washed and blocked with blocking solution (10% donkey serum in PBS) for 1 h at room temperature and then incubated with a primary antibody diluted in blocking solution overnight at 4 °C. Subsequently, slides were washed and incubated with secondary antibodies at room temperature for 30 min. After washed by PBS, the slides were visualized using DAB substrate (applied for less than 5 min until the desired color intensity was reached) and counterstained with hematoxylin.
Results were presented as mean ± SEM. Two-tailed Student’s t-test was conducted using Graph-Pad Prism Software. P values < 0.05 were considered statistically significant (*).
All animal experiments were conducted according to animal protocols approved by the Chinese Academy of Science Institutional Animal Care and Use Committee. Mice were housed under a 12-hour light/dark cycle and supplied with food and water ad libitum. Mice were anaesthetized using isoflurane and euthanized with CO2 followed by cervical dislocation.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
Raw sequencing data and processed data of RNA-seq, microRNA-seq and ChIP-seq have been deposited in the NCBI Gene Expression Omnibus (GEO) under the accession number GSE113117, GSE121029, GSE116303, GSE113181 and GSE130206. The whole genome sequencing data have been deposited in the NCBI Sequence Read Archive under accession number PRJNA541163. The proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD013856. The authors declare that all the data supporting the findings of this study are available within the article and its supplementary information files or from the corresponding author upon reasonable request. The source data underlying Figs. 1f, 1g, 2c-e, 2g, 2i-k, 3, 4c, 4f, 4g, 5, 6a, 6c-f, 7b-e, 8a, 9b-d and Supplementary Figs. 1h-k, 2b-d, 2f, 2h-i, 2k-m, 3f, 4d, 4f, 4g, 7a, 8a, 9c-j, and 10 are provided as a Source Data file. The uncropped blots and gels of Figs. 1b, 2b, 2f, 4a, 4b, 6b and Supplementary Figs. 1b, 1e, 1l-n, 2g, 2j, 3a, 3e, 4c, 4e, 7g, 7h, 9a and 9b are provided as Supplementary Figures 13-19.
High-throughput sequencing data analyses were performed using custom Python, Perl and R codes based on released softwares, as described in the Methods. All codes used in this study are available upon request from the corresponding authors. The software for data processing is available from the corresponding author upon request.
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We are grateful to L. Bai, R. Bai, J. Lu, Q. Chu, Y. Yang and S. Ma for administrative assistance; W. Li, J. Jia for assisting with animal experiments; J. Jia for the FACS; J. Hao for the immunohistochemistry; X. Shi, X. Wu, M. Wang, S. Yang, S. Duo, K. Du, and L. Zhou for management of laboratory animals. We thank all the members of the Liu lab for discussion and technical assistance. High-throughput sequencing data were processed on the “Era” petascale supercomputer of Computer Network Information Center of Chinese Academy of Sciences. This work was supported by the Strategic Priority Research Program of the Chinese Academy of Sciences (XDA16010100), the National Key Research and Development Program of China (2015CB964800, 2018YFC2000100, 2017YFA0102802, 2017YFA0103304, 2018YFA0107203, 2018YFC2000400), the National Natural Science Foundation of China (81625009, 81330008, 91749202, 91749123, 31671429, 81671377, 81771515, 31601158, 81701388, 81601233, 31601109, 81822018, 81870228, 81801399, 31801010, 81801370, 81861168034), Key Research Program of the Chinese Academy of Sciences (KJZDEWTZ-L05), Beijing Municipal Commission of Health and Family Planning (PXM2018_026283_000002), Advanced Innovation Center for Human Brain Protection (3500-1192012) and the State Key Laboratory of Membrane Biology. P.G. and J.C.I.B. were supported by Fundación Dr. Pedro Guillen, Fundación Teléfonica, Fundación MAPFRE and Universidad Católica San Antonio de Murcia (UCAM).
The authors declare no competing interests.
Peer review information: Nature Communications thanks Sandra Peiro and other anonymous reviewer(s) for their contribution to the peer review of this work.
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