DDX18 coordinates nucleolus phase separation and nuclear organization to control the pluripotency of human embryonic stem cells

Shi, Xianle; Li, Yanjing; Zhou, Hongwei; Hou, Xiukun; Yang, Jihong; Malik, Vikas; Faiola, Francesco; Ding, Junjun; Bao, Xichen; Modic, Miha; Zhang, Weiyu; Chen, Lingyi; Mahmood, Syed Raza; Apostolou, Effie; Yang, Feng-Chun; Xu, Mingjiang; Xie, Wei; Huang, Xin; Chen, Yong; Wang, Jianlong

doi:10.1038/s41467-024-55054-8

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Published: 30 December 2024

DDX18 coordinates nucleolus phase separation and nuclear organization to control the pluripotency of human embryonic stem cells

Nature Communications volume 15, Article number: 10803 (2024) Cite this article

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Abstract

Pluripotent stem cells possess a unique nuclear architecture characterized by a larger nucleus and more open chromatin, which underpins their ability to self-renew and differentiate. Here, we show that the nucleolus-specific RNA helicase DDX18 is essential for maintaining the pluripotency of human embryonic stem cells. Using techniques such as Hi-C, DNA/RNA-FISH, and biomolecular condensate analysis, we demonstrate that DDX18 regulates nucleolus phase separation and nuclear organization by interacting with NPM1 in the granular nucleolar component, driven by specific nucleolar RNAs. Loss of DDX18 disrupts nucleolar substructures, impairing centromere clustering and perinucleolar heterochromatin (PNH) formation. To probe this further, we develop NoCasDrop, a tool enabling precise nucleolar targeting and controlled liquid condensation, which restores centromere clustering and PNH integrity while modulating developmental gene expression. This study reveals how nucleolar phase separation dynamics govern chromatin organization and cell fate, offering fresh insights into the molecular regulation of stem cell pluripotency.

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Introduction

In eukaryotes, the linear genome is folded and packaged into three-dimensional chromatin structures in the nucleus, a highly organized and dynamic organelle^1,2,3. The organization and reconfiguration of chromatin into different compact versus open states allow fine-tuning gene regulation during cell differentiation and embryonic development. Pluripotent embryonic stem cells (ESCs) are ideal for studying gene expression programs and developmental control due to their unlimited self-renewal capacity and multilineage differentiation properties that often co-occur with the acquisition of the heterochromatin compartment. In particular, perinucleolar heterochromatin (PNH) formation is required for both mouse^4,5 and human⁶ ESC differentiation and early embryo development^7,8.

Transcription factors, structural RNAs, and RNA binding proteins (RBPs) have been identified as important regulators for chromatin organization through multivalent interactions. The self-assembly of these macromolecules and the immiscibility of each, known as phase separation^9,10, give rise to a series of membraneless sub-compartments (or nuclear bodies), including nucleoli¹¹, paraspeckles¹², P granules¹³, and nuclear speckles¹⁴. The mammalian nucleolus is the largest nuclear body assembled around the tandem repeated rDNA clusters. Each nucleolus is composed of three morphologically and functionally distinct sub-regions named the fibrillar center (FC), dense fibrillar center (DFC), and granular component (GC). Aside from the classical function in ribosome biogenesis (RiBi)^15,16, nucleoli are also involved in phase-separated protein quality control¹⁷ and stem cell differentiation, the latter of which co-occurs with the decrease in RiBi^4,18,19,20. Although widely recognized for their roles in providing an inter-chromosomal hub for 3D chromatin organization^21,22, nucleoli functions and the underlying mechanisms in chromatin organization and reconfiguration during cell differentiation or development remain poorly understood.

DEAD-box RNA helicases are evolutionarily conserved regulators of RNA-containing phase-separated organelles²³, including nucleoli²⁴. DDX18 was one of the 86 proteins whose depletion in the siRNA screen impacted nucleolar structure maintenance in HeLa cells²⁵. By studying DDX18 in human ESCs (hESCs), we discovered that DDX18 plays a critical role in controlling human pluripotency by maintaining high RiBi and highly organized membraneless nucleolar architecture. Mechanistically, DDX18 modulates liquid-liquid phase separation (LLPS) properties of the nucleoli through its intrinsically disordered regions (IDRs) and the dynamic multivalent interactions with nucleolar RNAs and proteins. Specifically, heterotypic interactions of DDX18 with nucleophosmin (NPM1) in nucleolar GC and with snoRNAs and rRNAs in the DFC/FC interface modulate the biophysical properties of the nucleolus, limiting the heterochromatin formation and preventing the initiation of differentiation program in hESCs. Finally, we developed a DDX18-mediated nucleolus-specific CasDrop²⁶, dubbed NoCasDrop. We successfully manipulated centromere clustering around the nucleolus, demonstrating that the redirection of constitutive heterochromatin organization towards a different compartment alone is sufficient to drive the derepression of development-related genes, initiating an early event associated with the exit of pluripotency and cell differentiation.

Results

DDX18 is required for hESC maintenance through the regulation of RiBi and protein synthesis

DDX18 is highly enriched in pluripotent hESCs relative to differentiated²⁷ or somatic²⁸ cells (Fig. S1a) and abruptly downregulated upon embryoid body (EB) differentiation at kinetics even faster than NANOG and OCT4 (Fig. S1b), suggestive of its specific function in hESC maintenance. shRNA-mediated DDX18 knockdown (KD) (shDDX18) at both protein (Fig. 1a) and RNA (Fig. 1b) levels, relative to control KD (shLuc), functionally validated its critical roles in maintaining hESCs, manifested by downregulation of the core pluripotency and stem cell surface marker genes and upregulation of lineage-specific and developmental genes (Fig. 1a–c), as well as reduced alkaline phosphatase staining (Fig. 1d). RNA sequencing (RNA-seq) in control and DDX18 KD cells (Fig. 1e) revealed 789 significantly downregulated genes for “stem cell population maintenance”, “rRNA processing”, and “translational initiation”, and 1903 significantly upregulated genes for multilineage differentiation and development based on both the gene ontology (GO) analysis (Fig. 1f) and gene set enrichment analysis (GSEA) (Figs. 1g, S1c, d). Of note, the “ectoderm differentiation” is not enriched for the upregulated genes (Fig. S1c, d), indicating lineage skewing towards mesendoderm differentiation of DDX18 KD hESCs (Fig. S1e). When compared to the transcriptomes of the three germ layers²⁹, the transcriptome of DDX18 KD cells clustered with those of endoderm and mesoderm but not ectoderm cells, further highlighting the skewed mesendoderm differentiation of DDX18 KD cells (Fig. S1f). Among the upregulated genes enriched with the “embryonic organ development” (Fig. 1g) are HOXA-D clusters (e.g., HOXA in Fig. 1h), key developmental regulators that are not expressed but held in bivalent chromatin domains in pluripotent stem cells to be resolved and activated upon differentiation^30,31,32.

**Fig. 1: DDX18 is required for hESC maintenance.**

The prevalent association of translation-related GO and biological processes with downregulated genes upon DDX18 KD prompted us to investigate the potential translational defects of DDX18 KD hESCs. First, a SUnSET assay to measure the newly synthesized peptides³³ revealed a marked decrease of total protein synthesis in DDX18 KD relative to control cells, a global translational defect that can be rescued with wild-type DDX18 (WT) but not the RNA helicase inactive DQAD mutant^34,35 (Fig. 1i). Next, SILAC-MS (stable isotope labeling by amino acids coupled with mass spectrometry) quantitative proteomics (Supplementary Data 1) identified 879 downregulated and 319 upregulated proteins with the enrichment of translation-related and differentiation-related GO terms, respectively, in DDX18 KD relative to control KD hESCs (Fig. S1g, h). By integrating the RNA-seq and SILAC data (Fig. S1i), we found that, while many genes/proteins are positively regulated at both transcriptional and translational levels (Groups I and VI), there are more genes/proteins regulated at translational levels without significant transcriptional changes (Groups II and V). Only a small number of genes/proteins exhibit anti-correlation between transcription and translation (Groups III and IV) (Fig. S1i; Supplementary Data 2). Furthermore, consistent with our previous findings in mouse ESCs (mESCs)¹⁹, our DDX18 ChIP-seq experiment in hESCs revealed that no significant DDX18 binding signals could be detected in other genomic regions beyond the rDNA loci (Fig. S5f), excluding the direct transcriptional effects of DDX18 on the pluripotency and development gene expression programs except for the rDNA gene expression.

Together, these results establish the roles of DDX18 and its RNA helicase activity in maintaining hESCs through its nucleolar functions in RiBi and translational control, a finding in line with emerging evidence linking translational control and protein synthesis to ESC maintenance and embryonic development^20,36,37.

DDX18 undergoes phase separation in vitro and in vivo

To understand how DDX18 may exert its nucleolar functions in regulating RiBi and translation for hESC pluripotency, we first performed 3D structured illumination microscopy (3D-SIM) and confirmed its sub-nucleolar localization in the NPM1-containing GC layer in hESCs (Fig. 2a, b). Nucleolus assembly is driven by liquid-liquid phase separation (LLPS), facilitated by nucleolar proteins harboring intrinsically disordered regions (IDRs) that mediate multivalent interactions to assist macromolecular assembly^24,38,39. Employing PONDER VSL2 webtool⁴⁰ for protein disorder predictions we identified two IDRs at DDX18’s N-terminal and C-terminal ends, respectively (Fig. S2a). Phase separation properties of DDX18 under or near physiological salt concentration (150 mM NaCl) were first verified in vitro using DDX18-GFP recombinant protein (Fig. 2c, conditions A’, B’, and C’), which does not require the molecular crowding reagent polyethylene glycol (PEG). DDX18 droplets can be disrupted by higher salt concentrations (Fig. 2c, d, condition series from D’ to F’); however, this disruption could be overcome by increasing the protein concentration (Fig. 2d). The LLPS nature of DDX18 in vivo was also confirmed in hESCs expressing DDX18-GFP at the endogenous DDX18 locus (Fig. 2e) by time-lapse microscopy, revealing that DDX18-GFP undergoes fusion and fission within minutes (Fig. 2f) comparable with the nucleolus fusion speed in vivo^11,38, and by fluorescence recovery after photobleaching (FRAP) with the average time for DDX18 to fully recover after the photobleaching around 104 ± 8 s (Figs. 2g, S2b), slightly slower than GC protein NPM1³⁸.

DEAD-box helicases are conserved from bacteria to mammals containing the N-terminal domain, helicase core domain, and C-terminal domain⁴¹. The interactions implicated in the phase separation include long-range electrostatic effects, prion-like property-mediated interactions, and aromatic amino acid-mediated π-π interactions⁴². To explore which domains and which types of molecular interactions confer DDX18 functions in phase separation, we analyzed the fraction of charged residues and net charge per residues (NCPR) (Fig. S2c, top track), presence of prion-like domain (PLD) (Fig. S2c, middle track), and propensity for π-π contacts (Fig. S2c, bottom track). We found neither PLD nor π-π contacts were involved in DDX18 phase separation. Instead, NCPR analysis identified positively charged basic tracts (blue) highly enriched within NIDR and CIDR of DDX18 (Fig. S2c). Furthermore, we found that both NIDR and CIDR are required for DDX18 phase separation: lacking either domain (ΔNIDR or ΔCIDR) or both (ΔIDR) greatly reduced the droplet formation in vitro (Fig. 2h, i) and in vivo (Figs. 2h, S2d). When fused with a far-red fluorescent protein (faRFP), NIDR alone, but not CIDR alone, was enough to drive the fusion protein into the nucleolus in vivo (Fig. S2d). Interestingly, we found NIDR harbors a predominant positively charged basic tract consisting of an evolutionarily conserved K-stretch (8 Lysines and 1 Arginine) (Fig. S2e, f), previously reported to be enriched in disordered regions and interact with RNA to drive and regulate cellular condensations⁴³. Accordingly, we found that the K stretch mutant (9 K/R to all A mutation; 9 A) was compromised in its nucleolar localization in vivo (Fig. S2g) and RNA-facilitated formation of the “core-shell” like structure⁴⁴ compared with WT (Fig. S2h). Of note, the DQAD helicase mutant of DDX18 failed to form any core-shell-like structure (Fig. S2h).

To further corroborate the above findings, we conducted a rescue experiment using wild-type (WT), DQAD, and IDR-deleted versions of DDX18 in DDX18 KD cells, followed by qPCR analysis. Our data demonstrated that only the wild-type DDX18 could fully restore hESC maintenance (Fig. S2i) and pluripotency/lineage gene expression programs (Fig. S2j) during differentiation, thus confirming the essential role of DDX18’s RNA helicase activity and phase separation property in its functionality. Together, these results establish the LLPS nature of DDX18 that is promoted by its RNA helicase activity and the RNA-interacting NIDR, suggesting that DDX18 may contribute to the LLPS of nucleoli in regulating RiBi and protein synthesis for pluripotency of hESCs.

DDX18 binds nucleolar localized RBPs and rRNAs/snoRNAs

To investigate how DDX18 exerts such nucleolar functions through its phase separation properties, we identified DDX18-interacting proteins and RNA targets in hESCs. First, we established a transgenic hESC line expressing 3xFlag-DDX18 for affinity purification of DDX18 and its interacting partners in the hESC nuclear extracts using anti-Flag beads (Fig. S3a). Then, with a stringent cut-off (number of unique peptides ≥4 and ratio of 3xFlag-DDX18/IgG ≥ 4), we identified 140 DDX18-interacting proteins (Supplementary Data 3). GO analysis of these proteins linked them to rRNA processing, translational initiation, and RNA secondary structure unwinding (Fig. 3a). As an important hub in maintaining genome stability, the nucleolus is highly enriched for DNA repair proteins, some of which were identified as DDX18-binding proteins with validated nucleolar localization and functional involvement in RiBi, including PARP1⁴⁵, SMC3⁴⁶, and TRIM28⁴⁷. However, we cannot exclude the possibility that DDX18 may also play a role in DNA damage repair, a topic worthy of future investigation.

**Fig. 3: DDX18 interacts with nucleolar RNAs and GC component proteins.**

Notably, the two well-known nucleolar GC marker proteins, NCL and NPM1, were also detected in the DDX18 interactome. We further confirmed the DDX18-NPM1 interaction by co-immunoprecipitation (co-IP) (Fig. 3b) and bimolecular fluorescence complementation (BiFC)⁴⁸ (Figs. 3c, S3b), in line with their GC co-localization (Fig. 2b) and supporting a GC-specific function of DDX18. Importantly, the DDX18-NPM1 interaction was attenuated by RNase A treatment in co-IP (Fig. 3b) and BiFC (Fig. 3c) assays, supporting an RNA-facilitated interaction between DDX18 and NPM1.

To identify the RNA targets of DDX18 in hESCs, we performed iCLIP (individual-nucleotide-resolution crosslinking and immunoprecipitation) with the established 3xFlag-DDX18 hESC line (Fig. S3c). DDX18 binds to a diverse set of RNAs, among which rRNAs were most highly represented (Fig. 3d; Supplementary Data 4), while some snoRNAs were also bound by DDX18, consistent with its nucleolus-specific localization (Fig. 2a, b). Upon evaluating our results in relation to iCLIP-seq data for ADAR1⁴⁹, a well-studied nuclear-localized RBP in hESCs, we noted an elevated degree of rRNA and snoRNA binding to DDX18 over ADAR1 (compare Fig. 3d with Fig. S3d). Interestingly, the C/D box snoRNA motif RUGAUGA was ranked among the top three in motif analysis (Fig. 3e), and DDX18 binding to C/D box snoRNAs (SNORDs) was confirmed by CLIP-qPCR, exemplified by a few SNORD members (Figs. S3e, S3f, top panel). We also noticed that DDX18 could bind protein-coding mRNAs (Figs. 3d, S3f, bottom panel; Supplementary Data 5). Of note, mRNA of genes residing in DNA nucleolus-associated domains (NADs) are highly enriched in the nucleolus⁵⁰, and we found that the DDX18-interacting mRNAs were also enriched for GO terms in RiBi and translation (Fig. 3f). Furthermore, unlike the DFC component FBL⁵¹, DDX18 does not bind to 1∼414 nt (5′ ETS-1) region of 47S pre-rRNAs localized in DFC (Fig. S3g), further supporting the nucleolar GC-specific functions of DDX18 in hESCs. These data establish physical and functional interactions between DDX18 and nucleolar RBPs and RNA species that underlie their roles in maintaining the pluripotency of hESCs.

DDX18 maintains nucleolar structural integrity through multiphase liquid miscibility and immiscibility with GC and DFC proteins, respectively

We next investigated how DDX18 may contribute to the structural integrity of the nucleolus, particularly in the context of DFC and GC. GC phase is known for enriching negatively charged proteins such as NPM1 and NCL, and phase separation of NPM1 is enhanced by positively charged peptides^52,53. As DDX18 binds to NPM1 both in vitro and in vivo (Figs. 3a–c, S3a, b), we compared the biophysical properties of DDX18 and NPM1 by testing their droplet miscibility. We found that NPM1-RFP cannot form droplets in vitro without the PEG crowding reagent (Fig. 4a), as reported⁵⁴. Interestingly, adding NPM1-RFP solution without droplets to the preformed DDX18 droplets, we observed a profound “core-shell” like structure containing homogeneous DDX18-GFP and NPM1-RFP mixture in the same compartment, phase-separated from the empty core structures, mimicking the three-layered structure of the nucleolus (Fig. 4b). The hollow-like condensates also underwent shell fusion and cavity fusion (see “Supplementary Movie”). In contrast, neither homogenous droplets nor “core-shell” structures were observed upon mixing preformed DDX18-GFP and RFP-FBL droplets together (Fig. S4a, b), verifying the immiscibility of DDX18 and FBL droplets. The observed differential miscibility of DDX18 with NPM1 and FBL droplets was consistent with NPM1 and FBL being the most representative components of the phase-separated GC and DFC regions within the nucleolus, respectively³⁸. To further probe the potential contribution of DDX18 to nucleolus architecture through its multiphase liquid (im)miscibility, we simultaneously mixed DDX18-GFP, NPM1-CFP, and RFP-FBL recombinant proteins. We observed a co-existing multiphase structure mimicking the regular nucleolus structure (Fig. 4c, top panel). As RNA helicase activity is critical for DDX18 functions (Figs. 1i, S2h–j), we tested how DDX18-bound RNAs (rRNA and snoRNA) may contribute to forming this particular multiphase structure. Intriguingly, the introduction of snoRNA or rRNA leads to a significant increase in the proportion of structures resembling the nucleolus (Fig. 4c, middle and bottom panels). This observation suggests a dynamic interplay between these RNA components and the formation of nucleolar-like structures.

**Fig. 4: Architecture of the nucleolus arises through multiphase liquid miscibility and immiscibility of DDX18 with NPM1 and FBL, respectively.**

To further understand its nucleolar functions in maintaining hESC pluripotency in vivo, we addressed how DDX18 depletion may modulate nucleolar architecture by immunostaining and 3D-SIM imaging in control and DDX18 KD hESCs. In hESCs with control KD (shLuc), the DFC marker FBL is completely encompassed (or included) in the NPM1-marked GC as immiscible multiphase droplets (Fig. 4d, top). In contrast, upon DDX18 depletion (shDDX18), the DFC region starts “leaking/escaping” (or excluded) from the GC and forming “nucleolar cap” or “nucleolar segregation” like structures (Fig. 4d, bottom) without significant changes of the core nucleolar protein levels including the GC markers NPM1 and NCL, and the DFC marker FBL (Fig. S4c, d). Quantification of the representative images on such “inclusive/exclusive” structures indicated that more than 40% of the DDX18 KD hESCs, compared to less than 10% of the control KD hESCs, showed mislocalization of FBL (Fig. 4e). Importantly, mislocalization of FBL, coincident with nucleolar enlargement, was also observed more prominently in the mesendoderm lineage than in the ectoderm lineage upon the directed trilineage differentiation of hESCs (Fig. S4e, f).

Our data thus establish a critical role of DDX18 in maintaining hESCs by preserving the structural integrity of the nucleolus through its multiphase liquid miscibility and immiscibility with the key GC and DFC proteins NPM1 and FBL, respectively (Fig. 4f, left). DDX18 depletion in hESCs, under the KD or differentiation condition, results in the mislocalization of DFC as “nucleolar caps” protruding from the GC (Fig. 4f, right) and lineage-specific nucleolar enlargement (Fig. S4e), causing the defects in RiBi and translation and, consequently, the loss of hESC pluripotency with the skewed differentiation towards mesendoderm lineage.

DDX18 restricts perinucleolar heterochromatin organization via its liquid miscibility and immiscibility with NPM1 and HP1 droplets, respectively

Nucleolar structure alteration is reported to cause chromatin reorganization^55,56,57. During ESC differentiation, the initially dispersed H3K9me3-marked and HP1-containing heterochromatin foci (i.e., PNH) start accumulating around nucleoli^5,20,58,59. Tethering heterochromatin at nucleoli promotes transcriptional activation of differentiation genes, leading to the exit of pluripotency in mESCs²⁰. We thus asked whether the PNH formation could be facilitated by DDX18 KD, leading to differentiation of hESCs as we observed (Fig. 1). Indeed, we found that DDX18 depletion led to the redistribution of dispersed HP1α/β foci, characteristic features of heterochromatin, from nucleoplasm in control cells (shLuc) closer to nucleolus in DDX18 KD cells (shDDX18) (Fig. 5a, b), colocalizing with heterochromatin-associated H3K9me3 (Fig. S5a), indicative of the PNH accumulation. These findings establish the requirement of nucleolus structural integrity for restricting PNH formation and maintaining the pluripotent dispersed heterochromatin state.

**Fig. 5: DDX18 KD disrupts nucleolar structure leading to perinucleolar heterochromatin organization.**

To explore the mechanisms by which DDX18 restricts PNH formation, we first sought to analyze the (im)miscibility of the droplets formed by GC phase proteins and the heterochromatin protein HP1α, given that HP1α phase separation regulates the liquid droplet-like properties of heterochromatin^60,61,62,63. By mixing RFP-HP1α with DDX18-GFP and NPM1-CFP, respectively, we observed that HP1α droplets were miscible with both DDX18 and NPM1 in droplet formation (Fig. 5c). NPM1 contains two evolutionarily conserved HP1α binding motifs (PXXVXL and PXVXL) (Fig. S5b) with reported roles in regulating HP1α oligomerization and promoting the formation of phase-separated droplets of HP1α⁶⁰, explaining the miscibility between HP1α and NPM1 droplets. The fact that PNH is not accumulated in undifferentiated hESCs despite normal expression levels of heterochromatin HP1 proteins suggests additional factors may act upon HP1α and NPM1 to restrict their liquid miscibility. We thus tested DDX18 action in this context by simultaneously mixing DDX18-GFP, NPM1-CFP, and RFP-HP1α recombinant proteins to explore their multiphase organization behaviors. We found that NPM1 preserves the homogeneous droplet formation with DDX18 (merged image of the cyan droplets in Fig. 5d), consistent with our prior in vitro observation (Fig. 4c). However, NPM1 suddenly lost its liquid miscibility with HP1α when DDX18 was present with HP1α being pushed outside of the DDX18/NPM1 condensates, resulting in the formation of a shell-like structure (Fig. 5d, e). These data indicate that, while maintaining its liquid miscibility with NPM1 droplets for GC integrity, DDX18 prevents the HP1α-NPM1 droplet miscibility, hindering the PNH accumulation at GC (see Discussion).

Next, we addressed why and how the heterochromatin prefers to dock/accumulate at the nucleolus region during ESC differentiation after DDX18 depletion. As multiphase organization could be regulated by interconnected complexes (nodes) of multivalent proteins^63,64, we sought to find the connecting node between PNH and the nucleolus. In mESCs, it has been reported that transcriptionally repressive regions near centromeres are often closer to the nucleolus²¹. Within the human genome, 45S rDNA tandem repeats are localized as rDNA gene arrays at the short arms of five chromosomes (Chr13, 14, 15, 21, and 22) flanked by the centromere and telomere (Fig. S5c). We detected centromere positioning by staining with the centromere markers CENPA and CENPT and observed a significant increase of centromere clustering around the nucleolus upon DDX18 depletion (Figs. 5f, g, S5d, e). Genome-wide occupancy of DDX18 by ChIP-seq revealed that DDX18 mainly binds to the rDNA transcribed regions (Fig. S5f), which is consistent with our previous study in mESCs¹⁹. In contrast, it is well established that the heterochromatin proteins HP1α/β bind to centromeres/pericentromeres⁶⁵ and that the GC protein NPM1 binds to the centromere^66,67,68,69. Thus, we hypothesized that centromere and centromere-binding proteins from the nucleolus might provide the connecting node for PNH to dock around the nucleolus. Indeed, we found that DDX18 depletion resulted in increased centromere binding of NPM1 within hESCs (Fig. 5h).

Together, these data support a model whereby DDX18 coordinates with NPM1 to safeguard nucleolar GC integrity and restrict centromere clustering and PNH formation (Fig. 5i, left). When DDX18 is absent or depleted, leading to the nucleolar cap formation (Fig. 4d, e), NPM1 droplets become miscible with HP1 droplets with increased centromere binding, promoting heterochromatinization (Fig. 5i, right), an early event required for ESC differentiation²⁰.

DDX18 depletion results in nuclear reorganization

To address how chromatin may be reorganized at a genome-wide scale contributing to changes in gene expression and, consequently, the differentiation of DDX18 KD hESCs, we applied the Hi-C seq technology to generate chromatin interaction data in control and DDX18 KD cells. Hi-C data allow the partition of the genome into two compartments called A/B compartments, containing relatively active (A) and inactive (B) regions, respectively⁶⁴. The saddle plots (Fig. S6a) revealed that DDX18 depletion leads to moderate changes in A/A (an increase from 1.369 to 1.419) and B/B (a decrease from 1.346 to 1.307) homotypic interactions, indicating a decrease in chromatin interaction within heterochromatin. In contrast, more minor changes in A/B or B/A heterotypic interaction were observed (Fig. S6a). Further using principal component analysis (PCA) at 100-kb resolution to classify the genome into A and B compartments, we identified regions that changed compartments upon DDX18 KD. We found 3.2% of regions switched from A to B and 3.3% from B to A (Fig. S6b). However, integrating these results with RNA-seq data showed that over 90% of significantly up- or down-regulated genes were not in compartment-switching regions (Fig. S6c). We then analyzed gene expression changes in 100-kb bins based on their PC1 value changes, indicating shifts towards active or inactive states. We found that a small proportion of bins with extreme PC1 changes correlated with significant gene regulation, with only 195 and 17 genes (out of >2000 differentially expressed) in bins showing PC1 changes greater than −2 and +2, respectively (Fig. S6d). Further analysis of upregulated lineage-associated genes showed most were in regions with moderate PC1 changes (−0.5 to +0.5) (Fig. S6e). This weak correlation between PC1 changes and transcription was also observed globally (Fig. S6f). Together, our results suggest that, while locus-specific transcriptional changes resulting from the compartment switch do occur (Fig. S6g), a vast majority of transcriptional changes observed upon DDX18 KD arise from mechanisms independent of compartment organization. This observation is consistent with the fact that DDX18 is an RBP rather than a typical transcription factor or chromatin remodeler and that DDX18 mainly binds to the rDNA transcribed regions in the nucleolus (Fig. S5f). Hence, it is more likely to influence gene expression via its unique nucleolar functions.

A/B compartments are primarily defined by intrachromosomal interactions within each chromosome⁷⁰. By analyzing the intrachromosomal interaction for all five rDNA-containing chromosomes (Chr13, 14, 15, 21, and 22), we only found two interacting regions (R1 and R2) of Chr21 showing significantly increased contact frequencies after DDX18 KD (Figs. 6a, S6h). Both interacting regions are close to the pericentromere. R1 is mediated by rDNA closer locus (Site I, Chr21: 9.4 MB-10.1 MB) and chromosome arm locus (Site III, Chr21: 24.9 MB-26.5 MB), while R2 is mediated by pericentromeric locus (Site II, Chr21: 14.7 MB-15.1 MB) and chromosome arm locus (Site III, Chr21: 24.9 MB-26.5 MB) (Fig. 6b). Interestingly, we found that Site III is localized in the lamina-associated domain (LAD)⁷¹ (Fig. S6i). The increased interactions of R1 and R2 in DDX18 KD relative to control KD samples suggest that DDX18 depletion could affect nucleolus-associated domain (NAD) and LAD exchange (Fig. 6c). Together with our findings that DDX18 depletion impairs nucleolus integrity (Fig. 4) and PNH organization (Fig. 5), we concluded that DDX18 could play a pivotal role in regulating centromere-telomere proximal rDNA-containing chromosome (i.e., Chr21) organization, i.e., heterochromatin formation.

**Fig. 6: DDX18 depletion results in chromatin reorganization.**

Since the nucleolus is a hub for interchromosomal interactions, we next examined the contacts between ribosomal DNAs (rDNAs) from nucleolar organizer region (NOR)-bearing chromosomes (Chr13, 14, 15, 21, and 22). The analysis of chromatin interactions in the short arms of NOR-bearing acrocentric chromosomes is challenging due to their repetitive nature and gaps in the human reference genome GRCh38. Utilizing the mHi-C pipeline⁷², which retains multimapping reads, enabled us to analyze rDNA interactions, revealing improved detection in the repetitive short arms of acrocentric chromosomes 15, 21, and 22 (Fig. S6j). However, gaps in the GRCh38 reference genome (Fig. S6k top) still limited the analysis. Remapping Hi-C data to the updated T2T-CHM13 reference genome⁷³ allowed us to fill many gaps, enhancing the resolution of contact maps in acrocentric chromosomes (Fig. S6k bottom). Focusing on the impact of DDX18 KD, we detected significant changes in chromatin organization, with 31% of interactions shared between control KD and DDX18 KD conditions (Fig. S6l). Next, to specifically study the impact of DDX18 KD on rDNA-containing chromosomal interactions, we initially calculated the change in interaction strength (observed/expected counts) upon DDX18 KD of all shared interactions across the genome and categorized our results by chromosome arm. Interestingly, this analysis showed a preferential weakening of Hi-C interactions, specifically in the short arms of acrocentric chromosomes containing rDNA repeats. In contrast, long arms and non-acrocentric chromosomes showed no significant change (Fig. 6d). Importantly, an increase in interchromosomal contacts among acrocentric chromosomes was observed (Fig. 6e and Fig. 6c model), suggesting that DDX18 plays a crucial role in organizing NORs.

Nuclear reorganization upon DDX18 depletion results in developmental gene expression changes

To understand how nuclear reorganization involving LAD-NAD exchange and NORs may lead to gene expression changes in DDX18-depleted hESCs, we studied the impact of DDX18 KD on the HOXA gene cluster for the following reasons. First, the HOXA gene cluster is in a gene-poor region of Chr7 that, in human cancer cells, is sequestered by the Nup93 sub-complex for HOXA repression at the nuclear periphery (LAD). The derepression of HOXA genes upon Nup93 depletion results in the LAD to nuclear interior (closer to NAD) relocation^74,75. Second, the HOXA gene cluster represents the CTCF-mediated chromatin loops and chromatin status changes during embryonic development and disease progression^76,77,78. Intriguingly, a published study reported the interchromosomal interaction between the NOR-bearing Chr21 and HOXA gene-residing Chr7 and its resultant t(7;21) translocation in a human leukemia patient⁷⁹. Third, the HOXA cluster genes were upregulated in our DDX18 KD hESCs (Fig. 1h) coincident with weakened interchromosomal interactions of HOXA-residing Chr7 with Sites II and III in Chr21 (Fig. S6m), suggesting that the DDX18 KD effect on the 3D conformation of those NOR-bearing chromosomes (e.g., Chr21) could indirectly affect chromatin organization (e.g., Chr7) and developmental gene expression (e.g., HOXA genes) (see Fig. 6c model).

To substantiate our model whereby DDX18 depletion could cause the NAD and LAD exchange and HOXA gene derepression (Fig. 6c), we performed the following studies. First, we performed additional Hi-C data analysis on chromatin interactions involving the HOXA locus and revealed that, despite the lack of a major compartment switch in the DDX18 KD hESCs (Fig. S6a–c), loops involving the HOXA locus within Chr7 appeared strengthened in the DDX18 KD hESCs (Fig. 6f). In addition, DDX18 KD hESCs showed reduced interaction frequencies within the HOXA gene cluster (Fig. S6n, o, left panels) and increased the external interactions between the HOXA cluster and the nearby genes (e.g., HOTTIP) (Fig. S6n, o, right panels). Furthermore, DNA FISH analysis revealed that HOXA9-13 loci are relocated away from the nucleolus (NAD) (i.e., reduced FISH signals) towards the nuclear periphery (LAD) (i.e., increased FISH signals) in DDX18 KD relative to control KD hESCs (Fig. 6g, h). Importantly, RNA FISH analysis also detected the corresponding upregulation of HOXA transcripts, exemplified by increased RNA FISH signals per nucleus of HOXA9 (Fig. 6i, k) and HOXA11 (Fig. 6j, l) at the nuclear periphery (LAD), while their NAD signals were slightly and appreciably reduced. These findings demonstrate that DDX18 depletion modulates the heterochromatin via the LAD and NAD exchanges associated with the NOR-bearing chromosomes, resulting in nuclear reorganization and derepression of developmental genes (e.g., Chr7 reorganization at the HOXA locus; Fig. 6c model).

Nucleolus-specific NoCasDrop restructures the genome and controls gene expression

We have thus far established the roles of DDX18 in maintaining the intrinsic nucleolar architecture (Fig. 4f) and extrinsic heterochromatin organization (Fig. 5i) for pluripotency and lineage commitment/differentiation of hESCs. However, it remains an open question whether the extrinsic mechanism, i.e., centromere clustering and PNH accumulation, is the cause or consequence of hESC differentiation. Therefore, inspired by CasDrop, a recently developed method successfully applied in mammalian cells to activate gene expression⁸⁰, we created NoCasDrop (Nucleolus specific CasDrop) to address whether tethering heterochromatin towards nucleolus would impair hESC pluripotency.

To this end, we took advantage of the nucleolus-specific phase separation by fusing full-length (FL) DDX18 and its individual domains (NIDR, HD, and CIDR) with the dCas9-mCherry (Figs. 7a, S7a). We found that both FL and NIDR of DDX18 fusion, but not the other domains, could recruit dCas9-mCherry protein to the nucleolus (Fig. S7a). However, FL, but not NIDR, could activate the pre-rRNA transcription (Fig. S7b). Therefore, to avoid any indirect effects resulting from rRNA overexpression, we used NIDR NoCasDrop for our further study (Fig. 7a). To overcome the challenge of manipulating a whole trunk of chromosome fragment by a single dCas9, we targeted tandem repeats in centromere and satellite DNA by infecting the NIDR NoCasDrop stable cell line with a human centromere-specific alpha-satellite sgRNA (sgAlpha) and a non-specific control sgRNA (sgNS), both of which have been functionally validated in human cell lines previously⁸¹. This allows multiple copies of dCas9 to coordinately target the same gene locus by the same sgRNA leading to a combined/concerted action in controlling the chromosome localization (Fig. 7a). Then, we performed immunostaining after 48 h of sgRNA infection and found that the centromere (Fig. 7b, c) and heterochromatin maker HP1α (Fig. 7d, e) could be tethered to the perinucleolar region by NoCasDrop and α-satellite specific sgRNA. More importantly, we found the derepression of developmental genes (e.g., HOXA cluster genes) by this NoCasDrop-mediated perinucleolar centromere clustering (Fig. 7f). In contrast, pluripotency genes and RiBi genes were minimally affected within the tested time window (Fig. S7c), indicating the specificity of NoCasDrop and corroborating the Hi-C data. These results also suggest that the derepression of HOXA cluster genes is an early event leading to the loss of pluripotency in DDX18 KD hESCs.

**Fig. 7: NoCasDrop of DDX18 restructures the genome positioning and controls gene expression.**

These data reinforce a major function of DDX18 in restricting centromere clustering around the nucleolus for the repression of developmental genes (e.g., HOXA cluster genes) in undifferentiated hESCs and provide a molecular tool to tether centromere clustering at the perinucleolar region for cell fate manipulation.

Discussion

Accumulating evidence indicates that the nucleolus functions as the ribosome biogenesis (RiBi) site for protein synthesis and plays a role in chromatin organization and transcriptional regulation in mESCs^47,82. Using a human pluripotent stem cell paradigm, we demonstrate that, as a nucleolus-specific RNA helicase, DDX18 controls RiBi and global translation while simultaneously preventing PNH formation through the modulation of nucleolus phase separation in maintaining human pluripotency. Specifically, under the pluripotent state, DDX18 maintains the nucleolar structural integrity and prevents PNH formation through its liquid miscibility with the core GC factor NPM1 and its immiscibility with the core DFC factor FBL (Fig. 7g). In contrast, when DDX18 is depleted (e.g., through genetic manipulation) or downregulated (e.g., during development or hESC differentiation), NPM1 increases its binding to the centromere and acquires the liquid miscibility with HP1, resulting in the escape of DFC from the center of the nucleolus and the formation of “nucleolar caps” with accumulated PNH (Fig. 7h). While specific nucleolar caps were reported to be located adjacent to the distal junction sequences nearby the short arm of the acrocentric chromosome anchored in PNH⁸³, it remains to be determined whether the “nucleolar caps” from those escaped DFCs in DDX18 KD hESCs (Fig. 4f) are the source of or overlapped with PNH. Notably, the mesendoderm lineage-specific increase of FBL mislocalization and enlargement of nucleoli in directed trilineage differentiation of hESCs (Fig. S4e, f) are reminiscent of enhanced nucleolar caps (Fig. 4d, e) and skewed mesendoderm differentiation (Figs. 1 and S1) of DDX18 KD cells. In addition, surface tension is crucial in biological processes, affecting how liquid droplets behave within cells. The phase separation of the initially miscible NPM1 and HP1 droplets in the presence of DDX18 (compare Fig. 5d with Fig. 5c) suggests that DDX18 may prevent the NPM1-HP1 droplet formation in pluripotent cells by its stronger interaction/miscibility with NPM1 than with HP1, evidently supported by DDX18-NPM1 droplets are being more homogeneous (Fig. 4a–c) than DDX18-HP1 droplets (Fig. 5c top; note the merged unpure yellow droplets with tainted red and green dots). Thus, our study uncovers a previously unappreciated role of the nucleolar RNA helicase DDX18 in coordinating nucleolus phase separation and chromatin organization to control the pluripotency of hESCs.

We previously reported that Ddx18 maintains mESC pluripotency by antagonizing PRC2 to protect rDNA from epigenetic silencing, ensuring high RiBi in mESCs¹⁹. However, this epigenetic mechanism is likely not active in hESCs. First, Ezh2/EZH2 depletion affects mESC differentiation but causes severe self-renewal and growth defects in hESCs^84,85. Second, the differentiation phenotype of DDX18 loss can be partly rescued by PRC2 depletion in mouse¹⁹ but not human (Fig. S7d) ESCs. Third, the DDX18-PRC2 partnership is detectable in mouse¹⁹ but not human (Fig. 3a; Supplementary Data 3) ESCs. Fourth, mouse and human ESCs represent distinct pluripotency states at functional^86,87 and molecular⁸⁸ levels, indicating that human pluripotency cannot be inferred from mouse studies. Our findings provide molecular insights into nucleoli’s role in chromatin organization, beyond being a RiBi compartment, in controlling stem cell pluripotency and lineage commitment.

The nucleolar GC protein NPM1 maintains mESCs, with its loss resulting in differentiation⁸⁹. NPM1 forms liquid-like droplets in the presence of RNA⁵² and is required for centromere clustering and anchoring to the nucleolus⁹⁰. An outstanding question is how such an essential and ubiquitous nucleolar function can meet the specific needs of different cell types and/or during various developmental stages. DEAD-box (DDX) or DEAH-box (DHX) RNA helicase proteins are evolutionarily conserved with cell/tissue-specific expression profiles and prominent roles in LLPS²³. In contrast to DDX18, which maintains the undifferentiated state in hESCs, DHX9 is dispensable for ESC maintenance but required for differentiation, processing IGS-rRNA into mature rDNA promoter RNA (pRNA)⁹¹ to promote TIP5 interaction and rRNA gene silencing, leading to low RiBi in differentiated cells⁹². This pRNA maturation initiates heterochromatin formation in the nucleolus, maintaining some undifferentiated ESC features while promoting the exit from pluripotency²⁰. We demonstrate that DDX18 binds to both rRNAs and snoRNAs, the latter assisting the 2’-O-methylation of the former^93,94. While the interaction of rRNA⁵² or nucleolar localized lncRNA^95,96 with nucleolar proteins was reported to contribute to their phase separation, our study also implicates snoRNAs and likely rRNA modifications in DDX18’s coordination of nucleolar multiphase separation. Furthermore, the requirement of the RNA helicase activity of DDX18 to rescue the DDX18 KD defects also suggests the involvement of rRNA/snoRNAs secondary structures in nucleolar LLPS properties. Future investigations are needed to unravel these details further.

It is noteworthy that HOXA clusters can change their nuclear location under physiological or leukemic conditions^97,98. Intriguingly, supporting the inter-chromosomal interaction between the NOR-bearing Chr21 and HOXA gene-residing Chr7 in hESCs (Figs. S6m, 6c), the interaction between Chr21-Chr7 and its resultant t(7;21) translocation were reported in a case of acute myeloid leukemia⁷⁹. In addition, while pluripotent cells were known to be relatively depleted for H3K9me3-marked constitutive heterochromatin^99,100, differentiation of hESCs is indeed associated with H3K9me3 accumulation at the HOXA loci for their activation, as reported³⁰, a finding that is itself counterintuitive but nonetheless supports our hypothesis. More importantly, published studies have established that HOXA genes, especially HOXA9, play an important role in the differentiation of hESCs towards mesodermal and hematopoietic fates¹⁰¹. In addition, mouse HoxA genes were also reported to play an essential role in specifying mesendoderm lineage during mESC differentiation and embryonic development¹⁰². Overall, HOXA genes’ upregulation appears to be an important step in the exit from pluripotency and commitment to the lineages derived from mesendoderm (reviewed by^31,32).

Our findings contribute to an increased understanding of the nucleolus in the nuclear (i.e., genome/chromatin) organization, which is currently under-investigated compared with the well-recognized role of the nuclear periphery in genome organization. Heterochromatic domains are always associated with the nuclear lamina (LADs) and nucleoli (NADs). Therefore, manipulating the chromatin organization within the nucleus to control gene transcription is desirable for studying cell biology. However, the membrane-less compartment and LLPS nature of the nucleolus²⁴ render it technically more difficult to identify NADs than LADs^71,103, a major challenge in studying NADs. Here, we developed a nucleolus-specific NoCasDrop technique to dissect how phase separation might target genomic domains in the nucleolus. By using NoCasDrop, we successfully tethered the centromere to the nucleolus and reprogrammed the developmental gene expression, providing direct support for DDX18 functions in controlling human pluripotency by maintaining nucleolus phase separation and restricting centromere clustering around nucleolus in hESCs. The molecular features of our NoCasDrop-treated hESCs are analogous to those of mESCs with nucleolar pRNA-tethered heterochromatin at rDNA loci, initiating structural remodeling toward a highly condensed nuclear heterochromatin, resulting in the upregulation of genes involved in cell differentiation while maintaining the undifferentiated state with normal ESC proliferation, expression of pluripotency genes, morphology, and AP staining²⁰. Our findings and the NoCasDrop tool should be applicable for further studies of nucleolar functions in development and disease and for cell fate manipulation.

Methods

Cell lines

H9 and HES2 hESCs were initially obtained from WiCell Research Institute and maintained in the Wang lab.

DDX18 C-terminal GFP knock-in cell line (H9-DDX18^GFP) was established by the CRISPR-Cas9 method and maintained in the Wang lab. First, H9 hESCs were transfected with a plasmid expressing Cas9/sgRNA (5’-GCAAGAAATCATCTGACAGC-3’) in the presence of a recombination GFP donor DNA flanked by right and left homology arms (HA) of DDX18. Then, three days after transfection, GFP-positive cells were selected by FACS, expanded, and validated by immunoblotting.

The 3xFlag-DDX18 transgenic cell line was established by PiggyBac transposon-mediated reversible genetic modification strategy and maintained in Wang lab. We co-transfected PB-3xFlag-DDX18 with PiggyBac transposase (PBase) to create the stable hESC lines (H9 and HES2) with the selection for hygromycin resistance. DDX18 gene and protein expression levels were confirmed by qPCR and western blotting analyses.

Plasmid construction

shRNA cloning

shRNAs were designed using the RNAi Consortium (TRC) siRNA selection program from MIT Whitehead Institute (http://sirna.wi.mit.edu/). pLKO.1-GFP is a replication-incompetent lentiviral vector chosen by the TRC for the expression of shRNAs. Oligos were first annealed and then cloned into pLKO.1-GFP vector following the protocol described by Addgene (www.addgene.org). Target sequences: DDX18 shRNA1 (shDDX18#1), CCAAGGTTCCATTAAGTGAAT; DDX18 shRNA3 (shDDX18#3), CGTGCATACCTATGGCTTGAT; shRNA targeting the firefly luciferase was used as a control (shLuc).

cDNA cloning

Human DDX18 cDNA was purchased from OriGene and was amplified by PCR using primers designed with 5’ NheI and 3’ PacI restriction sites (Supplementary Data 6) and cloned into the PiggyBac (PB) vector with 3xFlag tag in it.

NoCasDrop plasmid construction

Full-length (FL) DDX18 and its domains (NIDR, HD, and CIDR) were amplified by PCR with the primers containing 5’ NotI and 3’ XhoI restriction sites (Supplementary Data 6). PCR products and the dCas9-mCherry vector⁷⁰ were digested with restriction enzymes, purified, and ligated with T4 DNA ligase. All plasmids were confirmed by Sanger sequencing.

Construction of plasmids for recombinant proteins

Full-length human NPM1-RFP, NPM1-CFP, RFP-FBL, and RFP-HP1α were PCR amplified by G5 High Fidelity DNA Polymerases (EnzyArtisan, China) and then sub-cloned into a modified pET28b vector with a SUMO protein fused at the N terminus after the 6xHis tag by One-Tube Cloning Kit (EnzyArtisan, China). In addition, full-length human DDX18 and its truncations or mutants were PCR amplified and subcloned into a modified pET vector with an 8xHis tag followed by a GFP protein fused at the C-terminus. The plasmids were then extracted using the SPARKeasy Superpure Mini Plasmid Kit (Sparkjade Science Co., Ltd., China), and all inserted sequences of plasmids were validated by Sanger sequencing.

Human ESC culture

All hESC lines were cultured at 37 °C in a controlled humidified atmosphere of 5% CO_2, as described previously⁸⁸. The hESC lines used were H9 and HES2 (WiCell), authenticated by the provider, and routinely tested negative for mycoplasma. hESCs were maintained on irradiated inactivated MEFs in hESC medium composed of DMEM/F12 (Corning) containing 20% Knockout serum (Invitrogen), penicillin/streptomycin, L-glutamine, non-essential amino acids, β-mercaptoethanol (β-ME, Sigma), and bFGF (R&D Systems). For feeder-free culture, hESCs were cultured in mTeSR Plus (STEMCELL Technologies) on Matrigel hESC-Qualified Matrix (Corning) coated wells and passaged using ReLeSR (STEMCELL Technologies) every 3 to 4 days.

EB differentiation

For non-lineage-specific differentiation, we used the previously described EB formation protocol¹⁰⁴. H9 hESCs were cultured in a feeder-free mTeSR medium. The cells were treated with collagenase B (Roche) for 15–30 min and collected by scraping. After centrifuging, cell pellets were resuspended in differentiation media, StemPro 34 (Invitrogen) supplemented with 2 mM L-Gln, 4 × 10⁻⁴M monothioglycerol (MTG), 50 μg/ml ascorbic acid, and 1 mM transferrin (Sigma). EBs were grown in ultra-low adherent plates (Costar), and the medium was changed every three days. After eight days of differentiation, EBs were collected, resuspended in DMEM 10%, and transferred to gelatin-coated dishes to allow them to attach and differentiate for 4–7 additional days. Whole-cell lysates were collected at specific time points during the differentiation.

Direct differentiation of human embryonic stem cells into trilineage

Human ESCs (H9) were differentiated into three germ layers using STEMdiff™ Trilineage Differentiation Kit (STEMCELL Technologies, Cat#05230). HESCs were cultured in mTeSR™1 medium and differentiated following the trilineage differentiation protocol. Cell differentiation markers for each lineage were detected by immunostaining. PAX6 antibody (Santa Cruz, sc-81649), Brachyury antibody (R&D, AF2085), and CXCR-4 antibody (Santa Cruz, sc-53534) were used for ectoderm, mesoderm, and endoderm, respectively.

Production of viral supernatants

Calcium phosphate transfection of 293 T cells protocol was used to produce viruses. 293 T cells were cultured in a DMEM complete medium (DMEM supplemented with 10% FBS, Glutamine, and Penicillin/Streptomycin). 293 T cells were seeded at a density of 12 × 10⁶ cells per 10 cm plate in a DMEM complete medium without antibiotics 24 h before transfection. On the day of transfection, the culture was changed to 10 ml DMEM medium containing 25 nM chloroquine one hour before the transfection. The transfection mix was prepared by adding 14 µg target plasmid DNA, 7 µg pVSVg, 14 µg pCMVΔR8.2, and ddH₂O to 438 µl in a 15-ml sterile tube, followed by adding 62 µl 2 M CaCl₂ to a final volume of 500 µl. Next, 500 µl of 2 × HBS were added to the above transfection mix dropwise with gentle mixing. The whole mixture was incubated for 15 min at RT and then added directly to the 293 T cells dropwise through the medium. After 24 h incubation, the medium was changed without chloroquine for cell culture. Supernatants were collected 48- and 72-hours post-transfection. Viruses were concentrated using an Amicon Ultra-15 centrifugal filter unit (Millipore) at 3000 g at 4 °C for 20 minutes. Concentrated virus particles were aliquoted in sterile 1.5-ml centrifuge tubes and stored at –80 °C.

Virus infection

For virus infection, 30–50 μl of the concentrated virus was used to infect 500,000 hESCs in the presence of 6 μg/ml polybrene. A total of 1 ml of cells (~500,000) were plated per well of a 6-well plate coated with Matrigel. Cells were mixed with the virus gently and incubated overnight at 37 °C. After 20–24 h post-infection, the viruses were removed by changing to a fresh medium. After 48 h post-infection, shRNA-transduced cells showed GFP positivity under a fluorescent microscope, and puromycin (1 μg/ml for hESCs) was added to kill non-transduced cells. RNA and protein were isolated after four days of transduction.

Protein synthesis measurements using puromycin incorporation (SUnSET)

Puromycin incorporation was performed as previously described³¹. H9 hESCs with or without ectopic expression of shRNA-resistant DDX18 WT and the DQAD mutant were infected with DDX18 shRNA viruses. Four days later, control KD and WT or mutant rescue cells were incubated with puromycin (1 μM final concentration) for 30 min, followed by protein extraction and western blotting using an anti-puromycin antibody (MABE343, Millipore). Coomassie blue staining was used as the loading control.

Real-time quantitative PCR

Total RNA was extracted from cells using TRIzol Reagent (Invitrogen) and subsequently column-purified with the RNeasy Kit (Qiagen) and treated with RNase-free DNase (Qiagen). One microgram of total RNA was reverse-transcribed into cDNA using the qScript cDNA SuperMix (Quantabio). For ChIP-qPCR analysis, 1 ng ChIP-DNA was used for each PCR. All qPCR analysis was performed on the QuantStudio 5 PCR system with Fast SYBR Green Master Mix (Applied Biosystems). The results were analyzed and normalized to β-ACTIN gene expression.

Western blotting

Total cellular protein was extracted using RIPA lysis buffer (20 mM Tris-HCl, pH 7.5; 150 mM NaCl; 1 mM EDTA; 1 mM EGTA; 1% NP-40; 1% sodium deoxycholate; and protease inhibitor cocktail) and then quantified using the Bradford method (Thermo Fisher). The protein lysates were separated by SDS–PAGE, electro-blotted onto polyvinylidene difluoride (PVDF) membranes (Millipore), probed with the indicated primary antibody followed by the HRP-conjugated secondary antibody, and then detected by enhanced chemiluminescence.

SILAC and mass spectrometry (SILAC-MS)

To perform the quantitative mass spectrometry-based whole-proteome comparison of control KD and DDX18 KD hESCs, we cultured control KD and DDX18 KD hESCs in SILAC heavy medium and SILAC light medium, respectively. For the SILAC heavy condition, hESCs were grown for three passages in DMEM/F12 with corresponding complete supplements but deficient in both L-lysine and L-arginine and supplemented with heavy 13C615N4 L-arginine and 13C615N2 L-lysine (Cambridge Isotope Laboratories). The medium was supplemented with 10% dialyzed FBS for SILAC (Thermo Fisher Scientific), and all other gradients followed the normal condition for conventional primed hESC culture (with regular L-lysine and L-arginine as SILAC light condition).

Complete labeling (>95%) was confirmed by MS before initiating the experiment. To quantitatively identify changes in protein levels upon DDX18 depletion, cells were infected with shRNA-expressing viruses and selected with puromycin for four days. After washing with ice-cold PBS, the control and DDX18-knockdown cells were harvested and counted. An equivalent number of SILAC-labeled cells (5 × 10⁶) were lysed and sonicated in RIPA buffer on ice. After centrifugation at 14,000 rpm for 10 min at 4 °C, proteins of an equal number of cells were combined, and the concentrations were estimated using a BCA protein assay kit (Thermo Fisher). The proteins were boiled with SDS loading buffer and subjected to SDS-PAGE followed by colloidal blue staining. Gel fragments were excised, washed, and sent for liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) to identify proteins.

Affinity purification coupled with mass spectrometry (AP-MS)

To isolate endogenous DDX18-associated protein complexes, affinity purification coupled with mass spectrometry (AP-MS) was performed as described¹⁰⁵ with some modifications. First, nuclear extraction was prepared from 10 × 15 cm dishes of 3xFlag-DDX18 hESCs with the nuclear extraction buffer [20 mM HEPES (pH 7.9), 25% glycerol (v/v), 0.42 M NaCl, 1.5 mM MgCl₂, 0.2 mM EDTA, 0.5 mM DTT (add fresh), 0.2 mM PMSF (add fresh), 1x protease inhibitor cocktail (add fresh)], then, nuclear extraction was pre-cleared with protein G agarose beads (Roche), using 100 μl beads (200 μl slurry) per 10 mg total protein. 25 μg of Flag antibody (Sigma, F1804) and 25 μg IgG (Millipore) per IP were incubated with the pre-cleared nuclear extraction in the cold room with rotation overnight without Benzonase nuclease to maintain the nucleic acid-mediated protein interactions. The next day, equilibrate protein G agarose beads (200 μl beads) with the nuclear extraction buffer, transfer the beads to the lysate/Ab mixture, and rotate in the cold room for 3 h. The immunoprecipitated samples were washed with wash buffer [20 mM HEPES (pH 7.9), 20% glycerol (v/v), 100 mM KCl, 1.5 mM MgCl₂, 0.2 mM EDTA, 0.5 mM DTT (add fresh), 0.02% NP40, 0.2 mM PMSF (add fresh), 1x protease inhibitor cocktail (add fresh)] five times, dry the beads by aspirating all the washing buffer using 20 μl tip at the last wash, beads were boiled with an equal volume of 2× SDS loading buffer for 5 min, and separated by SDS-PAGE. Finally, whole lanes were excised from the gel for LC-MS/MS to sequence and identify DDX18-associated proteins.

Immunoprecipitation (IP) and co-immunoprecipitation (Co-IP)

For immunoprecipitation, 2 μg Flag antibody was coupled to Dynabeads at 4 °C for 2 h in 200 μl lysis buffer, and nuclear extracts from 3xFlag-DDX18 hESCs were prepared and incubated with anti-Flag antibody overnight. For RNase A treatment, the immunoprecipitant was incubated in a lysis buffer containing 200 μg/ml RNase A (Sigma) at 37 °C for 10 min. The beads were washed four times with lysis buffer, resuspended in 1xSDS loading buffer, and boiled for analysis by SDS-PAGE. 3xFlag-DDX18 and NPM1 were identified by western blotting using anti-Flag M2 (Sigma) and anti-NPM1 (Santa Cruz, sc-47725) antibodies, respectively.

RNA-seq library preparation, sequencing, and data analysis

RNA was isolated with TRIzol (Invitrogen). RNA integrity was evaluated using a Eukaryotic RNA 6000 Nanochip on an Agilent 2100 Bioanalyzer (Agilent Technologies). Up to 1 µg of total RNA from each sample was used for library preparation with the TruSeq RNA Sample Preparation Kit (Illumina). A common adapter was used for all samples, and barcode sequences present in the reverse primers were introduced by 12–16 cycles of amplification. Each library was assessed for quality and size distribution using an Agilent High Sensitivity Assay bioanalyzer chip and quantified by PCR. Equimolar amounts of each barcoded library were mixed, and single-end sequenced on an Illumina HiSeq Sequencing System. For each sample, 14–21.7 M 50-nt reads were obtained, pre-processed with the FASTX-toolkit suite (http://hannonlab.cshl.edu/fastx_toolkit/), and aligned to the human genome (hg19 assembly) using TopHat mapper.

ChIP-sequencing and data analysis

ChIP-seq was performed as described previously¹⁰⁶. Chromatin was sheared by sonication to 100-500 base pair fragments. ChIP DNA libraries were made following the Illumina ChIP-Seq Library preparation kit (New England BioLabs). Samples were subjected to Illumina Solexa Cluster Station sequencing. Sequencing was performed on Illumina HiSeq2500 with a single read of 100 nt. Images acquired were processed through the image extraction pipeline and were aligned to the human genome hg19 using Bowtie. The reads that mapped to multiple locations in the genome were discarded. To map DDX18 ChIP-seq data to the rDNA, we used the consensus DNA sequence of the 43-kb ribosomal locus from NCBI (GeneBank ID: U13369.1). Employing this specific 43-kb region, we utilized the Bowtie algorithm with standard parameters for mapping to the Hg19 human genome build. The coverage was then visualized using IGV.

Hi-C sequencing and data analysis

Human ESCs were infected with shLuc and shDDX18 viruses. Samples were collected on day 4. ESCs were washed with PBS and fixed with 2% formaldehyde for 10 min at RT. Hi-C samples were prepared with the Arima Hi-C Kit, and libraries were prepared with the KAPA HyperPrep Kit following the manufacturer’s instructions. Paired-end reads were combined and pre-processed by Trim Galore (v.0.6.3) and Cutadapt programs. Hi-C library reads were processed using HiC-Pro (v3.0.0) and aligned to the hg19 reference genome using Bowtie2. All valid pairs of contacts were generated after filtering and removing duplicates in HiC-Pro. Valid contact read pairs of samples were applied to obtain the correlation coefficient matrices using Homer (v4.11), and PC1 values were exported in bedgraph.

Fluorescence recovery after photobleaching (FRAP) assay

H9-DDX18^GFP cells were cultured on a Matrigel (Corning) pretreated 35 mm 60 μ-dish (ibidi). 60% confluent cells were used for the FRAP experiment. All FRAP assays were performed on a ZEISS microscope with a 63× oil immersion objective and operated with the software. Spots of approximately 1 μm in diameter were photobleached with 20% laser power for 2 s using 488 nm lasers. Time-lapse images were acquired over 2 min after bleaching. Intensity recovery curves were normalized and corrected for photobleaching. The recovery curves were plotted by GraphPad Prism 9.

Measurement of centromere numbers and perinucleolar heterochromatin size

H9 hESCs were infected with DDX18 and control shRNAs as described previously. Three days after infection, cells were fixed and permeabilized for immunostaining. A primary antibody against NPM1 (Santa Cruz, sc-47725) was used for nucleolus staining. In addition, the primary antibody against HP1α (Abcam, ab202533) was used for perinucleolar heterochromatin marker, and antibodies against CENPA (GeneTex, GTX13939) and CENPT (Invitrogen, PA583540) were utilized for centromere staining. The images acquired with a confocal microscope (Leica SP8) were then measured by Image J software using particle analysis function. The average number of CENPA/CENPT foci and volume of HP1α foci in each cell were calculated and plotted.

Live cell imaging

All live-cell imaging was carried out on a Leica SP8 imaging system, equipped with a 60× /1.42 NA Plan Apo oil-immersion objective, 100×/1.40 NA Plan Apo oil-immersion objective (Olympus), and live-cell imaging environment control system (Live Cell Instrument). H9-DDX18^GFP cells were cultured on 29 mm No.1.5 glass-bottom (ibidi). In addition, DDX18-GFP knockin hESCs were cultured in mTeSR Plus medium. For imaging of the single time point, six z sections with 0.2 μm spacing were acquired, and for imaging of time, cells were imaged in a single layer per 15 s for a single channel or per 30 s for dual channels. For presentation in figures, raw data were processed by Fiji/ImageJ.

Structured illumination microscopy (SIM) procedure

For all SIM experiments, H9 hESCs were seeded on High-Performance No.1.5 18 × 18 mm glass coverslips (ibidi), fixed and permeabilized. To visualize DDX18, NPM1, and FBL, cells were blocked with 1% BSA for 1 h. Then, antibodies were 1:200 diluted in blocking buffer and incubated for 1 h at RT. After washing with 1xPBS 3 times, fluorescent secondary antibodies were 1:2,000 diluted in blocking buffer and incubated for 1 h at RT. Samples were mounted in ProLong Diamond Antifade Mountant medium (Thermo Fisher). Images were acquired with a 100×/1.4 Apo-TIRF objective lens on an Eclipse Ti microscope with 3D N-SIM (Nikon Instruments, Melville, NY). Imaging and SIM reconstruction parameters were set empirically for each field of view to achieve optimal signal intensity and reconstruction. Focus stabilization (Perfect Focus System, Nikon) was used to maintain focus during image acquisition. Z-stacks were collected with a step size of 0.2 μm and a total depth of 2.5–3 μm.

DNA fluorescence in situ hybridization with immunofluorescence (immuno-DNA FISH)

FISH probes were procured from Empire Genomics, and the BAC clone RP11-1025G19, including a part of HOXA9-13, was utilized to label the HOXA locus. The FISH procedure involved modifications to a previously employed protocol¹⁰⁷. Cells were firstly fixed in 3% paraformaldehyde in PBS for 10 min at room temperature and washed in PBST for 5 min 3 times. Cells were then permeabilized with 0.1% Triton-X at room temperature for 10 mins, then washed in PBST for 5 min 3 times. Cells were then incubated with rabbit anti-NPM1 antibodies diluted 1:1000 in blocking buffer overnight at 4 °C, washed with PBST 5 min for three times at room temperature, incubated for 1 hour at RT with Alexa488-labeled secondary goat anti-rabbit IgG (Thermo Fisher). Cells were then subjected to the FISH protocol. Coverslips were treated with PBST washing and ice incubation with 0.1 M HCl/0.7% Triton X-100/2× SSC for 10 min. Afterwards, coverslips were washed with 2× SSC, and 2 μl FISH probes were added, followed by mounting and sealing with rubber cement. Samples were denatured on a heat block at 76 °C for 3 min and then hybridized for 18–72 h at 37 °C on a heat block. Coverslips underwent washing at 37 °C in 2× SSC and 60 °C in 0.1× SSC, and at room temperature in 4× SSC. Finally, coverslips were mounted in DAPI-containing antifade mounting media, and images were captured using a Nikon A1plus-RSI confocal microscope.

Hybridization chain reaction RNA fluorescence in situ hybridization (HCR RNA-FISH)

HCR RNA-FISH probes for each target mRNA were designed and ordered from Molecular Instruments (www.molecularinstruments.com). Human ESCs (H9) were cultured in mTeSR1 Plus medium (StemCell Technologies) on Matrigel-coated 6-well plates. Lentiviral shDDX18 and shLuc were added to H9 cells with polybrene. After 24 h post-infection, viruses were removed, and puromycin (1 µg/ml) was added to the fresh medium to kill non-transduced hESCs for 24 h. On Day 3, the cells were trypsinized, counted, and seeded into 8-well chambered glass bottom #1.5 µ-slides (ibidi). Cells were grown to the desired confluency (~70%) for 24–48 h with puromycin in the medium. Growth media were aspirated, and each chamber was washed with DPBS. Cells were fixed with 4% formaldehyde and then permeabilized overnight with 70% ethanol at −20 °C. The Multiplexed HCR RNA-FISH (V3.0) encompassing the detection and amplification stages was performed following the protocol provided by the Molecular Instruments. DAPI was used to counterstain the nucleus. Nucleolus Bright Green Dye (Dojindo Cat#N511), which reacts to RNAs present in both nuclear (predominantly nucleolar) and cytoplasmic RNAs, was used to detect nucleolus when co-stained with DAPI. The cells were examined under a Stellaris confocal microscope.

Individual nucleotide-resolution UV crosslinking and immunoprecipitation (iCLIP) and analysis

The iCLIP protocol was performed as described previously¹⁰⁸ with several modifications. First, 3xFlag-DDX18 H9 cells were irradiated by UV once with 160 mJ/cm² using Stratalinker 1800 at 254 nm. Next, 3xFlag-DDX18 was immunoprecipitated with protein A Dynabeads (10002D, Invitrogen) conjugated to mouse anti-Flag (Sigma-Aldrich, F1804-1MG). The region corresponding to 75–100 kDa complexes was excised from the membrane to isolate the RNA and sequenced using Illumina HiSeq, generating 50-nt single-end reads. The iCLIP analysis is based on the instructions of the CLIP Tool Kit (CTK, https://zhanglab.c2b2.columbia.edu/index.php/CTK_Documentation). Analysis of the reproducibility of crosslinked sites, identification of the significant iCLIP crosslinked clusters, and z-score analysis of enriched pentamers were performed as described. Data were processed by the iCount software (https://github.com/tomazc/iCount). Duplicates were collapsed before alignment. After collapsing duplicates, barcode sequences were trimmed. Adapters were cleaned after adapter removal. Alignment was done according to CTK instructions. Sequences were aligned to hg19 using Burrows-Wheeler Aligner (BWA) and reads shorter than 15 bp were removed. Peak calling was done using tag2Peak.pl script with statistical assessment.

Protein expression and purification

For protein expression, Escherichia coli Transetta (DE3) cells (TransGen Biotech, China) bearing expression plasmids were grown to OD 0.6–0.8 at 37 °C in LB media containing kanamycin and chloramphenicol and then induced at 18 °C with 0.1 mM Isopropyl β-D-1-thiogalactopyranoside for an additional 16–18 h. Cell pellets were harvested and resuspended in lysis buffer [50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 10% glycerol, 1 mM phenylmethylsulfonyl fluoride (PMSF), 2 mM 2-mercaptoethanol, and protease inhibitor cocktail]. Cells were then broken by sonication and cleared by ultracentrifugation at 18,000 rpm for 50 min at 4 °C. Next, the supernatant was subjected to Ni-NTA agarose beads (QIAGEN, USA) for affinity purification. The resin was then washed with lysis buffer, and the bound protein was eluted in elution buffer (50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 10% glycerol, and 300 mM imidazole). For NPM1-RFP, NPM1-CFP, RFP-FBL, and RFP-HP1α, the His-SUMO tag was removed by on-column digestion with Ulp1 protease at 4 °C for 12 h. Proteins were further purified by size-exclusion chromatography on a Hiload Superdex 75 or Hiload Superdex 200 column (GE Healthcare, USA), equilibrated with 25 mM Tris-HCl, pH 8.0, and 500 mM NaCl.

Furthermore, full-length DDX18-GFP proteins and their truncations or mutants were injected on a Mono S 5/50 GL cation-exchange column (GE Healthcare, USA) and further purified by size-exclusion chromatography on a Superdex 200 column. NPM1-RFP and NPM1-CFP were injected on a Mono Q 5/50 GL anion-exchange column (GE Healthcare, USA) and further purified by size-exclusion chromatography on a Superdex 200 column. The purified proteins were concentrated and stored in the high salt buffer (25 mM Tris-HCl, pH 8.0, 500 mM NaCl, and 2 mM DTT) at −80 °C to inhibit droplet formation. Concentrations were measured by UV absorbance at 280 nm.

Phase separation assays in vitro

Purified human DDX18-GFP, NPM1-RFP, NPM1-CFP, RFP-FBL, and RFP-HP1α proteins were mixed with varying volumes of salt buffer (25 mM Tris-HCl, pH 8.0, and varying NaCl concentrations) to obtain desired protein/salt concentrations. To visualize the droplet formation, 3–6 μl of the protein solution was immediately loaded onto a homemade chamber comprising a glass slide with a coverslip fixed by two parallel strips of double-sided tape. For HP1α droplet formation, we used nonphosphorylated HP1α and added 5% PEG8000 to facilitate its phase separation. Fluorescent images were taken by a Zeiss LSM 710 microscope.

DDX18 protein disorder plots

Intrinsic disorder regions (IDR) for DDX18 (NCBI Reference Sequence: NP_006764.3) were performed using the PONDR VSL2 webtool (http://www.pondr.com/). Prion-like domains (PLD) prediction was performed on PLAAC. Pi-Pi (π-π) contacts predictor was used online on Forman-Kay’s laboratory website (http://pound.med.utoronto.ca/~JFKlab/). Finally, CIDER was used to analyze the net charge per residue (NCPR) distribution.

Quantification and statistical analysis

Statistical analyses were carried out using Excel or R (version 3.3.0). Data are presented as mean ± SD. The statistical tests are analyzed using the student’s t-test if it is not stated in the relevant figure legends. All the statistical details can be found in the figure legends or results.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The DDX18 affinity purification and SILAC mass spectrometry (MS) data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD057813. The RNA-seq, ChIP-seq, iCLIP-seq data generated in this study have been deposited in the Gene Expression Omnibus (GEO) database under accession code GSE182804 and Hi-C sequencing GSE182855. Source data are provided with this paper.

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Acknowledgements

We thank Drs. Ling-Ling Chen for providing NPM1-RFP and RFP-FBL bacterial expressing plasmids, Dan Li at the Wang lab for eCLIP-seq expertise, and Tajda Klobucar at the Modic lab for technical advice on HCR-FISH. We also thank Columbia Confocal and Specialized Microscopy Shared Resource for 3D-SIM imaging, Columbia Medicine Microscopy Core for Stellaris confocal microscope services (funded by NIH-1S10OD032447), the CCTI Flow Cytometry Core for cell sorting services. Laboratories of Drs. Chen and Li were supported by grants (2023YFA1800403 to Y.C., NSFC32071195, 31900934, and 23QC1400600 to Y.L.). Research in the Wang laboratory was supported by NIH (GM129157, HD095938, HD097268, HD114122, and R21HD116446).

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These authors contributed equally: Xianle Shi, Yanjing Li, Hongwei Zhou, Xiukun Hou.

Authors and Affiliations

Department of Medicine, Columbia Center for Human Development and Stem Cell Therapies, Columbia University Irving Medical Center, New York, NY, USA
Xianle Shi, Hongwei Zhou, Jihong Yang, Vikas Malik, Xin Huang & Jianlong Wang
School of Life Sciences, Tsinghua University, Beijing, China
Xianle Shi & Wei Xie
Shanghai Institute of Biochemistry and Cell Biology, Center for Excellence in Molecular Cell Science, Chinese Academy of Sciences, Shanghai, China
Yanjing Li & Yong Chen
State Key Laboratory of Systems Medicine for Cancer, Shanghai Cancer Institute, Shanghai, China
Yanjing Li
Department of Thyroid and Neck Cancer, Tianjin Medical University Cancer Institute and Hospital, Tianjin, China
Xiukun Hou
Black Family Stem Cell Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA
Francesco Faiola & Junjun Ding
Guangzhou Institutes of Biomedicine and Health, Chinese Academy of Sciences, Guangzhou, China
Xichen Bao
The Francis Crick Institute and University College London, London, UK
Miha Modic
College of Life Sciences, Nankai University, Tianjin, China
Weiyu Zhang & Lingyi Chen
Department of Medicine, Sandra and Edward Meyer Cancer Center, Weill Cornell Medicine, New York, NY, USA
Syed Raza Mahmood & Effie Apostolou
Department of Molecular Medicine/Cell Systems and Anatomy, University of Texas Health Science Center at San Antonio, San Antonio, TX, USA
Feng-Chun Yang & Mingjiang Xu

Authors

Xianle Shi
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Yanjing Li
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Hongwei Zhou
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Xiukun Hou
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Jihong Yang
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Vikas Malik
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Francesco Faiola
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Junjun Ding
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Xichen Bao
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Miha Modic
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Weiyu Zhang
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Lingyi Chen
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Syed Raza Mahmood
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Feng-Chun Yang
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Mingjiang Xu
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Wei Xie
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Xin Huang
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Yong Chen
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Jianlong Wang
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Contributions

X.S., Y.L., H.Z., and X. Hou designed and conducted the experiments and wrote the manuscript with contributions from all other authors. V.M., F.F., J.D., and L.C. performed the experiments. J.Y., M.M., X. Huang, X.B., and W.Z. performed the computational analysis. S.R.M. and E.A. performed Hi-C data analyses presented in Figs. 6d, e and S6b-f,l. F.-C.Y., M.X., and W.X. provided technical advice and helpful discussions. J.W. conceived the project and, together with Y.C., designed the experiments, interpreted the results, and prepared and approved the manuscript.

Corresponding authors

Correspondence to Yong Chen or Jianlong Wang.

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Nature Communications thanks Fulai Jin, and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

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Shi, X., Li, Y., Zhou, H. et al. DDX18 coordinates nucleolus phase separation and nuclear organization to control the pluripotency of human embryonic stem cells. Nat Commun 15, 10803 (2024). https://doi.org/10.1038/s41467-024-55054-8

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Received: 14 August 2024
Accepted: 26 November 2024
Published: 30 December 2024
DOI: https://doi.org/10.1038/s41467-024-55054-8

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