Introduction

Chromatin dynamically regulates DNA accessibility to orchestrate essential processes such as transcription, replication, and DNA repair^1–5. These processes are dynamically regulated by chromatin-associated complexes, such as the human silencing hub (HUSH) complex⁶, which control the accessibility of genetic material to regulatory machinery. The Microrchidia (MORC) protein family, characterized by a conserved N-terminal GHL-ATPase domain and a structurally diverse C-terminal region, regulates gene expression and chromatin organization^7–10. These C-terminal domains are thought to contribute to dimerization and functional specificity^11–14. Notably, MORC2 has been identified as a component of the HUSH complex and is required for HUSH-mediated transcriptional repression¹².

GHL ATPases, including DNA gyrase, Hsp90, and MutL, rely on ATP-dependent dimerization to drive conformational changes essential for substrate engagement and enzymatic regulation^15–19. Similar mechanisms have been proposed for MORC family members. In humans, the MORC family comprises five members—MORC1, MORC2, MORC3, MORC4, and SMCHD1, and different MORC paralogs target distinct genomic loci, including transposable elements, viral DNA, and long-exon genes^7,20,21. The conserved N-terminal GHL-ATPase domain of MORC family proteins has been extensively characterized, whereas the evolutionarily diversified, multi-domain C-terminal regions are thought to act as key determinants of chromatin targeting and functional specificity. Despite these shared structural features, the molecular basis for MORC-mediated transcriptional regulation remains poorly understood, partly due to limited structural insight into the full-length proteins and C-terminal regions.

MORC2 has recently been identified as a key regulator of chromatin architecture and gene silencing, primarily through its recruitment to HUSH-target loci via C-terminal CC2 domain-mediated interactions with TASOR and MPP8¹². While its silencing activity was initially attributed to the N-terminal ATPase domain, genetic complementation studies have revealed that the CW domain and all three coiled-coil (CC) regions are also indispensable for its function^12,13. Beyond its structured domains, sequence analyses and structural predictions have uncovered an extended intrinsically disordered region (IDR) spanning the central to C-terminal regions of the protein, suggesting potential regulatory mechanisms by facilitating multivalent and dynamic interactions beyond enzymatic activity²².

Transcriptional silencing is frequently spatially organized within membrane-less nuclear compartments, often driven by the process of biomolecular condensation^23,24. Sequence analyses of MORC2 reveal an extended intrinsically disordered region (IDR) spanning the central to C-terminal domains, suggesting a capacity for the multivalent, weak interactions that underpin phase separation^25,26. While IDRs are common in chromatin-associated proteins, their presence alone does not guarantee condensation; rather, phase separation often emerges from the coordinated interplay between disordered segments and structured oligomerization hubs. How such mesoscale organization modulates MORC2’s enzymatic turnover and its ability to remodel the genome remains a fundamental question in the field.

In this study, we demonstrate that MORC2 undergoes phase separation to form dynamic nuclear condensates. We define the structural basis for this behavior, identifying a dimeric CC3 scaffold that, together with multivalent interactions between the IDR and a newly identified IDR-binding domain (IBD), drives condensation. We further show that DNA binding acts as a molecular scaffold to promote MORC2 assembly, which in turn allosterically enhances its ATPase activity. Using a combination of cellular “killswitch” assays and in vivo models, we establish that the dynamic material properties of MORC2 condensates—rather than mere protein recruitment—are strictly required for transcriptional regulation. Finally, we provide mechanistic insights into how neuropathic disease-associated mutations in the MORC2 N-terminus perturb this biophysical balance, linking condensate dysregulation to human pathology.

Results

Coiled-coil 3 (CC3) mediates dimerization of full-length MORC2

To determine whether MORC2 has N– and C-terminal dimerization interfaces similar to those in other GHL-type ATPases²⁷, which previously identified the role of CC3 (previously designated as CC4) in MORC2 dimerization¹⁴, we established an in vitro system to purify full-length (FL) human MORC2 (MORC2FL, residues 1-1032) from HEK293F cells, yielding nucleic acid-free, highly pure proteins for structural and functional analyses (Figure 1a, b). Analytical size-exclusion chromatography coupled with static light scattering (SLS) demonstrated that MORC2FL forms dimers, with a molecular weight of approximately 239 ± 1 kDa under buffer conditions containing 1000 mM NaCl even in the absence of ATP (Figure 1c). Notably, the addition of 601 DNA resulted in a detectable increase in molecular weight (303 ± 2 kDa), indicative of DNA-protein complex formation, but did not disrupt the dimeric assembly of MORC2 in the same buffer condition (Figure 1c).

Figure 1.

To elucidate the atomic structure of MORC2FL, we initially employed negative-stain electron microscopy to reveal that MORC2FL particles were poorly resolved and exhibited considerable heterogeneity in buffer containing 300 mM NaCl. Notably, a subset of particles displayed a morphology consistent with dimer formation (Figure S1a). To further investigate this, we performed structural prediction of MORC2FL using AlphaFold3 and identified a predicted structure that closely resembled the observed reconstruction (Figure S1b). We then examined the oligomerization properties of various MORC2 constructs. MORC2^1-900 (FLΔCC3-IBD domain) was found to exist as a monomer, whereas MORC2^901-1032 and MORC2^901-1003 exhibited dimeric properties, confirming the critical role of the CC3 domain (residues 901-1003) in mediating MORC2 dimerization (Figure 1c, and Figure S1c).

To gain further structural insights, we determined the 3.1 Å crystal structure of the CC3 dimer using X-ray diffraction (Table S1). The structure revealed that the dimer interface is stabilized predominantly by hydrophobic interactions. Three key hydrophobic residues—L911, L915, and F922—form distinct layers within the dimer interface (layers 1-3, Figure 1e, and Figure S1e, f). The side chains of these residues are buried within the dimerization interface, creating a hydrophobic core critical for maintaining the stability of the dimer.

We systematically introduced single-point mutations in these hydrophobic residues to assess their contributions to dimerization. SLS analysis demonstrated that the wild-type (WT) MORC2^901-1032 (with Trx-tag) fragment predominantly exists as a dimer (∼61 kDa). In contrast, all three key mutants (L911Q, L915Q, and F922Q) displayed altered molecular weight distributions indicative of disrupted dimer formation and enhanced higher-order oligomerization (more than 61 kDa) (Figure S1g). However, none of these mutations completely disrupted dimerization or converted the dimer to a monomeric state (Figure S1g). Further analyses of additional single-site mutants (e.g., I908Q, Y921A, F951Q, Y954A, L958Q) showed minimal effects on dimer stability, underscoring the central importance of L915 and its neighboring hydrophobic layers in stabilizing the CC3 dimer (Figure S1g). Moreover, we examined the dimeric status of truncated CC3 constructs (residues 901-975), which confirmed that the CC3 N-terminal region is sufficient to drive dimer formation (Figure 1f). To delineate the structural basis of these interactions, we mapped the positions of hydrophobic residues within the CC3 backbone (Figure S1e). The distances between interacting residues, such as I908-I908’ (3.4 Å), L911-L915’ (3.7 Å), F922-F922’ (3.9 Å), and L958-L958’ (4.0 Å), revealed a tightly packed hydrophobic network. The robustness of this network ensures the integrity of the CC3 dimer, even under varying salt concentration and protein concentration tested in vitro conditions. These findings collectively suggest that the CC3 domain plays a critical role in MORC2 dimerization. The dimeric assembly of CC3 is essential for maintaining the structural integrity of the protein, which likely underpins its functional activities, including associations with chromatin¹⁴.

MORC2 forms condensates both in vivo and in vitro

In the predicted structure of MORC2FL, we observed an extended unstructured region at the C-terminus of the protein (Figure S1b). This region is classified as an intrinsically disordered region (IDR), a protein architecture frequently reported to facilitate multivalent interactions and can act as a scaffold for phase separation^{25, 28}. Specifically, IUPred2A analysis predicts that the residues 593-735 predominantly exhibit high disorder scores, prompting us to define this segment as the IDR domain (Figure 1a).

In vitro, sedimentation assays showed that when the salt concentration was reduced to 150 mM, more than half of the EGFP fused MORC2FL (EGFP-MORC2FL) precipitated into the pellet fraction (Figure 2a, b). Fluorescence imaging revealed that at this near-physiological ionic strength, EGFP-MORC2FL spontaneously demixed to form well-defined spherical droplets (Figure 2c, d). These droplets exhibited a characteristic polydispersity in size and showed significant coarsening as protein concentration increased (Figure S2a). Sedimentation assays further showed that the pellet fraction of EGFP-MORC2FL after centrifugation was concentration-dependent (Figure S2b). The EGFP-MORC2FL-enriched condensates exhibited dynamic behavior, including droplet fusion (Figure S2c) and fluorescence recovery after photobleaching (FRAP) (Figure 2e, f), indicating a dynamic state.

Figure 2.

Using antibody labeling, we observed that endogenous MORC2 forms discrete, dense punctate structures within the nuclei of HeLa cells (Figure 2g). The specificity of these puncta was confirmed by their absence in MORC2-knockout (KO) HeLa cells (Figure S2d, e), suggesting that MORC2 may natively organize into higher-order nuclear condensates under physiological conditions. To further characterize these assemblies, we transfected HeLa cells with EGFP-MORC2FL and observed the formation of prominent, droplet-like nuclear condensates (Figure 2h). Notably, consistent with the endogenous staining patterns, EGFP-MORC2FL condensates preferentially localized to regions of reduced chromatin density, as visualized by the H2A-mCherry signal. This distinct spatial distribution implies a potential role for MORC2 condensates in organizing or regulating the heterochromatin landscape. FRAP analysis of these EGFP-MORC2FL puncta revealed fluorescence recovery on a timescale of seconds (Figure 2i, j). Furthermore, these puncta underwent active fusion events (Figure S2f), a hallmark of dynamic liquid-like condensates with a highly mobile internal environment.

To determine whether MORC2 forms biomolecular condensates under physiological conditions, we generated mice expressing EGFP-MORC2 protein from the endogenous locus (Figure S3a). We focus on neurons in the brain and spinal cord as missense mutations in MORC2 cause neuropathies including Charcot-Marie-Tooth and spinal muscular atrophy disease^11,13. Imaging of NeuN-positive neurons revealed that EGFP-MORC2FL condensates occur in neurons under endogenous expression level (Figure 2k). FRAP analysis in freshly prepared brain slices further demonstrated rapid and partial fluorescence recovery, supporting the dynamic nature of endogenous MORC2 assemblies (Figure 2l). In addition, unlike MORC3, which shows cell cycle-dependent nuclear puncta formation²⁹, MORC2 condensates were absent in mitotic cells (Figure S3b). Together, these results indicate that MORC2 forms phase-separated biomolecular condensates under physiological conditions in vivo and in vitro.

The MORC2 C-terminal domain (CTD) drives condensate formation

To identify the key determinants governing the assembly of MORC2 nuclear condensates, we partitioned the protein into two functional modules based on its domain architecture: the N-terminal domain (NTD, residues 1–472), comprising the ATPase and CC1 motifs, and the C-terminal domain (CTD, residues 473–1032), which encompasses the IDR and multiple coiled-coil regions. A series of EGFP-tagged truncation constructs were then expressed in HeLa cells to systematically map the regions essential for condensate formation (Figure 3a). Deletion of the CTD completely abolished puncta formation, resulting in a diffuse localization across the nucleus and cytoplasm (Figure S4a). By contrast, NTD deletion had negligible effects on nuclear localization or the capacity for condensation. To exclude the possibility that the NTD’s diffuse pattern was merely due to its loss of nuclear entry, we appended an exogenous nuclear localization signal (NLS, PKKKRKV³⁰) to the NTD; however, while this redirected the protein to the nucleus, it failed to restore puncta (Figure 3a), confirming that the CTD is the primary driver of MORC2 condensation. To assess whether the CTD is intrinsically prone to undergo condensation in vitro, we purified a CTDΔCW (residues 537–1032) variant from E. coli, as the full CTD exhibited suboptimal biochemical stability (Figure S5a, b). At 150 mM NaCl, Cy3-labeled CTDΔCW readily formed discrete condensates in a concentration-dependent manner (Figure 3b, c), and displayed a sedimentation profile characteristic of phase-separated assemblies (Figure 3d, e). Fluorescence microscopy and static light scattering (SLS) confirmed that Cy3 labeling did not perturb its condensation behavior, morphology, or molecular weight distribution (Figure S5c–f).

Figure 3.

Further dissection of the CTD revealed that deletion of either the CC2 or TCD domains did not impair the formation of nuclear puncta (Figure S4b). In contrast, removal of the IDR-NLS region (residues 593-761) abolished both puncta formation and nuclear localization (Figure S4b). Notably, appending an exogenous NLS (PKKKRKV) to CTDΔIDR-NLS restored its nuclear entry but failed to rescue condensate assembly (Figure S4c), identifying the IDR as a prerequisite for condensation. Similarly, deletion of the CC3-IBD segment maintained nuclear localization but eliminated puncta formation, indicating that this region is also indispensable (Figure S4b). Intriguingly, neither the IDR-NLS fragment nor the CC3-IBD fragment alone (even when NLS-fused) was sufficient to trigger condensation (Figure S4d), implying a cooperative mechanism between these modules. Consistent with these cellular findings, in vitro assays showed that neither the MORC2ΔCC3-IBD (residues 1–900) nor the isolated CC3-IBD fragment could undergo condensation (Figure S6a-b). However, mixing the IDR with the IBD fragment reconstituted robust condensate formation at higher concentration (Figure S6c), confirming their synergistic role. Finally, the CW domain appears to modulate condensate morphology, as its deletion led to enlarged and frequently fused puncta. We conclude that the IDR, CC3, and IBD constitute the minimal structural core required for MORC2 condensation, while the CW domain serves to restrict condensate overgrowth and maintain proper morphology (Figure 3a, and Figure S4e).

Multivalent interactions between the IDR and IBD promote MORC2 condensation

Next, we elucidated the mechanistic basis for MORC2 assembly mediated by the IDR and CC3-IBD. We hypothesized that multivalent interactions between these modules drive the formation of a condensed phase that coexists with a dilute aqueous phase. Using nuclear magnetic resonance (NMR) spectroscopy, we characterized the interaction between the IDR and the C-terminal segment (residues 901–1032). ¹⁵N-HSQC titration assays using the ¹⁵N-labeled CC3 (residues 901-1003) or the 1004-1032 fragment revealed that residues 1004-1032 constitute the primary binding site for the IDR, which we termed the IDR-Binding Domain (IBD), whereas the CC3 region exhibited negligible interaction (Figure 4a-c).

Figure 4.

Further subdivision of the IDR into three segments—IDRa (residues 593-643), IDRb (residues 644-694), and IDRc (residues 695-735)—pinpointed IDRa as the key region responsible for interaction with IBD (Figure 4a, d-f). As is typical for IDR-mediated interactions involved in condensation, the observed chemical shift perturbations (CSPs) were relatively small but highly region-specific. Notably, the CSP profile of IDRa closely mirrors that of the full-length IDR, whereas IDRb and IDRc showed minimal perturbations (Figure S7). Detailed sequence analysis revealed that the CSPs are clustered around charged residues (D, E, K, R) and their neighboring residues (Y, T, N, I, L, P). This suggests that the association is driven by a concerted action of multiple non-covalent forces: beyond the foundational electrostatic attraction, the involvement of aromatic (Y) and aliphatic (I, L, P) residues points to the contribution of π-π stacking, cation-π interactions, and hydrophobic effects, while polar residues (T, N) likely facilitate intermolecular hydrogen bonding. This hierarchical assembly, characterized by both charge complementarity and hydrophobic-like contributions, aligns with the ‘sticker-and-spacer’ model. In this framework, these multivalent residues act as ‘stickers’ that provide the requisite valency to scaffold a robust yet dynamic network, while the intervening sequences serve as ‘spacers’ to maintain the fluidity necessary for the condensed state (Figure 4g). Collectively, these quantitative CSP data support a model of weak, dynamic, and region-specific interactions, a hallmark of IDR-driven condensation mechanisms^31–33.

To assess the role of IDRa-IBD multivalent interactions in cells, we transfected HeLa and HEK293T cells with MORC2FL variants harboring individual deletions of the IBD, IDRa, IDRb, or IDRc regions. Deletion of the IBD abolished condensate formation in both HeLa cells (Figure 3a, and Figure S4e) and HEK293T cells (Figure S8). Similarly, deletion of IDRa significantly impaired condensation, resulting in a marked decrease in both the number and area of nuclear puncta, whereas IDRb or IDRc deletions had minimal impact (Figure 4h, i). Based on these findings, we propose a model in which multivalent ‘sticker’ interactions between IDRa and the IBD drive MORC2 intermolecular assembly, a process further potentiated by CC3-mediated dimerization, which collectively provides the valency threshold required for condensation (Figure 4j).

DNA binding promotes MORC2 condensation and stimulates ATPase activity

Notably, MORC2 condensates form exclusively within the nuclei of transfected HeLa cells, implying that chromatin-associated factors—most likely DNA—serve as essential cues for their nucleation or stabilization. While the N-terminal CC1 domain was previously identified as a DNA-binding module¹³, we hypothesized that the full-length MORC2 architecture utilizes additional interfaces to facilitate complex chromatin interactions.

To test this, we performed a comprehensive domain screen using electrophoretic mobility shift assays (EMSA). Beyond confirming the robust DNA-binding activity of the CC1 domain (Figure 5a, b), our screen revealed that the CC2, IDR, and TCD (Tudor Chromo-like domain) also possess significant DNA-binding capabilities (Figure 5c-e). In contrast, the CC3–IBD fragment showed no affinity for DNA (Figure 5f). These results indicate that MORC2 employs a multivalent DNA-binding architecture distributed across both its N-terminal and C-terminal regions.

Figure 5.

Building on this, we investigated whether these multivalent DNA-protein interactions could scaffold MORC2 condensation in vitro. At 0.9 μM, MORC2FL alone failed to form discrete assemblies; however, the addition of 25 nM 601 DNA induced robust condensate formation, demonstrating that DNA acts as a molecular seed to promote MORC2 phase separation (Figure 5g, h). Interestingly, while the NTD (residues 1–536) binds DNA, it remains diffuse (Figure S6d,e), whereas the CTDΔCW (residues 537–1032) undergoes DNA-driven condensation (Figure 5i). Systematic evaluation across a matrix of protein and DNA concentrations yielded a phase diagram mapping the transitions between clear solutions, liquid-like condensates, and amorphous precipitates, consistently showing that DNA significantly lowers the concentration threshold for MORC2 condensation (Figure S9). This functional divergence suggests a division of labor within MORC2: while multiple domains provide DNA affinity, the CTD provides the essential protein-protein interaction network required for higher-order assembly.

Finally, we explored how this assembly impacts the MORC2 enzymatic core. The addition of 601 DNA markedly stimulated the ATPase activity of wild-type MORC2, but not the ATP-binding-deficient N39A mutant (Figure 5j). This reveals a synergistic interplay: DNA binding does not merely recruit MORC2 to chromatin; it acts as a molecular scaffold that facilitates condensation, which in turn may optimize the productive conformation or local density required for allosteric activation of the ATPase domain. In conclusion, these findings highlight a coordinated mechanism that integrates DNA binding, condensation, and enzymatic activity, providing a biophysical basis for the multivalent assembly and allosteric regulation of MORC2 on chromatin.

Condensate formation and dynamics are required for MORC2-mediated transcriptional regulation

Establishing a causal link between biomolecular condensation and biological function—particularly in transcriptional regulation—remains a central challenge. A key question is whether MORC2 condensates represent discrete functional entities or merely inactive byproducts. To address this, we sought to decouple condensate formation from internal dynamics, allowing for a rigorous interrogation of their functional relevance. While deletion of the CC3 domain abolishes MORC2 condensation, CC3 is also indispensable for dimerization, confounding functional interpretations. Furthermore, although deleting IDRa markedly weakens MORC2 condensation, a tool to specifically manipulate the solubility and internal mobility of these assemblies was previously lacking.

To overcome these limitations, we employed a micropeptide-based ‘killswitch’ (KS) strategy fused to the C-terminus of MORC2 (Figure 6a). This approach selectively reduces condensate dynamics without disrupting protein expression, folding, or domain architecture³⁴. Unlike the ΔCC3 or ΔIDRa mutants, the MORC2+KS variant robustly form nuclear condensates but exhibits markedly reduced internal dynamics, as demonstrated by FRAP analyses showing minimal fluorescence recovery (Figure 6b, c). This strategy allowed us to perturb the material properties of the condensates independently of structural integrity.

Figure 6.

To evaluate how the modulation of MORC2 condensates influences its gene regulatory capacity, we compared the transcriptional consequences of rescuing MORC2-knockout HeLa cells with MORC2FL, condensation-deficient mutants (ΔCC3 and ΔIDRa), and the dynamics-defective MORC2+KS (Figure 6d). Bulk RNA-seq was performed 48 hours post-transfection. We hypothesized that impairing either the assembly or the fluidity of the condensates would compromise MORC2-dependent gene regulation. Notably, despite being expressed at substantially higher levels than MORC2FL (Figure 6e), all three mutants showed a striking and consistent failure to restore the MORC2-dependent transcriptional profile (Figure 6f–h). This defect was particularly pronounced for transcriptionally repressed genes, including two sets of high-confidence MORC2 targets identified in prior studies^{12, 22} (Figure 6i, and Figure S10). Gene Ontology analysis revealed that the MORC2-regulated genes are involved in DNA damage responses, viral infection, and cell cycle regulation (Figure 6g). These findings demonstrate that neither increased protein abundance nor the mere presence of static, condensate-like structures is sufficient to restore MORC2 function.

Collectively, these data support a model in which the dynamic nature of MORC2 condensates is essential for full transcriptional activity. While soluble MORC2 complexes are likely involved in initial target recognition, our results indicate that proper condensate formation—and critically, internal fluidity—are required for effective transcriptional control. The inability of the MORC2+KS mutant to rescue transcriptional defects, despite forming visible nuclear assemblies, strongly argues against a model where MORC2 condensates are merely inert byproducts of its activity.

Pathogenic MORC2 variants differentially modulate condensation, DNA binding, and ATPase activity

MORC2 is essential for gene silencing at H3K9me3-marked loci and retrotransposon repression^6,35–37. Pathogenic mutations in its GHL-type ATPase domain have been linked to severe neuropathies and cancer^38–42. To explore whether these mutations influence the condensation behavior and biochemical activities of the full-length protein, we examined a panel of disease-associated variants, including those linked to Charcot–Marie–Tooth disease type 2Z (CMT2Z: E236G, R252W, Q400R, D466N) and spinal muscular atrophy (SMA: S87L, S218L, F256L, R266A, T424R) (Figure 7a).

Figure 7.

In HeLa cells, all variants retained the ability to form nuclear puncta. However, the CMT2Z-linked E236G and, notably, the SMA-linked T424R mutants significantly increased the proportion of cells exhibiting nuclear condensates (Figure 7b). These observations suggest that the N-terminal region plays a nuanced role in modulating the phase boundary of MORC2. Specifically, previous structural studies indicated that the T424R mutation has minimal effects on the overall N-terminal architecture but primarily alters the assembly and dissociation kinetics of the N-terminal dimer¹³. This suggests that the conformational cycling or dimerization state of the N-terminus directly contributes to the macroscopic regulation of MORC2 condensates. Interestingly, while deletion of the N-terminal region did not abolish puncta formation, FRAP analyses revealed a marked loss of condensate fluidity compared with the full-length protein (Figure S11). This highlights the N-terminus as a critical regulator of the material properties of MORC2 assemblies. Consequently, we evaluated the dynamics of E236G and T424R condensates; FRAP measurements showed substantially reduced recovery for E236G, indicating decreased internal fluidity, whereas T424R exhibited minimal effects (Figure 7e, f).

In vitro ATPase assays revealed that T424R significantly increases enzymatic activity (Figure 7c), consistent with its reported role in promoting ATPase dimer cycling¹³. Additionally, S218L and F256L variants enhanced ATP hydrolysis, whereas others, such as Q400R and D466N, showed activity comparable to WT (Figure 7c). Due to severe solubility issues, E236G could not be biochemically characterized in vitro (Figure S12e).

We next assessed DNA binding affinity of MORC2 variants using fluorescence polarization (FP) assays. Despite subtle differences, none of the variants showed statistically significant changes in DNA-binding capacity (Figure 7d), indicating that these mutations primarily perturb enzymatic turnover and condensation dynamics rather than DNA recruitment.

In summary, our results support a model in which MORC2 mediates transcriptional regulation through its intrinsic ability to undergo biomolecular condensation. The N-terminal ATPase domain undergoes conformational cycling between open and closed states, while the C-terminal CC3-mediated dimer acts as a stable scaffold for condensate formation (Figure 7g). MORC2 condensation is further fine-tuned by multivalent interactions between the IBD and IDRa regions, as well as by DNA binding. Together, these features enable MORC2 to bridge distal chromatin regions and establish repressive condensates, underscoring its central role in chromatin organization and genome stability

Discussion

Biophysical Principles of MORC2 Organization

While the MORC family of ATPases is recognized as a central player in epigenetic regulation⁴³, genome stability^44,45, and meiotic progression^46,47, the biophysical principles governing their spatiotemporal organization have remained elusive. In this study, we establish a causal link between MORC2 condensation and its transcriptional regulatory function. By integrating high-resolution structural insights with functional assays, we reveal that MORC2 does not act merely as a solitary chromatin remodeler^3,5,48; instead, it functions as phase separation-driven DNA compaction machinery. This discovery demonstrates how mesoscale material properties—specifically the fluidity and density of protein-DNA assemblies—are directly translated into precise gene regulation.

A Modular Engine for Condensate Assembly

The architectural basis of MORC2 assembly relies on a finely tuned multi-domain interplay. The C-terminal region, comprising the structured CC3-IBD and the intrinsically disordered IDR, provides the mechanistic engine for this process. CC3 serves as a critical structural scaffold (Figure 1e, and 3a), mediating homodimerization that effectively doubles the molecular valency, thereby facilitating multivalent “sticker” interactions between the IDR and the defined IBD (Figure 4). We found that mutations selectively abrogating condensate formation, or those that arrest condensate dynamics without blocking assembly, impair MORC2’s ability to repress core target genes (Figure 6). This underscores that high internal fluidity is functionally paramount, ensuring the rapid exchange kinetics required for continuous ATP hydrolysis and dynamic chromatin engagement.

The “Viscoelastic Window” and Molecular Stoichiometry

A defining insight from our study is that the mere presence of nuclear puncta is insufficient for MORC2’s repressor activity. Instead, its function is governed by the precise material state of the condensates. We demonstrate that DNA acts as an essential functional scaffold (Figure 5), driving condensation potentially mediated by nonspecific electrostatic interactions with the arginine/lysine-rich IDR. Crucially, these assemblies are exquisitely sensitive to molecular stoichiometry. This establishes a narrow, functionally permissive “viscoelastic window”: Too Diffuse: MORC2 fails to achieve the local density necessary for mechanical DNA compaction. Too Rigid: If the system crosses the threshold into a solid-like, arrested state (as seen in the MORC2 +KS mutant or E236G variant), it loses the fluidity required for enzymatic turnover and co-repressor engagement. Within the nucleus, the CW domain likely serves as a regulatory node, orchestrating the compositional regulation of these condensates by reading specific epigenetic marks to fine-tune local valency and morphology (Figure 3 and Figure S3).

Mechanistic Insights into Pathogenic Variants

Our analysis of disease-associated variants—particularly E236G (CMT2Z) and T424R (SMA)—highlights how mutations in the ATPase domain can pleiotropically affect both enzymatic turnover and phase behavior (Figure 7). While T424R primarily accelerates the ATPase dimer cycle¹³, E236G triggers a transition toward a less dynamic, “hardened” condensate state. These results emphasize that disease mutations do not simply “turn off” the protein; they perturb the biophysical context of the assembly. The observation that some variants exhibit elevated ATPase activity in vitro suggests that in the native cellular environment, condensation-mediated local enrichment may be required to properly couple ATP hydrolysis to productive chromatin remodeling.

Our findings clarify previous discrepancies regarding MORC2’s DNA binding. By utilizing the 601 nucleosome-positioning sequence, we demonstrate a much more robust interaction than previously captured with short, random dsDNA. Furthermore, the use of full-length MORC2 reveals that C-terminal regulatory domains are essential for maintaining enzymatic homeostasis—a feature masked in truncated constructs. Whether MORC2-mediated silencing is achieved through direct modulation of ATPase cycles or an independent mechanical compaction mechanism remains an open question.

While our study establishes the essential role of MORC2 condensation in transcriptional control, this exploration represents an initial step toward deciphering the complex coordination between enzymatic turnover and phase behavior. Future research is required to resolve how the ATP-dependent “DNA clamp” cycle is spatially coupled to the physical compaction of chromatin, and whether additional co-repressors are selectively partitioned into these dynamic assemblies to maintain epigenetic stability.

Materials and Methods

Plasmid construction, cell culture and transfection

WT MORC2 (UniProt: Q9Y6X9) cDNA was amplified from a cDNA library derived from HeLa cells via reverse transcription. The MORC2 cDNA was then subcloned into the p3xFLAG-Myc-CMV-24 vector between BglII and KpnI restriction sites using seamless cloning. The seamless cloning process was facilitated by the ClonExpress II One-Step Cloning Kit (Vazyme, #C112-02) according to the manufacturer’s instructions. An EGFP sequence was subsequently inserted into the BglII site at the N-terminal of MORC2 to generate the EGFP-MORC2 construct. To generate deletion mutant constructs of MORC2 for intracellular studies, specific regions of MORC2 were targeted and deleted using a polymerase chain reaction (PCR)-based approach. An 18– or 21 bp homologous sequence originating from one of the forward or reverse primers was introduced into the other primer, and the resulting PCR products were subjected to homologous recombination cloning, facilitated by the 18-bp homologous sequence. For protein expression and purification, MORC2FL and truncated MORC2 were cloned into prokaryotic expression vectors with MBP or Trx fusion proteins for bacterial expression.

In addition, MORC2FL and a series of truncation mutants were purified using a HEK293F eukaryotic expression system for in vitro phase separation and static light scattering (SLS) assays. A260/280 measurements, agarose gel analysis, or EMSA information on protein purifications and their experimental applications in this study is provided in Supplementary Table 2. HeLa, HEK293T cells were cultured in DMEM medium (Biological Industries, DMEM, high re maintained in a humidified incubator at 37°C with 5% CO₂. Cell transfection was performed using Lipofectamine 3000 (Invitrogen) according to the manufacturer’s instructions. Briefly, cells were seeded in appropriate culture plates the day before transfection and allowed to reach 40-60% confluency. The separately diluted Lipofectamine 3000 reagent and plasmids in Opti-MEM reduced serum medium (Gibco) were mixed, and the transfection complexes were added to cells after 20 minutes. After a 6-hour incubation, the transfection medium was replaced with fresh complete DMEM medium.

Recombinant protein expression, purification

Plasmids containing His-tagged genes were transformed into E. coli BL21 (DE3) cells and cultured in Luria-Bertani medium with 50 mg/L ampicillin. When the culture reached an optical density (OD) of 0.6-0.8 at 600 nm, the temperature was lowered to 16°C. Isopropyl β-D-1-thiogalactopyranoside (IPTG) (0.25 mM) was added to induce protein expression for 18-20 hours. Cells were harvested by centrifugation at 13,000 × g for 15 minutes, then resuspended in Buffer A (50 mM Tris-HCl, pH 8.0, 1000 mM NaCl, 10 mM imidazole, 1 mM dithiothreitol (DTT), and 1 mM phenylmethylsulfonyl fluoride (PMSF)). The cell suspension was snap-frozen and stored at –80°C for subsequent protein purification. HEK293F suspension-adapted cells were cultured in conical flasks at 37°C, 120 rpm, and 5% CO₂, until a density of 1.0 × 10[ cells/mL was reached. The cells were transiently transfected with 1 mg of MORC2-6 × His plasmid using 2 mL of branched polyethyleneimine (1 mg/mL). After 48 hours of incubation, cells were harvested by centrifugation at 3,000 × g for 5 minutes, resuspended in Buffer A, snap-frozen, and stored at –80 °C for further purification.

All purification steps were performed at 4°C to preserve protein activity. Frozen cell suspensions were rapidly thawed in water, lysed by sonication (200 W for ∼0.3 hours), and centrifuged at 20,000 × g for 30 minutes at 4°C. The supernatant was filtered through a 0.22 μm syringe filter to obtain soluble material. A His-Trap Ni-NTA 6FF chromatography column was equilibrated with five column volumes of Buffer A (Buffer A: 50 mM Tris-HCl, pH 8.0, 1000 mM NaCl). The clarified extract was loaded onto the column, washed with Buffer A to remove non-specific proteins, and eluted with a linear gradient of nickel elution buffer (Buffer B: 50 mM Tris-HCl, pH 8.0, 1 M NaCl, 300 mM imidazole, 1 mM DTT). Eluted fractions were analyzed by SDS-PAGE and Coomassie staining. The protein of interest was concentrated, exchanged into Buffer C (50 mM Tris-HCl, pH 8.0, 1000 mM NaCl, 1 mM EDTA, 1 mM DTT) using an Amicon Ultra centrifugal filter unit, following overnight treatment with Benzonase nuclease to eliminate nucleic acid contamination or HRV 3C Protease to cut His-tag. The purification process was subsequently refined through gel filtration using a Superdex 6 Increase column (GE Healthcare) for the MORC2FL. Following this, fragments were conjugated with a GPGS linker, and the resulting mixture was subjected to a size-exclusion chromatography process (Superdex 200/75 column, GE Healthcare) to yield a final solution comprising 50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 1 mM EDTA, and 1 mM DTT. Peak fractions were analyzed by SDS-PAGE and Coomassie staining, confirming the successful purification of highly homogeneous MORC2FL (1-1032) and various fragments (Figure S7), then flash-frozen in liquid nitrogen and stored at –80°C.

Crystallization, data collection and processing

The CC3 domain (901-1003) and various mutations of MORC2constructs were cloned into a modified pET32M vector (with Trx-tag) using standard PCR-based methods and confirmed by DNA sequencing. Due to the lack of homologous structures, protein crystals with heavy atom derivatives are required to determine their phases. Therefore, we used the methionine-deficient strain B834 to express the target protein with selenomethionine (SeMet) in an inorganic selenosubstituted medium. Recombinant proteins were expressed in E. coli B834 cells in M9 medium at 16°C. MORC2 CC3 fused with a GPGS linker were finally changed by step of size-exclusion chromatography (Superdex 75 increase column, GE Healthcare) into 50 mM Tris-HCl, pH 8.0, 100 mM NaCl, 1 mM EDTA and 1 mM DTT). The freshly purified protein was concentrated to 10∼20 mg/mL. Crystals were obtained by the static sitting drop method in 0.1M Citric acid (pH 3.5) and 25% w/v PEG 3350 at 25°C. Glycerol (20%) was added as the cryo-protectant. A 3.1-Å resolution X-ray dataset was collected at the beamline BL18U1 of the Shanghai Synchrotron Radiation Facility (SSRF). The diffraction data were processed and scaled by HKL2000 (http://www.hkl-xray.com/). Using the structure of auto-build model by SAD data and AlpahFold2 predicted model as the search model, the initial structural model was solved by molecular replacement in PHASER (https://www.ccp4.ac.uk/). Further manual model adjustment and refinement were completed iteratively using COOT and PHENIX. The final structure was validated by PISA and MolProbity (https://www.phenix-online.org/). The final refinement statistics of the complex structure are listed in Supplementary Table 1. All structure figures were prepared using the programme PyMOL (http://pymol.sourceforge.net/).

Protein labeling with fluorophore

The Cy3 NHS ester (AAT Bioquest) fluorophores were dissolved in DMSO at 10 mg/mL. Purified proteins were prepared in reaction buffer at a concentration of 10 mg/mL. The fluorophores were added to the protein solution at a 1:1 molar ratio. The reaction was conducted with shaking at room temperature for 1 hour in the dark. After the reaction, labeling was quenched with 200 mM Tris-HCl, pH 8.3, and the labeled proteins were desalted using a HiTrap desalting column (GE Healthcare). Fluorescence labeling efficiency was assessed using a Nanodrop 2000 spectrophotometer (ThermoFisher #ND-2000). The final labeling efficiency of each protein was adjusted to 1% by mixing the labeled protein with an excess of unlabeled protein (1:99 ratio).

Lentiviral mediated H2A-mCherry inducible expression in HeLa cells

Stable expression of H2A-mCherry in HeLa cells was achieved through lentiviral-mediated transduction. First, H2A-mCherry gene sequence was inserted into the inducible lentiviral vector pTRIPZ to generate the expression plasmid pTRIPZ-H2A-mCherry. Lentiviral particles were generated by triple transfection of HEK293T cells with pTRIPZ-H2A-mCherry, PMD2.G, and PSPAX2 plasmids using Lipofectamine 3000. The viral supernatant was collected 48 hours after transfection, filtered with 0.45-μm filter to remove cell debris, and used to transduce HeLa cells. Following transduction, HeLa cells were selected with 50 µg/mL hygromycin to establish stable cell lines. H2A-mCherry expression was induced with 1 μg/mL doxycycline.

Live cell imaging and immunofluorescence staining

For live-cell imaging, HeLa cells were seeded onto 35 mm culture dishes with glass-bottom plates (Biosharp) and transfected with EGFP-MORC2 or its mutant constructs. Images were captured 12 hours post-transfection using a Nikon confocal microscope equipped with an incubator system (37°C, 5% CO₂). Nuclei were stained with Hoechst 33342 or visualized using mCherry-labeled H2A fusion protein. For immunofluorescence, cells were fixed with 4% paraformaldehyde, permeabilized with 0.1% Triton X-100, and blocked with 10% goat serum. Primary antibodies were incubated overnight at 4°C, followed by incubation with fluorescence-conjugated secondary antibodies for 2 hours at room temperature in the dark. Fluorescence images were acquired using a Nikon confocal microscope.

CRISPR-Cas9 mediated MORC2 knockout cells

MORC2 knockout (KO) HeLa cell lines were generated using the CRISPR-Cas9 system. A CRISPR single-guide RNA (sgRNA) targeting exon 5 of MORC2 (5′-ACACCTGAGTCTACTCAGAT-3′) was designed using the online tool (http://crispor.tefor.net/). Two DNA oligos with complementary sgRNA targeting sequence and additional bases were designed as previously described^49,50. These oligos were annealed and cloned into pLentiCRISPR v2 (Addgene plasmid #52961) to generate the MORC2 knockout vector. After transfection and puromycin selection, MORC2-deleted cells were obtained. Individual cells were sorted into separate wells of a culture plate to establish monoclonal cell lines using flow cytometry. To validate the knockout cell lines, genomic DNA was extracted from both WT and KO cells. The specific regions targeted by the sgRNA were amplified using PCR with primers: 5′-CAACATTCTCGAGCTGGACCTACAG-3′ (forward) and 5′-CCACGACAAGACTGGAAACGTGACTC-3′ (reverse). The PCR products were purified and subjected to Sanger sequencing, revealing the deletion and insertion mutations in the target sites of the MORC2 allelic genes (Figure S2d).

Generation of endogenous EGFP-MORC2 chimeric mice

CRISPR-Cas9 was used to generate EGFP-MORC2 chimeric mice. The EGFP and linker sequence were inserted at the N-terminus of the mouse MORC2a gene. Briefly, a single-guide RNA (sgRNA) was designed to target exon 1 of MORC2a, with the sequence GACTGAAGACTCATTGCTGTCA. A donor DNA fragments containing the EGFP-linker region flanked by homologous arms was constructed. Fertilized mouse zygotes were collected, and a mixture of Cas9 mRNA (50 ng/μL), sgRNA (25 ng/μL), and donor DNA (50 ng/μL) was microinjected into pronucleus of the zygotes. The injected embryos were transferred into the oviducts of pseudopregnant recipient female mice. Resulting pups were genotyped by PCR and Sanger sequencing to confirm transgene integration (Figure S2f). The genotyping primers for the EGFP-MORC2 allele were as follows: Left-F1: GCGGTGGTTGAGTTCCAATTCC; Left-R1: CTTCAGGGTCAGCTTGCCGTAG; Right-F1: GAACGGCATCAAGGTGAACTTC; Right-R1: TGTATCCAAGGTAATCTCTGTGGTA.

Mice were bred at the Animal Center of the University of Science and Technology of China (USTC). Animals were housed under controlled conditions (22[°C, 12-hour light/dark cycle) with free access to food and water ad libitum. All experimental procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Science and Technology of China (Hefei, China).

Fluorescence recovery after photobleaching (FRAP)

The FRAP assay was performed to investigate the dynamics of protein mobility within condensates, either in in vivo or in vitro. Specific regions of interest (ROIs) approximately 2 μm² in size were selected and subjected to photobleaching using 100% laser power (488 nm) for approximately 20 ns. One droplet was left unbleached as a control. Fluorescence recovery within the ROI was monitored for 100-300 seconds. FRAP was performed using a 63x oil immersion objective mounted on a Nikon confocal microscope (in cells) or a Zeiss LSM 880 (in vitro). Fluorescence intensity was normalized to the pre-bleach signal to analyze the recovery kinetics. For brain slice FRAP, organotypic brain slices were prepared from adult chimeric mice expressing endogenous EGFP-MORC2, following a slightly modified protocol previously described⁴⁹. After decapitation, brains were rapidly extracted and placed in ice-cold dissection medium consisting of Hibernate A, 2% B27 supplement, 2[mM L-glutamine, and 1% penicillin–streptomycin. The cerebellum and midbrain were removed, and the remaining cerebral hemispheres were coronally sectioned at 250[μm using a McIlwain tissue chopper. Slices were gently separated in chilled dissection medium and transferred onto glass-bottom dishes containing culture medium composed of Neurobasal A, 2% B27 supplement, 2[mM L-glutamine, and 1% penicillin–streptomycin. Imaging was initiated immediately after brain slice preparation. FRAP experiments were performed using the Nikon confocal microscope. Photobleaching was carried out with 5-7 laser pulses (50[μs dwell time), and images were acquired at 2-s intervals. Post-bleach images were acquired 20[s after photobleaching.

Sedimentation assay

To assess concentration-dependent partitioning behavior, purified proteins at the indicated concentrations were incubated under the same buffer conditions used for in vitro phase separation assays and equilibrated for 10 min at room temperature. Samples were then centrifuged at 16,000 × g for 10 min to separate the supernatant and pellet fractions. Following centrifugation, the supernatant was carefully collected, and the pellet was resuspended in an equal volume of buffer. Equal volumes of supernatant and pellet fractions were analyzed by SDS–PAGE and quantified by densitometry.

Electrophoretic mobility shift assays (EMSA)

Widom 601 DNA (147 bp, 25 nM) was amplified by PCR and incubated with increasing concentrations of MORC2 or its domain constructs in binding buffer (20 mM Tris-HCl, pH 7.5, 45 mM KCl, 4 mM MgCl₂, 5% glycerol, 5 mM DTT). The reaction mixture was loaded onto a 6% polyacrylamide gel and electrophoresed at 120 V for 70 minutes. DNA was visualized using SYBR Gold stain, and images were captured using the ChemiDoc MP Imaging System.

Size-exclusion chromatography coupled with static light scattering

The analysis was performed using an AKTA FPLC system (GE Healthcare) coupled to a static light scattering detector (miniDawn, Wyatt) and a differential refractive index detector (Optilab, Wyatt). Protein samples (70[μM) were filtered and applied to a Superdex 200 increase or Superose 6 increase column, pre-equilibrated with a column buffer containing 50[mM Tris-HCl, pH 8.0, 150[mM NaCl or 1000[mM NaCl (MORC2FL), 1[mM EDTA, and 1[mM DTT. Data were analyzed using ASTRA6 software (Wyatt).

In vitro phase separation and fluorescence microscopy imaging

Droplet formation was monitored using differential interference contrast (DIC) and fluorescence microscopy. To induce phase separation, protein was added to phase separation buffer (50 mM Tris-HCl, pH 8.0, 150[mM NaCl, 1[mM DTT). For consistency, all MORC2 samples were diluted to a final concentration of 4[μM with phase separation buffer containing 200[mM NaCl, unless otherwise specified. All preparation steps were performed at 4[°C to preserve protein activity. The phase separation mixtures were then equilibrated at room temperature for 20[min. A 5-10[μL aliquot of each sample was placed onto a custom-made glass slide, covered with an 18[mm diameter coverslip, and incubated at room temperature for 20[min before imaging.

Protein isotope labeling and NMR Spectroscopy

Isotopically labeled protein samples were expressed in minimal M9/H₂O or M9/D₂O medium. Uniformly ¹⁵N-labeled (or ¹³C– or ¹⁵N-labeled) protein samples were prepared in M9/H₂O medium supplemented with 1[g/L of ¹⁵NH₄Cl and 2[g/L of glucose (or ¹³C-glucose). The cells were harvested at an OD₆₀₀ of approximately 1.0. Isotope-labeled precursors and amino acids were purchased from Cambridge Isotope Laboratories (CIL). Isotopically labeled samples were prepared in 50[mM Tris-HCl, pH 8.0, 100[mM NaCl, and 7% D[O. Protein concentrations ranged from 0.5 to 1.0[mM. NMR experiments were performed using Bruker Avance III 600[MHz spectrometers equipped with a cryo-probe. NMR spectra of MORC2 and its complex were recorded at 298[K. Backbone assignments of MORC2^1004-1032 were obtained using SOFAST versions of standard triple-resonance experiments, including HNCACB and CBCACONH, recorded at 298 K. Residual assignment ambiguities were resolved using ¹⁵N-edited HMQC-NOESY-HMQC spectra. All NMR spectra were processed using NMRPipe and analyzed with NMRViewJ software (http://www.onemoonscientific.com). CSPs were calculated in Hz at 600 MHz using the following equation:

Fluorescence polarization Assay (FP)

Fluorescence polarization (FP) assays were performed at room temperature (25[°C) using a BioTek H1 fluorescence spectrophotometer to evaluate the binding affinity between MORC2 protein and 5-carboxyfluorescein (5-FAM)-labeled 601 DNA (FAM-601) across a concentration range of 0-2000[nM. The 147-bp Widom 601 DNA fragment was generated via PCR amplification using a 5-FAM-labeled primer. The labeled primer was dissolved in 0.1[M NaHCO[ buffer, pH 8.3, and the amplified product was purified by agarose gel extraction to eliminate residual primers and nonspecific amplification products.

Fluorescence measurements were carried out in assay buffer (50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1 mM EDTA, 1 mM DTT, and 10% glycerol) containing purified FAM-601 DNA at a final concentration of 0.4[ng/μL. Excitation and emission wavelengths were optimized according to the instrument manual, with excitation set at 485[nm and emission at 525[nm. Fluorescence polarization values were recorded in triplicate, and control measurements were included to correct for background signals.

Data were analyzed to determine the equilibrium dissociation constant (K_d) for the MORC2-FAM-601 interaction by fitting data to a log(agonist) vs. response model. In parallel, the effect of small molecules on this binding interaction was assessed by introducing compounds into the assay mixture and monitoring changes in fluorescence polarization.

ATPase activity measurements

ATPase activity was measured⁵² using 1[µM MORC2 or its mutant variants at 37°C in 50[mM Tris-HCl, pH 8.0, 150[mM NaCl, 8[mM MgCl₂, 10% glycerol and 5[mM ATP. All experiments were repeated independently at least three times.

RNA-seq and bioinformatic analysis

MORC2-knockout HeLa cells were seeded at equal density and transfected with 3[µg of plasmid DNA and 12[µg of PEI (polyethylenimine) per dish. After 6[hours, the transfection medium was replaced with fresh complete medium, and cells were incubated for an additional 48[hours. Cells were then washed twice with ice-cold PBS and lysed directly in RNA Isolator (R401, Vazyme) on ice. Total RNA was extracted using the RNeasy Mini Kit (Qiagen, 74004) according to the manufacturer’s protocol. RNA quality was assessed using NanoDrop 2000 (Thermo Fisher Scientific), Agilent 2100 Bioanalyzer, and LabChip GX (PerkinElmer). For library preparation, 1[µg of total RNA was used per sample, employing the VAHTS Universal V6 RNA-seq Library Prep Kit for MGI®. The resulting double-stranded DNA libraries were denatured, circularized, and digested to obtain single-stranded circular DNA. DNA nanoballs (DNBs) were generated from these templates through rolling circle amplification (RCA). Sequencing was performed on the MGI DNBSEQ-T7 platform (MGI Tech Co., Ltd) using combinatorial Probe-Anchor Synthesis (cPAS) chemistry.

Before performing differential expression analysis, lowly expressed genes were filtered out by retaining only those with a raw count of ≥10 in at least three samples (baseMean). Differential gene expression analysis between sample groups was carried out using the DESeq2 package (v1.26.0). Genes with p-adjust < 0.05 were defined as differentially expressed genes (DEGs). These DEGs were subsequently subjected to Gene Ontology (GO) enrichment analysis in DAVID (Knowledgebase v2025_1).

Statistical Analysis

All statistical analyses were performed in GraphPad Prism. All data underwent normality testing (D’Agostino & Pearson test and Anderson-Darling test) in advance. Two-tailed unpaired Student’s t-test post-hoc Welch’s correction and one-way ANOVA with Tukey’s post hoc test was used when data were distributed normally. The significance level was set as p < 0.05. All statistical results are provided as Source Data.

Supplemental Tables and Figures

Table S1

Table S2.

Figure S1.

Figure S2.

Figure S3.

Figure S4.

Figure S5.

Figure S6.

Figure S7.

Figure S8.

Figure S9.

Figure S10.

Figure S11.

Figure S12.

Data availability

All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. Raw gene expression data are available in Source Data. The atomic coordinates of the MORC2 CC3 domain have been deposited to the Protein Data Bank under the accession code: 9KQL.

Acknowledgements

We thank Dr. Xuebiao Yao (University of Science and Technology of China, USTC), Dr. Zhi Qi and Dr. Linyu Zuo (Peking University) for helpful discussion. We thank members of the Shanghai Synchrotron Radiation Facility (SSRF, China; beamlines BL02U1, BL18U1, and BL19U1) for X-ray beam time. This work was supported by grants from the National Key R&D Program of China (2019YFA0508402), the National Natural Science Foundation of China (22122703, 32470808, 32170767, 32471273, 31971144, and T2221005), the Center for Advanced Interdisciplinary Science and Biomedicine of IHM (QYPY20220014), the Strategic Priority Research Program of the Chinese Academy of Sciences (XDB0490000), the Major Frontier Research Project of USTC (LS9100000002), USTC Research Funds of Double First-Class Initiative (YD9100002507 and YD9100002054), the Anhui Medical University Foundation (9101224201), and National Natural Science Foundation Incubation Program of The Second Affiliated Hospital of Anhui Medical University (2021GQFY02).

Additional information

Author contributions

Y.Z., W.X., W.D., C.H., C.W., and Y.B. designed the experiments. Y.B., Y.Z., W.X., and W.D. conducted the experiments. Y.Z. determined the crystal structure. Y.W., W.J. and F.Z. assisted with the experiments and reagents. All authors analyzed the data. Y.Z., Y.B., C.H., and C.W. wrote the manuscript with input from all authors. All authors approved the final version of the manuscript. C.H., and C.W supervised the project.

Funding

MOST | National Natural Science Foundation of China (NSFC) (22122703)

• Weiya Xu

MOST | National Natural Science Foundation of China (NSFC) (32470808)

• Weiya Xu

MOST | National Natural Science Foundation of China (NSFC) (32170767)

• Weiya Xu

MOST | National Natural Science Foundation of China (NSFC) (32471273)

• Weiya Xu

MOST | National Natural Science Foundation of China (NSFC) (31971144)

• Weiya Xu

MOST | National Natural Science Foundation of China (NSFC) (T2221005)

• Weiya Xu

Anhui Medical University Foundation (9101224201)

• Yihui Bi

MOST | National Key Research and Development Program of China (NKPs) (2019YFA0508402)

• Chengdong Huang

Additional files

PDB X-ray Structure Validation Report