TCL1A mediates DNA methylation defects in recurrent hydatidiform mole with NLRP7 pathogenic variants

Domain: chemistry
Published: 2026-03-05
Editorial outcome: minor_revision

Abstract
Pathogenic variants in NLRP7, implicated in 55% of recurrent hydatidiform mole characterized by hypomethylation at maternally methylated imprinted regions, are proposed to disrupt de novo DNA methylation in human oocytes. However, the precise mechanism remains unclear. Here, we identify TCL1A, a DNMT3A inhibitor, as an endogenous NLRP7-interacting partner. The cryo-EM structure of the NLRP7-TCL1A complex reveals its fundamental architecture. Comprehensive analysis demonstrates that the majority of recurrent hydatidiform mole-causing NLRP7 variants impair its interaction with TCL1A. Mechanistically, NLRP7 potentially safeguards oocyte methylome by sequestering TCL1A in the cytoplasm, thereby preventing its nuclear entry and subsequent suppression of DNMT3A-mediated de novo methylation. Combining in silico predictions and interaction analysis, we identify L766R as a pathogenic variant. These findings propose a cytoplasmic regulatory mechanism governing nuclear DNA methylation, explaining the hypomethylation pathogenesis in NLRP7 variant-associated recurrent hydatidiform mole.

Introduction
Hydatidiform mole (HM) is a severe reproductive disease characterized by aberrant embryonic development and trophoblast hyperproliferation1–3. Women with a history of molar pregnancy are at an increased risk of recurrent HM (RHM) and potential progression to malignant gestational trophoblastic neoplasia3,4. As an imprinting disorder, HM is closely related to defective DNA methylation in imprinted germline differentially methylated regions (gDMRs), which primarily results from the loss of the maternal genome, an excess of the paternal genome, or abnormal establishment or maintenance of maternal imprinting5–8. Notably, biallelic pathogenic variants in components of the human subcortical maternal complex (hSCMC), a conserved cytoplasmic complex essential for oocyte and early embryo development, have been implicated in these imprinting disorders1,9–12. The hSCMC consists of four proteins (NLRP5, OOEP, TLE6, and KHDC3L) and three putative proteins (NLRP2, PADI6, and human-specific NLRP7)13–15. Pathogenic variants in NLRP7 and KHDC3L account for the majority of RHM cases, while those in PADI6 and NLRP5 lead to a spectrum of reproductive disorders, including RHM, multilocus imprinting disturbances (MLID), and early embryonic arrest1,9–12.

NLRP7 is a human maternal-effect gene identified from familial RHM (FRHM) and accounts for 55% of patients with RHM1,9. Unlike the most androgenetic HM, NLRP7 variant-related RHMs are typically normal diploid biparental1,9,16. Investigations of molar tissues from FRHM patients with biallelic NLRP7 pathogenic variants have revealed widespread loss of methylation at maternally imprinted gDMRs, while paternally imprinted gDMRs remain relatively unaffected17,18. Furthermore, although the DNA methylation landscape in oocytes carrying NLRP7 pathogenic variants remains to be determined, whole-genome bisulfite sequencing (WGBS) analysis of the oocytes from a patient with biallelic KHDC3L variants, which account for 5% patients with RHM, revealed genome-wide hypomethylation19. These findings suggest that pathogenic variants in NLRP7/KHDC3L disrupt de novo DNA methylation during human oogenesis. Interestingly, heterozygous variants in NLRP7 have also been observed in mothers of offspring with MLID20–22. However, the mechanism by which cytoplasmically localized NLRP7 influences nuclear DNA methylation remains unclear23,24.

The lack of relevant murine models and scarce human materials hinder the recapitulation of NLRP7 physiological functions in vivo. The identification of its interacting proteins has predominantly relied on in vitro cell systems25–28. Although there may be significant differences in mechanisms and patterns between humans and mice, mouse maternal factors essential for de novo methylation in oocytes, including DNMT3A, DNMT3L, KDM1A, KDM1B, SETD2, HDAC1/HDAC2, SIN3A, CFP1, SALL4, and HIRA, have been identified29,30. Knockout of these factors results in hypomethylation across the entire genome and at maternal gDMRs in oocytes or embryos, similar to that in some patients carrying pathogenic NLRP7/KHDC3L variants17,19. To date, no experimental evidence has confirmed the direct interaction between NLRP7 and these epigenetic factors. In addition, NLRP7 variants are associated with significant phenotypic heterogeneity, manifesting as a spectrum of reproductive and developmental disorders, such as early embryonic arrest, recurrent miscarriage, MLID, and, in rare cases, live births20,21,31–33. This phenotypic variability complicates the genetic diagnosis of NLRP7 pathogenic variants. Therefore, the identification of the in vivo binding partners is critical for understanding the functional mechanism and accurately evaluating the pathogenicity of hSCMC/NLRP7 variants, particularly in cases involving sporadic missense variants.

Human T-cell leukemia/lymphoma 1A (TCL1A), an oncoprotein of the TCL1 family, drives T-cell leukemias and lymphomas by dysregulating the AKT pathway34,35. Notably, nuclear TCL1A has been reported to disrupt de novo DNA methylation by inhibiting methyltransferase 3A/3B (DNMT3A/3B) in B-cell chronic lymphocytic leukemia36. Interestingly, TCL1A is also highly expressed in human oocytes37. However, its function and physiological significance remain elusive.

In this study, we screen the binding partners of hSCMC in vivo and in vitro, leading to the identification of a direct interaction between TCL1A and the human-specific NLRP7. Most significantly, our findings reveal that the impaired NLRP7-TCL1A binding represents a potential pathogenic mechanism underlying the NLRP7 variants. This defective interaction facilitates the translocation of hTCL1A into the nucleus, where it likely binds to and inhibits DNMT3A methylation activity, ultimately leading to DNA methylation defects in the RHM associated with NLRP7 pathogenic variants. In addition, the evidence we provide suggests that the NLRP7-TCL1A interaction is a validated molecular indicator to assess the pathogenicity of NLRP7 variants in clinical practice.

Results
Maternal deficiency of Nlrp5 or Khdc3 has limited effects on the establishment of genomic DNA methylome in mouse oocytes
We first investigated whether mouse models could recapitulate the defects of de novo DNA methylation establishment observed in human oocytes carrying pathogenic variants in components of the hSCMC. Nlrp5 (encoding one of SCMC core components) and Khdc3 (the ortholog of human KHDC3L) knockout mouse lines were used as experimental models38,39. The genomic DNA methylome in MII-stage oocytes from normal and maternally deficient mice was analyzed using STEM-seq (small-scale TELP-enabled methylome sequencing)40. Compared with the controls, DNA methylation levels in SCMC-defective oocytes only showed minor changes both at the global level and at maternal imprinting control regions (ICRs) (Supplementary Fig. 1a–e). These findings suggest that current SCMC-deficient mouse models inadequately represent the mechanisms of aberrant de novo DNA methylation establishment in the oocytes from patients with pathogenic variants in KHDC3L or NLRP7, highlighting the necessity for human-derived models in such studies.

TCL1A is integrated into the hSCMC through interacting human-specific NLRP7
To identify the endogenous interactors linking hSCMC to DNA methylation, we performed Co-immunoprecipitation (Co-IP) and mass spectrometry (MS) assays for multiple components (NLRP7, KHDC3L, and NLRP5) using human abnormal oocytes and arrested cleavage-stage embryos discarded from assisted reproduction (Fig. 1a). Five interactomes were conducted: two for NLRP7 or KHDC3L and one for NLRP5. Proteins were filtered using the criteria of ≥2 unique peptides and ≥1.5-fold enrichment. However, the duplicate KHDC3L Co-IP-MS analyses yielded poor results, with only a limited number of proteins identified. Thus, we focused the analysis on the interactomes of NLRP7 and NLRP5. Ten candidate proteins shared in three interactomes of NLRP7 and NLRP5 were identified, including four known hSCMC members (NLRP5, TLE6, KHDC3L, and OOEP), three putative components (NLRP2, NLRP7, and PADI6), and three other proteins (CCNI2, CDK5, and TCL1A) (Fig. 1a, and Supplementary Table 1).

Co-IP followed by western blotting analysis confirmed that in addition to the four known hSCMC subunits, NLRP7 and NLRP2 were likely the components of the hSCMC in vivo (Fig. 1b). However, PADI6 was not obviously detected. In addition, CCNI2, CDK5, and TCL1A were precipitated by the NLRP7 antibody, while the NLRP5 antibody only pulled down CDK5 and TCL1A, but not CCNI2 (Fig. 1b), suggesting more extensive interactions of NLRP7. To further screen the direct partners of NLRP7, systematic pairwise Co-IP assays were conducted in HEK293T cells overexpressing those candidates. Our Co-IP results were mostly consistent with prior studies (Supplementary Fig. 2a)31. Notably, our assays demonstrated that NLRP7 exhibited distinct interaction strengths with other components in vitro. Weaker interactions of NLRP7 with NLRP5 and PADI6 were observed, while its binding to CDK5 and KHDC3L was minimal. However, our assay showed the strong binding between NLRP7 and NLRP2, CCNI2, itself, TCL1A, and the core hSCMC component TLE6 (Fig. 1c). To further investigate the direct interaction between NLRP7 and TLE6 in living cells, we developed an assay termed VPICONE (Visual Protein-protein Interaction by Co-translocation into Outer Nuclear Envelope) (Supplementary Fig. 2b). In this assay, mCherry tagged an outer nuclear membrane (ONM) anchor (termed as mCDTK) was fused to the C-terminus of bait (bait-mCDTK), thereby recruiting GFP-tagged prey to the ONM for visualization (Supplementary Fig. 2b, c). As a validation, direct interaction between TLE6 and OOEP was successfully detected as their co-localization on the ONM in HEK293T cells (Supplementary Fig. 2d). The VPICONE assay revealed that co-expression of NLRP7-mCDTK with GFP-TLE6 induced robust co-localization on the ONM, but was absent in controls (Fig. 1d). The in vivo interaction between NLRP7 and TLE6 was also detected by PLA in the discarded human MI oocytes (Fig. 1e, f). Collectively, these results indicate that human NLRP7 is an authentic component of hSCMC, directly binding to the core protein TLE6.

Among these candidate interactors of NLRP7, CCNI2, a cyclin family protein, binds to CDK5, probably regulating cell cycle in cancer cells41, while TCL1A is linked to DNA methylation via DNMT3 inhibition and thus maybe associated with the hSCMC’s function in imprinting disorders36. Thus, we further screened the interactions between TCL1A and those components of the hSCMC through pairwise Co-IP assays in co-expressing HEK293T cells. Those assays showed TCL1A specifically binds to NLRP7 and itself, consistent with its dimeric formation (Fig. 1g)42,43. Additionally, TCL1A interacted with NLRP7 isoforms 2 (NP_996611.2) and 3 (NP_001120727.1), but not isoform 1 (NP_631915.2), in their overexpressing HEK293T or COS-7 cells, indicating that the residues 643-672 of NLRP7 are essential for TCL1A binding (Supplementary Fig. 2e–g). NLRP7-TCL1A interaction remained stable even after washing with high salt buffer (Supplementary Fig. 2h), underscoring the interaction robustness. A variant in TCL1A (residues 36-38: PLT-to-AAA mutation) that was reported to disrupt its dimerization did not dramatically affect its interaction with NLRP7 (Supplementary Fig. 2h)42. Although multiple NLRP family proteins are present in human oocytes44, in vitro Co-IP assays demonstrated that TCL1A specifically interacted with NLRP7, but not directly with other proteins. It also suggests the functional specialization of NLRP7 compared to its most homologous paralog NLRP2 (Supplementary Fig. 2i). The presence of minimal KHDC3L binding did not significantly affect the interaction between NLRP7 and TCL1A (Supplementary Fig. 2j). Notably, the application of the VPICONE assay could visualize the recruitment of GFP-TCL1A to the ONM by NLRP7-mCDTK (Fig. 1h). PLA analysis confirmed the in vivo interaction between NLRP7 and TCL1A in the subcortex and cytoplasm of discarded human oocytes and cleavage embryos (Fig. 1i, j). Taken together, these in vitro and in vivo experiments suggest that TCL1A is a specific and endogenous binding partner of human-specific NLRP7.

We further investigated how the NLRP7-TCL1A complex is incorporated into the core hSCMC45. Our results demonstrated that the core NLRP5 precipitates TCL1A only in the presence of NLRP7 and was also dependent on TLE6/OOEP but not KHDC3L (Fig. 1k). The VPICONE assay further demonstrated that TLE6-mCDTK recruited GFP-TCL1A to the ONM in an NLRP7-dependent manner, supporting the formation of a TLE6-NLRP7-TCL1A complex in cells (Fig. 1l). Altogether, these findings suggest that the NLRP7-TCL1A complex is involved in hSCMC through its direct binding to TLE6 (Fig. 1m).

Tcl1, the murine ortholog of TCL1A is a maternal-effect gene crucial for preimplantation embryonic development46. Mouse Tcl1 RNA remained dormant until germinal vesicle breakdown. The protein was undetectable in fully grown oocytes (FGO) (Supplementary Fig. 3a, c, d)47,48. In contrast, consistent with the high translational activity detected at the latest in FGO, human TCL1A was easily detected in the sample of only three human FGO, suggesting its high abundance (Supplementary Fig. 3b–d)37,49. Furthermore, we observed no interaction between exogenous mouse Tcl1 and mSCMC core TLE6 in mouse oocytes (Supplementary Fig. 3e). Unlike that of the well-defined SCMC components, the level of mouse Tcl1 was also unaffected by maternal NLRP5 knockout at the 2-cell stage (Supplementary Fig. 3f)50–54. These results indicate that the relationship between hSCMC and NLRP7-TCL1A is specialized in humans.

Overall structure of the NLRP7-TCL1A complex
Human NLRP7, akin to other NLRP family members such as NLRP5 and NLRP3, comprises three typical domains: an N-terminal pyrin domain (PYD), a central NACHT domain (NBD, HD1, WHD, and HD2 subdomains), and a C-terminal region composed of 14 canonical leucine-rich repeats (LRR1-14) (Fig. 2a). TCL1A is an all-beta protein consisting of an eight-stranded antiparallel beta-barrel (Fig. 2a)43. Building on our recent structures of the mouse and human SCMC core (NLRP5/TLE6/OOEP)45,55, we attempted to model an extended hSCMC complex containing NLRP7 and TCL1A. However, despite multiple attempts, we were unable to purify a complex containing all these five proteins, likely due to the low efficiency of co-transfection with multiple plasmids.

We therefore focused on elucidating the structure of the NLRP7-TCL1A complex. For this purpose, Flag-tagged full-length NLRP7 isoform 3 (residues 1-1037) and Strep-tagged full-length TCL1A (residues 1-114) were co-expressed in HEK293F cells, and the NLRP7-TCL1A complex was purified (Fig. 2b). Single-particle cryo-EM analysis yielded two maps of NLRP7-TCL1A at resolutions of 3.58 Å (state I) and 3.48 Å (state II) (Fig. 2c–f, Supplementary Figs. 4–5, Supplementary Table 2). Interestingly, the NLRP7-TCL1A complex in state I contains two NLRP7 and four TCL1A molecules, while state II contains only one NLRP7 and two TCL1A molecules (Fig. 2c–f). This structural observation suggests that NLRP7-TCL1A adopts a dynamic oligomerization equilibrium, similar to human and mouse SCMC core45,55. The structural comparison revealed no conformational changes between State I and State II (Supplementary Fig. 6a). Therefore, subsequent analyses focused on the State I complex.

The overall structure of the state I complex forms an approximately 135 Å × 113 Å heterohexamer (Fig. 2c). The two NLRP7 molecules (a and b) embrace each other via extensive interactions with a PYD domain inserted into the cavity between the two dimerized NACHT domains (Fig. 2c, d). NLRP7 molecule (a) adopts a relatively intact structure, with the exception of two disordered regions: a flexible linker between the PYD and NACHT domains (residues 94-114) and a flexible segment within LRR2 (residues 637-655) (Fig. 2c, d). In NLRP7 (b), only the NACHT and partial LRR domains were resolved (Fig. 2c, d). Due to the absence of density for the flexible linker to PYD in both NLRP7s, the identity of the observed PYD domain could not be unambiguously assigned. For clarity in description, we assigned it to the NLRP7 molecule (a) in the current model. Each LRR domain of NLRP7 interacts with one molecule of the TCL1A dimer (Fig. 2c, d). The homo-dimeric forms of NLRP7 and TCL1A in the complex were in agreement with the Co-IP results (Fig. 1c, f).

In the complex, two NLRP7 molecules adopt the nearly identical conformation with a RMSD at 0.85 Å over 653 Cα atoms (Supplementary Fig. 6b). In addition, the structural models between TCL1A (a) in this complex and TCL1A monomer (PDB: 1JSG) are indistinguishable, with a RMSD value at 0.78 Å (Supplementary Fig. 6c), suggesting that the binding of NLRP7 does not significantly alter the conformation of TCL1A.

Distinctive structural characteristics of human NLRP7
Further structural comparisons among NLRP7, NLRP3, and mouse NLRP5 suggest that NLRP7 has relatively low ATPase activity. By superimposing the NBD domain of NLRP7 with that of NLRP3 (PDB for inactive form: 7PZC and active form: 8EJ4)56,57, the putative ATP/ADP binding pocket of NLRP7 is different from that in NLRP3 (Supplementary Fig. 7a, b), but similar to that in mouse NLRP5 (PDB: 8H96) (Supplementary Fig. 7c). The NACHT domain of NLRP7 is more extensive than that of NLRP3 (PDB: 7PZC), showing the HD1 and WHD subdomains of NLRP7 shifted more outwardly (Supplementary Fig. 7a). Specifically, in the Walker A and sense 2 motifs, which are critical for ATP binding and hydrolysis, the distances from key residues K184, T185, and H468 of NLRP7 to ADP are much longer than those in NLRP3 (PDB: 7pzc) by superposition (Supplementary Fig. 7d, e). When comparing the NACHT domain of mouse NLRP5 (PDB: 8h96) with that of NLRP7, the conformations of the WHD and HD2 subdomains exhibit a relatively high similarity, except for the HD1 domain, which shows a notable deflection (Supplementary Fig. 7c)45,55. The ATP hydrolysis activity assay further supported the structural analysis (Supplementary Fig. 7f). These data indicate the unique regulations of the conformation in the reproduction-related NLR proteins.

Interestingly, NLRP7 dimerizes in a distinct pattern, which contrasts with the LRR-facilitated dimerization of mouse NLRP5 and the autoinhibitory oligomerization characteristic of NLRP355,56. According to the calculation from PISA, the dimer interface of NLRP7s exceeds 1000 Å2. The entire dimer interface of NLRP7s in the complex could be divided into four distinct interfaces in three main regions (Fig. 3a). The internal PYD (a) is stabilized by interaction between the NBD domain of NLRP7 (a) and the HD1 and HD2 domains of NLRP7 (b), forming the interface I and II. The amino acid residues D36, Q39, W66, N69, E76, N73, and E77 of PYD (a) form hydrogen bonds with R154/R156 and E322/E327 in NBD and HD1 domains of NLRP7 (b) (Fig. 3a, I and II). Additionally, F32, P33, N65, W66, and N69 of PYD (a) mediates hydrophobic interactions with F157 of NBD(b) (Fig. 3a, III). The two LRR domains of NLRP7s form a parallel stack, resulting in the C-terminal region of LRR of NLRP7 (a) directly contact the NACHT domain of NLRP7 (b) (Fig. 3a, IV). The Q914, R917, K973, and S974 of LRR (a) form the hydrogen bonds with E340, D148, and D144 of NACHT domain in NLRP7 (b) (Fig. 3a, IV). Nevertheless, several mutations of polar residues and even the deletion of the PYD domains failed to significantly disrupt the NLRP7 dimerization (Fig. 3b), indicating that the extensive hydrophobic interactions likely also contribute to NLRP7 dimerization.

TCL1A directly binds to the LRR domain of NLRP7
In the NLRP7-TCL1A complex, TCL1A directly interacts with four α-helices of LRR domains of NLRP7 (LRR3-6) (Fig. 3c). Key residues R693, S702, D722, A726, R746, T747, N758, K760 and E780 in NLRP7 directly form hydrogen bonds with the residues W19, M62, T65, L78, Y79, R85, S89, D113 and D114 of TCL1A (a) (Fig. 3c). Substitutions at these interface residues or deletions of the LRR4-5 motif in either protein seriously disrupted the complex stability, with the exceptions of S702A in NLRP7 and W19A/T65A in TCL1A (Fig. 3d, e). Notably, NLRP7 pathogenic variants at R693 and D722 residues linked to RHM and reproductive failure1,58, abolished its TCL1A binding, suggesting that the interaction mediates the pathogenicity of NLRP7 variants.

Sequence alignment of other human reproductive NLRPs, including NLRP2, NLRP4, NLRP5, NLRP7, NLRP9, and NLRP14, identified the residues uniquely conserved in NLRP7 that enable TCL1A binding (Supplementary Fig. 8a). Comparative analysis across mammalian species revealed the conservation in TCL1A-binding interface of NLRP7 (Homo sapiens vs. Macaca: 80%, vs. Pongo: 95%, vs. Sus: 56.7%, vs. Capra: 58.9%) and TCL1A itself (Supplementary Fig. 9b, c). Co-IP assays suggest that this interaction is primate-specific, but is absent in porcine homologs (Supplementary Fig. 9d).

The majority of disease-associated NLRP7 variants impair its interaction with TCL1A
As of 2023, the Infevers database (https://infevers.umai-montpellier.fr) documented that approximately 91 non-synonymous variants in NLRP7 are associated with reproductive disease (Supplementary Table. 3). Based on the impact on protein sequences, they were classified into 45 non-truncating variants (nTVs, including 44 missense variants and a deletion variant T185del) and 46 truncating variants (TVs) (Fig. 4a). Based on the associated human phenotypes, 45 nTVs were mainly categorized into RHM (marked in red, 29 cases including T185del), non-RHM (marked in blue, 10 cases), and miscarriage (marked in orange, 6 cases), and TVs were mainly linked to RHM (45 cases) and nRHM (1 case with E776Gfs*14) (Fig. 4a). Approximately 44.8% (13/29) of the RHM-related nTVs was structurally located within the TCL1A-binding interface within LRR3-6, including the hotspot pathogenic variants R693W/P/Q and L750V, as well as R701C, C704Y, P716L, A719V, R721W/Q, D722G, H735P, and C761Y (Fig. 4b). It further indicates that disruption of binding with TCL1A contributes to the RHM caused by pathogenic variants in NLRP7. We selected 52 of the total 91 variants for further investigation, encompassing all 45 nTVs and 7 TVs, five of which are the pathogenic variants outside the interface and closest to the C-terminus (Supplementary Table 3).

First, we evaluated the impact of these NLRP7 pathogenic variants on protein expression levels in HEK293T cells. Approximately 30.8% (16/52) of the pathogenic variants significantly reduced NLRP7 expression, potentially due to increased protein instability (Supplementary Fig. 9a–e). They were mainly clustered in the NACHT domain (T185del, F250L, K379N, G380R, R390H, L398R) and LRR domain (C668R, R693W, C704Y, P716A, A719V, R721W, D722G, I858Hfs11, G981Rfs19). These findings suggest that the NLRP7 instability resulting from these variants is one of the potential pathogenic mechanisms.

We next evaluated the interaction of NLRP7 pathogenic variants-TCL1A in HEK293T cells. Notably, TCL1A stabilized the expression of normal NLRP7 but had a minor effect on certain mutants when co-expressed (Supplementary Fig. 9d–g). To reduce variability of NLRP7 expression in the experiments, we employed a semiquantitative pull-down assay to quantify the binding affinity of TCL1A to NLRP7 (Supplementary Fig. 9h). Approximately 51.1% (23/45) of nTVs significantly disrupted the NLRP7-TCL1A interaction (Fig. 4c, e), further suggesting that the impaired binding serves as a key pathogenic mechanism. Among these pathogenic variants, about 87.0% (20/23) were localized to the LRR domains, the region with the highest mutational burden observed in patients (Fig. 4c, e)1. Notably, all seven TVs abolished NLRP7-TCL1A binding (Fig. 4d, e). The VPICONE assay confirmed the defective interaction for several RHM-linked variants (Fig. 4f).

When focusing on the major clinical phenotype of those RHM-associated pathogenic variants, ~65.5% (19/29) of nTVs and ~71.4% (25/35) of total tested variants impaired the NLRP7-TCL1A interaction in our assays (Fig. 4g). Given that all five TVs closest to the C-terminus disrupting the interaction (Fig. 4d, Supplementary Table. 3), we reasonably hypothesized that TVs occurring upstream of these sites disrupt TCL1A binding. When incorporating all 46 pathogenic TVs into our analysis, approximately 75.8% (69/91) of total disease-linked variants affected the NLRP7-TCL1A interaction. Notably, when considering only RHM-causing non-synonymous variants, this proportion rose to ~86.5% (64/74) (Fig. 4g). Interestingly, we also observed that multiple pathogenic variants disrupt NLRP7 dimerization (Fig. 4h). Above all, these data demonstrate that NLRP7 pathogenic variants associated with reproductive disorders, especially RHM, predominantly result in protein instability and/or impair TCL1A binding capacity.

TCL1A modulates the pathogenicity of NLRP7 variants, potentially through its aberrant nuclear accumulation and inhibition of DNMT3A-mediated methylation
TCL1A and NLRP7 were predominantly localized in the cytoplasm of human GV oocytes, likely with a minor fraction of TCL1A detectable in the nucleus (Fig. 5a, b). Interestingly, the nuclear TCL1A was progressively increased in cleavage-stage embryos (Fig. 5a, b). Given the robust NLRP7-TCL1A interaction and their high expression, we hypothesized that NLRP7 functions to sequester TCL1A in the cytoplasm of human oocytes and early embryos, probably similar to the observations in which the SCMC/PADI6/NLRP14 restricts the cytoplasmic localization of epigenetic regulators SPIN1/DNMT1/UHRF1 by their direct binding in mouse oocytes and early embryos54,59–61.

We examined whether NLRP7 influences the subcellular localization of TCL1A in HEK293T cells (Supplementary Fig. 10a). Consistent with prior reports36,62, GFP-TCL1A exhibited a diffuse distribution throughout the cells with distinct nuclear localization when co-expressed with mCherry or the non-binding NLRP7 isoform 1-mCherry. In contrast, co-expression of NLRP7 isoform 3, but not most disease-associated variants, markedly reduced TCL1A nuclear localization (Supplementary Fig. 10a, b). Furthermore, the proportion of cells exhibiting TCL1A nuclear localization was negatively correlated with the NLRP7/TCL1A expression ratio (Supplementary Fig. 10c–e). A similar pattern was observed in mouse oocytes microinjected with mRNAs encoding normal NLRP7, its mutants, and GFP-TCL1A (Fig. 5c, Supplementary Fig. 10f). These results suggest that NLRP7 prevents the nuclear localization of TCL1A in a dose-dependent manner.

Given the pivotal role of DNMT3A in mediating de novo DNA methylation in oocytes and the association between TCL1A and DNMT3A36,63, we further investigated how nuclear TCL1A functions in DNMT3A methylation activity through several in vitro assays. First, the Co-IP assays confirmed a direct physical interaction between TCL1A and DNMT3A/3B, but not with DNMT1 and DNMT3L (Fig. 5d). Second, in vitro methylation assays using purified DNMT3A and TCL1A demonstrated that TCL1A binding significantly inhibited DNMT3A enzymatic activity (Fig. 5e). Third, although DNMT3L is not expressed in human oocytes64, and the role of alternative protein DNMT3B3 in human oocytes remains controversial65,66, we found that overexpressing TCL1A reduced the interactions between DNMT3A and DNMT3B3 or DNMT3L in HEK293T cells (Fig. 5f). Notably, our simultaneous study revealed that TCL1A acted through competitive binding to DNMT3A, thereby inducing conformational rearrangements that consequently impair its interactions with DNA and S-adenosylmethionine67.

To further investigate the impact of NLRP7/TCL1A on the establishment of de novo DNA methylation in cells, we utilized a cellular differentiation system in which hypomethylated mESCs differentiate into hypermethylated formative pluripotent stem cells (fPSCs), with the methylation process mediated by DNMT3A and DNMT3B67,68. An mESC line with inducible co-expression of NLRP7 and TCL1A by doxycycline (Dox) and Shield1 (S1) were established (Fig. 5g, and Supplementary Fig. 10g)69. In fPSCs, we found that TCL1A overexpression severely disrupted methylation, reducing global levels from ~85.6% in control to 29.6% (Fig. 5h–j). Notably, NLRP7 coexpression counteracted TCL1A-driven hypomethylation, partially restoring methylation level to ~37.3% (Fig. 5h–j). All these results support a hypothesis that NLRP7 pathogenic variants impair the interaction with TCL1A and result in its accumulation in nuclear, where it binds to DNMT3A and inhibits de novo DNA methylation during human oogenesis (Fig. 5k).

Affinity to TCL1A is a predictive indicator for assessing the pathogenicity of NLRP7 variants
Subsequently, we explored the potential of our findings to predict the pathogenicity of mutated NLRP7. By comparing the experimental results of the NLRP7-TCL1A interaction, structure-based predictions from mCSM Affinity and commonly used in silico software PolyPhen-270,71, we found that these three methods demonstrated a high degree of consistency in predicting the pathogenic variants, especially for RHM-related variants located in the LRR domain (Fig. 6a). This suggests that structure-based predictions and NLRP7-TCL1A interaction testing are valuable detection methods for assessing the pathogenicity of NLRP7 variants.

To test it, we performed whole-exome sequencing on two patients with RHM and their families, and identified a homozygous L766R variant, as well as a previously reported homozygous R721W variant in NLRP7 (Fig. 6b). Detailed case reports for these two patients were provided in the Methods section. L766 and R721 were located within the LRR3-6 of NLRP7 (Fig. 6c). The mCSM Affinity and in silico analysis both predicted L766R and R721W to be reducing complex stability or potentially pathogenic, although the PolyPhen-2 prediction for R721W was an exception (Fig. 6d). To validate the accuracy of this prediction, we examined the effects of these two variants on their protein expression and their interaction with TCL1A. Our results showed that both variants, especially L766R, significantly reduced the expression of NLRP7 in HEK293T cells and severely disrupted their interaction with TCL1A, aligning with our predictions (Fig. 6e, f). Furthermore, we observed that L766R and R721W resulted in the nuclear accumulation of GFP-TCL1A in the injected mouse oocytes (Figs. 6g and 5c). In conclusion, these findings support the idea that assessing TCL1A binding affinity is a valuable approach for verifying and complementing traditional genetic diagnostic methods for NLRP7 pathogenic variants.

Methods
Ethical approval
Human Gamete and Embryo Research: The use of discarded human oocytes or embryos was approved by the Medical Ethics Committee of the Third Affiliated Hospital of Guangzhou Medical University, China (Approval numbers: 2020–139 and 2022–120). The collection and use of human samples were strictly conducted in accordance with the Declaration of Helsinki and the 2021 ISSCR Guidelines for Stem Cell Research and Clinical Translation. The abnormal human oocytes or cleavage embryos discarded during embryo culture were collected from donors (aged 25–40 years) who seek assisted reproductive treatment (ART) in the reproductive medicine center from 2021 to 2024. The discarding of oocytes and early embryos during ATR was based on rigorous evaluation by embryologists. Oocyte/embryo assessment followed standardized clinical protocols: (1) Abnormal oocytes: immature (GV/MI stage) or fertilization-failed (no two pronuclei at 16–18 h post-ICSI); (2) Suboptimal embryos: developmental arrest (>24 h stagnation) or Veeck grade < III cleavage-stage embryos (Day 3). To gather sufficient samples, there were no restrictions on the types of disease causes, treatments, and superovulation protocols for donors. Collected samples were immediately frozen in liquid nitrogen and stored at −80°C, or fixed at room temperature. The discarded human embryos and oocytes were only used for immunofluorescent staining, proximity ligation assay, and immunoprecipitation-mass spectrometry or western blotting. No further culture was performed on any abnormal embryos. Samples were randomly pooled prior to experimental use. Written informed consent was obtained from all donors.

Genetic Pedigree Study: The study about families of NLRP7 mutations was approved by the Medical Ethics Committee of West China Second University Hospital, Sichuan University, China (Approval number: 2024–243). The design and conduct of this study complied with regulations on the use of human participants and were in accordance with the principles of the Declaration of Helsinki. Venous blood samples from the patient and their family members underwent whole-exome sequencing following standard clinical diagnostic procedures. Written informed consent for the use of samples and relevant information in research and publication was obtained from all participants. In accordance with our institutional policies, remaining samples will not be shared with other researchers without a formal application and approval. Requests for materials should be addressed to Z.G., L.L.*, or D.D.

Animal Research: All animal maintenance and manipulations were conducted in accordance with the guidelines of the Institutional Animal Care and Use Committee of the Institute of Zoology, Chinese Academy of Sciences (IOZ-IACUC-2021-052). Mice were housed under standard conditions (12-h light/12-h dark cycle, temperature 23–25°C, humidity 40–65%) with ad libitum access to standard diet and water.

Case report
RHM patient 1: A 28-year-old female, as the only child of her non-consanguineous parents, at the time of presentation had experienced two complete hydatidiform mole (CHM) by the same partner. For the second CHM, chromosomal microarray of the products of conception showed normal results except for complete paternal uniparental disomy or triploidy, while immunohistochemical analysis of the histopathological specimen showed expression of cyclin-dependent kinase inhibitor 1C (CDKN1C, also known as p57Kip2). Despite the absence of molar pregnancies in other family members, a diagnosis of FRHM was suspected based on two CHM and no normal pregnancies.

RHM patient 2: The proband in the second case was a 41-year-old female, who presented with a history of four CHM and no normal pregnancies during a 10-year period. Following her fourth CHM, at the age of 36, she developed persistent trophoblastic disease requiring chemotherapy. Her non-consanguineous parents have another three daughters. The oldest sister has experienced two normal pregnancies resulting in a son and a daughter, while her youngest sister has given birth to a healthy daughter. The patient’s second sister has also experienced three consecutive CHM with no normal pregnancies, and has not attempted conception thereafter. Consequently, a diagnosis of FRHM was confirmed.

Co-IP coupling with MS
For Co-IP-MS, a total of 150 discarded human oocytes and embryos per subgroup were lysed in 150 μl lysis buffer (25 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% NP-40, 5% BSA, and 1× protease inhibitor) for 30 min on ice, then centrifuged at 13,400 × g for 15 min at 4°C. One microgram of anti-NLRP7 (Santa Cruz, cat#sc-377190, lot#J1218) or anti-NLRP5 (ABclonal, A9886, 0204550201) antibody was added into the lysates and incubated for 4 h (h) at 4°C, and 20 μL of protein G agarose beads were added into the lysates and incubated for 2 h. The beads were washed with the lysis buffer four times and boiled in 20 μl of 2× loading buffer.

For further protein identification, mass spectrometry analysis was performed. Briefly, the immunoprecipitates from two independent NLRP7 IPs (along with their respective controls) and one NLRP5 IP (with its control) were separated by short-run SDS-PAGE and stained with Coomassie Brilliant Blue. The gel bands were excised, destained, reduced with DTT, and alkylated with iodoacetamide, followed by overnight digestion with trypsin. The resulting peptides were extracted sequentially with increasing concentrations of acetonitrile. The digested peptide mixture was analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) on a nanoLC-QEXACTIVE system (Thermo Scientific) under the following conditions: column, reversed-phase C18 (75 μm × 20 cm, 3 μm particle size); flow rate, 300 nL/min; mobile phase A, 0.1% formic acid in 100% H₂O; mobile phase B, 0.1% formic acid in 100% acetonitrile; HPLC gradient program, 100 min gradient from 4 to 90% mobile phase B; capillary temperature, 320°C; electrospray voltage, 2.0 kV; scan range, 300–1600 m/z; dynamic exclusion time, 40 s; data acquisition mode, one full MS1 scan followed by MS2 scans of the 20 most abundant ions.

The database searching and protein identification were performed using the SEQUEST HT search engine in Thermo Proteome Discoverer (version 1.4.0.288) against the uniprot_human_2022.fasta database. Search parameters included: tryptic digestion with up to two missed cleavages, a precursor ion mass tolerance of less than 10 ppm, and a fragment ion mass tolerance of less than 0.02 mDa. Carbamidomethylation of cysteine was set as a fixed modification, while oxidation of methionine was set as a variable modification. For result filtering, Percolator was used for spectrum filtering with a Delta Cn threshold of less than 0.1, an FDR set to 1%, and a peptide confidence set to High for peptide filtering.

Plasmid construction
To construct the vectors for expression in HEK293T cells, the coding sequences of human SCMC-related genes, including NLRP5 (NM_153447.4), NLRP7 (NM_001127255.2), NLRP2 (NM_001174081.3), PADI6 (NM_207421.4), TLE6 (NM_053254), KHDC3L (NM_001017361.3), OOEP (NM_001428256.1), CCNI2 (NM_001039780.4), CDK5 (NM_004935.4), TCL1A (NM_001098725.2), pig TCL1A (NM_001435428.1) and NLRP7 (XM_021094979.1), Macaca mulatta TCL1A (XM_015144451.2) and NLRP7 (NM_001114354.1) were respectively sub-cloned into the p3×Flag-CMV10 and p5×Myc-CMV10 vectors, which is derived from the p3×Flag-CMV10 vector by replacing the 3×Flag tag with a 5×Myc tag83. To introduce mutations, a polymerase chain reaction (PCR) was performed using corresponding primers containing the mutated bases, with the wild-type vector as the template. The PCR products were subsequently digested with restriction endonuclease Dpn1 (New England Biolabs, R0176V) and transformed into TOP10 competent cells (TSINGKE, TSC-C12). To construct the vectors for the expression of NLRP7-TCL1A complex, full-length human NLRP7 and TCL1A were individually sub-cloned into the pCAG vector with N-terminal Flag-tag and N-terminal Strep-tag II. To construct vectors for VPICONE, the CDS of bait genes, mCherry, and the sequences coding the disordered region, transmembrane domain, and KASH domain (DTK) of mouse SYNE1 were amplified and were fused and sub-cloned into the pCDNA3.1(-) vector, while the CDS of prey genes were sub-cloned into pEGFP-C1. All vectors were confirmed by Sanger sequencing.

Transfection of vectors, western blotting, Co-IP, and Flag pull-down assay
HEK293T cells (EK-Bioscience, CC-Y1010) or COS-7 (HyCyte, TCO-C016) were pre-plated in each well of a 6-well plate and transfected with a total of 1.0 μg of plasmids and 3.0 μg of polyethylenimine (MKbio, 24765-1). The cells were further cultured for 24–36 h before harvesting. For Co-IP, similar to the method mentioned above, the cells were lysed in lysis buffer for 15–30 min on ice. Then, the lysates were centrifuged at 13,400 × g for 15 min at 4°C. A 300 μL aliquot of the lysate was incubated with 200 ng primary antibody and 15 μL of protein-G agarose beads (Beyotime, P2055) or protein A/G Magnetic Beads (Selleck, B23202) at 4°C for 2–4 h. The beads were incubated in TBS containing 0.1% Triton X-100 and 300 μg/mL 3×Flag peptides (Beyotime, P9801) for 1 h at 4°C. The immunoprecipitants and 15 μL of the input lysates were subjected to electrophoresis on a 4–20% gradient PAGE gel (WSHT, GSH2001). After transfer, the membranes were blocked with 5% non-fat milk for 1 h at RT. The membranes were then incubated with primary antibody overnight at 4°C or HRP-conjugated anti-tag antibody for 2 h at RT. The ImageJ Software (NIH) was used for analysis of band intensity. Flag antibody (ABclonal, AE092, 6100008467, 1:500) and Myc antibody (ABclonal, AE070, 6100005026, 1:500) were used for Co-IP.

For the Flag pull-down assay, Flag-NLRP7 or its mutants were expressed in HEK293T cells in 12-well plates, and Myc-TCL1A was expressed in 10 cm dishes. Cells were harvested after 36 h. Cell lysates containing Flag-NLRP7 or mutants were incubated with anti-Flag antibody and agarose protein A/G beads for 2 h at 4°C. After removing the lysates, the beads were incubated with an equal amount of lysate containing Myc-TCL1A for another 2 h at 4°C. Binding efficiency was calculated as [Co-IP(TCL1A)/Co-IP(NLRP7)]/Input(TCL1A), using band intensities from western blotting of co-immunoprecipitated samples25. All experiments were normalized against the wild-type control, which was assigned a reference value of 1.0.

The primary antibodies were used in this study included: HRP-conjugated Flag-tag antibody (ABclonal, AE024, 9200024001, 1:3000), HRP-conjugated Myc-tag antibody (ABclonal, AC206, 36000010268, 1:3000), HRP-conjugated anti-Tubulin antibody (ABclonal, AC030, 91000030001, 1:3000), HRP-conjugated anti-Actin antibody (ABclonal, AC043, 3600001946, 1:3000), CCNI2 antibody (Abcam, ab97767, GR241014, 1:1000), TCL1A antibody (Abcam, ab225718, 1032990, 1:1000), NLRP7 antibody (Santa Cruz, sc-377190, 1218, 1:1000), CDK5 antibody (ABclonal, A23502, 0016370101, 1:1000), NLRP2 antibody (ABclonal, A8233, 0038960201, 1:1000), TLE6 antibody (Abcam, ab76858, 1:1000), NLRP5 antibody (ABclonal, A9886, 0204550201, 1:1000), PADI6 antibody (Abcam, ab173952, GR194647, 1:1000), OOEP antibody (Abcam, ab185478, GR301408, 1:1000), GAPDH antibody (Cell signaling, 2118L, 10, 1:2000). KHDC3L antibody (homemade, 1:1000)72, mouse TLE6 antibody (homemade, 1:1000)50, mouse NLRP5 antibody (homemade, 1:1000)50, and mouse OOEP antibody (homemade, 1:1000)50 were used in previous studies. Secondary antibodies were as follows: HRP-conjugated goat anti-mouse antibody (ZSGB-BIO, ZB-5305, 1:5000) and HRP-conjugated goat anti-rabbit antibody (ZSGB-BIO, ZB-2301, 1:5000).

Protein and RNA level detection in transiently transfected cells, and Cycloheximide (CHX) Chase Assay
For protein and RNA level detection of NLRP7 and its variants, 250 ng of wild type NLRP7 and its mutant plasmids were mixed with 20 ng of pEGFP-C1 plasmid (serving as a spike-in control) and transfected into HEK293T cells. At 24 h post-transfection, the cells were divided into two aliquots for the detection of protein (Flag-NLRP7, GFP, and TUBB) levels by semi-quantitative western blot analysis and their corresponding mRNA levels by semi-quantitative PCR analysis, respectively. Primers were included for TUBB1 (forward primer, 5′-CAGTCTGGGGCAGGTAACAA-3′; reverse primer, 5′-CACTTTGGGTGAAGGCACCA-3′), GFP (forward primer, 5’-ATCATGGCCGACAAGCAGAA-3′; reverse primer, 5′-GGTACCGTCGACTGCAGAAT-3′), and Flag-NLRP7 (forward primer, 5′-ACCTGGGCCAGAATCATTTGT-3’; reverse primer, 5′-AGGCCCTACAGGTTGTCTTC-3′).

For the CHX chase assay, the cells were further cultured with medium containing 40 μg/mL CHX (MedChemExpress, HY-12320), and samples were collected at 0, 4, 8, and 12 h time points. Semi-quantitative western blot analysis were performed.

In vitro transcription and microinjection of mRNAs into mouse follicle and fully grown GV stage oocytes
Full-grown GV stage oocytes and growing oocytes were isolated from 7 to 8-week and 14-day-old ICR mice (Charles River). The mRNAs were transcribed in vitro with mMESSAGE mMACHINE™ SP6 Transcription Kit according to the manufacturer’s instructions (Life Technologies, AM1340). The concentration of mRNAs for injecting was about 100–1000 ng/μL. After injection, the oocytes were cultured in M2 medium supplemented with 100 ng/mL 3-isobutyl-1-methylxanthine (IBMX, Sigma, I7018) for 12 h, then were imaged or released for oocyte maturation. The live images were captured with a laser scanning confocal microscope (Carl Zeiss, LSM880).

Immunofluorescent staining, live imaging, and proximity ligation assay
The discarded human oocytes and embryos, as well as the transfected cells, were fixed with 4% paraformaldehyde (Sigma Aldrich, P6148) at RT for 30 min, washed three times with PBS/0.1% BSA (PBSA), and permeabilized with 0.5% Triton X-100 in PBSA for 30 min. After blocked with 1% BSA in PBSA at RT for 1 h, the samples were incubated with primary antibodies at 4°C overnight and then incubated with the fluorescent secondary antibody at RT for 2 h. DAPI (Sigma Aldrich, D9542) was used for staining DNA. For quantitative analysis experiments, photographs were taken for each sample under the same excitation intensity. PLA was performed according to the manufacturer’s protocol of Duolink® PLA Fluorescence kit (Sigma Aldrich, DUO92008). For pre-blocking the antibodies, the antigenic peptide (amino acids 16–170 in NLRP7, 0.4 μg/mL) specific to NLRP7 antibody (Santa Cruz, sc-377190, J1218, 1:200) and purified full-length TCL1A (0.5 μg/mL) specific to a TCL1A antibody at a molar ratio of 100:1 were incubated with antibodies for 2 h at RT. For immunofluorescent staining and live imaging in HEK293T cells, the cells were cultured in a 96-well glass-bottom plate specifically designed for laser confocal microscopy (Cellvis, P96-1-N). The images were captured with a Laser scanning confocal microscope (Carl Zeiss, LSM880). ImageJ software (NIH) and ZEISS ZEN 2012 software (Carl Zeiss) were used for image analysis. The primary antibodies were included: rabbit anti-Flag antibody (ABclonal, AE092, 1:100), TCL1A antibody (Abcam, ab225718, 1:200), mouse anti-NLRP7 antibody (Santa Cruz, sc-377190, 1:200), TLE6 antibody (Abcam, ab76858, 1:200), rabbit anti-NLRP7 antibody (Abcam, ab105405, GR7127, 1: 200), LAMIN B antibody (BioWord, AP6001, aa045181, 1:200). The Fluorescent dyes-conjugated secondary antibodies were included: FITC-Donkey anti-rabbit antibody (Jackson ImmunoResearch, 711097003, 1: 500), Alexa Fluor 647-conjugated-Donkey anti-rabbit antibody (ThermoFisher, A-31573, 1:500) and Alexa Fluor 546-conjugated-Donkey anti-mouse antibody (ThermoFisher, A-11030, 1:500).

Protein expression and purification
HEK293F cells (Invitrogen, R79007), maintained in 800 mL SMM 293-TII Expression Medium (Sino Biological, M293TII), were transiently co-transfected with 0.4 mg of NLRP7 plasmid and 0.4 mg of TCL1A plasmid using 2.4 mg of PEI. For protein purification, cells were harvested after transfection 72 h at 1800 × g, 15 min, 4°C. The cell pellet was resuspended in lysis buffer (25 mM Tris, pH 8.0, 150 mM NaCl) containing 1 mM phenylmethylsulfonyl fluoride, 0.8 μM aprotinin, 2 μM pepstatin A, and 5 μg/mL leupeptin and crushed with a high-pressure disrupter at 4°C. The cell debris were centrifuged at 50,000 × g, 45 min, 4°C, and then the supernatant was flowed through Flag resin (Merck) for two times. Resins were washed 4 times by 40 mL lysis buffer. Bound proteins were eluted with 0.2 mg/mL Flag peptide in lysis buffer. The proteins were concentrated and further purified by size-exclusion chromatography (SEC, GE, Superose 6 increase, 10/300 GL) with 25 mM HEPES pH 7.5, 150 mM NaCl, and 5 mM DTT. The peak fractions were collected and concentrated to 2.3 mg/mL for EM experiments.

Cryo-EM sample preparation, data acquisition, and analysis
The NLRP7-TCL1A complex was applied to glow-discharged 300 mesh gold grids (Quantifoil Au 300, R1.2/1.3) and subsequently vitrified using Vitrobot Mark IV (FEI), blotted for 2.5 s under 100% humidity at 4°C. The data were collected using a Titan Krios microscope (Thermo Scientific) operated at 300 kV equipped with a K2 Summit detector (Gatan) and an EPU 2.5 software (Thermo Scientific) at Sichuan University. Details are presented in Supplementary Table 2.

Single-particle data processing was performed using cryoSPARC 4.1.2. 2054 movies were collected and processed for motion correction and contrast transfer function estimation. A total of 1,195,156 particles were extracted from micrographs with Blob picker and used to generate 2D class averages. Subsequently, ab initio reconstruction was performed, resulting in low-resolution maps for template picker. For the NLRP7-TCL1A dimer state, a total of 1,259,946 particles were automatically picked and extracted from micrographs, using the template derived from one good map in the dimer state. After several rounds of 2D classification, ab initio reconstruction, and heterogeneous refinement, 167,314 good particles were selected and applied to non-uniform refinement and local refinement, yielding a map of 3.58 Å. For the NLRP7-TCL1A monomer state, template picker was also performed, ultimately resulting in the acquisition of 943,858 particles. As mentioned above, a subset of 165,276 particles was ultimately selected for non-uniform refinement, local refine and Deep EMhancer, resulting in a 3.48 Å map.

The initial model of NLRP7-TCL1A was generated by docking the AlphaFold2-predicted NLRP7 and TCL1A into the cryo-EM map with UCSF ChimeraX software for structure visualization, and then performed the manual adjustments by COOT84,85. All models were refined against the cryo-EM map and validated using the real-space refinement tool with PHENIX86.

MTase-Glo methyltransferase assay
Nucleosomes were reconstituted following a previously described method87. Briefly, four types of histone (H2A tagged with a C-terminal 6×His tag, H2B, H3, and H4) were expressed in Escherichia coli BL21(DE3) (Invitrogen, EC0114). The cells were lysed in buffer (25 mM Tris, 150 mM NaCl, pH 8.0, 1 mM PMSF) using a high-pressure cell disruptor. Inclusion bodies containing histones were collected by centrifugation, resuspended in denaturation buffer (50 mM Tris, 8 M guanidine hydrochloride, 1 M NaCl, pH 8.0), and clarified by centrifugation at 27,000 × g for 30 min. The supernatants containing the histones were quantified and mixed at a 1:1:1:1 molar ratio. The mixed proteins were dialyzed twice at 4°C against dialysis buffer (25 mM Tris, 2 M NaCl, 1 mM DTT, pH 8.0)—first for 2 h, then overnight in fresh buffer. Aggregated proteins were removed by centrifugation at 27,000 × g for 30 min. The reconstituted histone octamer was bound to Ni-NTA resin by passing through the column twice, washed with 25 mM Tris, 2 M NaCl, 5 mM imidazole (pH 8.0), and eluted with 25 mM Tris, 2 M NaCl, 300 mM imidazole (pH 8.0). The eluate was concentrated and further purified by SEC in 25 mM Tris, 2 M NaCl, 2 mM DTT (pH 8.0).

The purified histone octamer (0.85 mg/mL) and DNA (0.96 mg/mL) were mixed in 25 mM Tris, 2 M NaCl, 2 mM DTT (pH 8.0), then dialyzed by a salt-gradient method against 25 mM Tris, 2 mM DTT (pH 8.0). Protein aggregates were removed by centrifugation at 17,000 × g for 5 min, and the sample was further purified by SEC in 20 mM HEPES, 1 mM DTT (pH 7.4). The DNA sequence used was: 5′-GAGAAAGGATCCCTCGAGCGCGGCCGCCCTGGAGAATCCCGGTGCCGAGGCCGCTCAATTGGTCGTAGACAGCTCTAGCACCGCTTAAACGCACGTACGCGCTGTCCCCCGCGTTTTAACCGCCAAGGGGATTACTCCCTAGTCTCCAGGCACGTGTCAGATATATACATCCTGTGCATGTATCTCGAGCGCGGCCGCCCTG-3′.

The activity of DNMT3A (residues 220-912) was assessed using the MTase-Glo Methyltransferase Assay Kit (Promega, V7601) according to the manufacturer’s protocol. The reaction mixture was optimized to a final volume of 12.8 μL, containing 0.5 μM DNMT3A or DNMT3A-TCL1A complex, 10 μM SAM, and 2 μM nucleosome. The reactions were performed in buffer containing 25 mM Tris (pH 8.0), 50 mM NaCl, 1 mM DTT, 5% glycerol, and 0.2 mg/mL BSA, and incubated at 37°C for 1 h. Subsequently, 3.2 μL of 5× Methyltransferase-Glo reagent was added, and the mixture was incubated at 30°C for 30 min, followed by the addition of 16 μL of Methyltransferase-Glo detection solution and a further 30 min incubation at 30°C. Luminescence was measured using a Synergy H1 microplate reader (Synergy H1, BioTek).

Constructing mouse embryonic stem cells lines (mESCs) and inducing formative pluripotent stem cells (fPSCs)
The mESCs (Millipore, CMTI1) on a 129/SVEV strain background were maintained in N2B27/2i/Lif medium, including N2B27 medium supplemented with 1 μM PD035901 (LC Laboratories, P-9688), 3 μM Chir99021 (LC Laboratories, C-6556), and 1000 units/mL hLIF (Millipore, ESG1107).

To construct the inducible TCL1A expressing mESCs (mESCS1(-)), a destabilizing domain (DD) was fused with the N-terminus of Flag-TCL1A and cloned and inserted into a modified piggyBac transposon vector. Next, the mESCs were electrotransfected. The positive clones were screened in 2i/Lif medium with G418 for 1 week. The clones of cells, expressing TCL1A after induction with Shield1 (S1, MCE, HY-112210), were selected for further research. An mESCS1(-) clone were further electrotransfected with a piggyBac transposon vector harboring a TETON system controlling the expression of 5×Myc-NLRP7. The cells were selected in 2i/LIF medium containing puromycin for 1 week and designated as mESCs S1(-) DOX(-). The superscripts (+) and (-) indicate the presence and absence of each inducer (S1 or Dox), respectively.

fPSCs were generated. On day 0, the mESCs S1(-) Dox(-) line were digested into single cells. Then, the cells were suspended and mixed with an equal volume of Matrigel (BD, BD356230). The cells were plated in a dish and further incubated for 1 h at 37°C until the Matrigel solidified. The cells in Matrigel were subsequently cultured in prewarmed fPSC induction medium (N2B27 medium supplemented with KSR, activin A, and bFgf2) at 37°C and 5% CO2 for 3 days. To induce the protein expression, the small chemical Shield1 at a working concentration of 0.05 μg/mL or doxycycline at working concentrations of 2.0 μg/mL was added to the fPSC induction medium on day 0. On day 3, the fPSCs were harvested.

WGBS for fPSCs and data processing
The genomic DNA of fPSCs cultured in 6-well plates was isolated with a DNA isolation mini kit (Vazyme, DC112). WGBS was conducted, following the instructions of the bisulfite kit (Vazyme, EM101) and library kit for bisulfite conversion and library preparation (Vazyme, NE103), sequentially. The libraries were sequenced on the Illumina NovaSeq X plus platform to generate 150-base-pair paired-end reads, achieving an average sequence yield of 20 gigabases per sample. The quality of the raw sequencing reads was initially evaluated using FastQC v0.11.9. The quality of the raw sequencing reads was assessed using FastQC v0.11.9. Adapter contamination and low-quality bases were subsequently trimmed from the reads using Trim_galore v0.6.10 with default parameters. The processed read pairs were aligned to the mm9 mouse genome assembly using Bismark v0.22.3. After alignment, the deduplicate_bismark command within Bismark was employed to identify and remove PCR duplicates, thereby optimizing the dataset for downstream analyses. Methylation states were extracted from deduplicated reads using MethylDackel v0.6.1. Following a consistency check between replicates, methylation profiles were combined to improve coverage depth and resolution by aggregating C/T counts at each CpG site.

For genomic segment analysis, the genome was segmented into continuous intervals of 100 base pairs (bp), 1 kilobase (kb), or 1 megabase (Mb). On average, a minimum coverage criterion of three CpG sites per 100 bp was applied. The methylation level within each bin was determined as the ratio of methylated cytosines (C) to the sum of methylated and unmethylated cytosines (C + T) across all CpG sites within the interval. When comparing overexpressed samples to controls, only bins covered in both conditions were included in the analysis. The total methylation score for each interval was calculated as the sum of the methylation levels across all the CpG sites within that region. All analyses were performed using custom LINUX and R scripts, which are available upon request to facilitate verification and replication of our results.

STEM-seq for mouse oocytes and data processing
Oocytes were lysed with 10 μL of STEM-seq lysis buffer (10 mM Tris-HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl2, 0.1 mM EDTA, pH 8.0, 0.5% NP-40) containing 1 μL of protease K (New England Biolabs, P8107S) for 3 h at 55°C, followed by heat-inactivation for 1 h at 72°C. Samples were then treated with 1 μL of dsDNA Fragmentase (New England Biolabs, M0348AA) for 30 min before being subjected to bisulfite conversion using EZ DNA Methylation-Lightning Kit (ZYMO, D5030). Libraries were prepared using the EpiArt DNA Methylation Library Kit (Vazyme, NE103) according to the manufacturer’s instructions. In brief, a 3′ adapter was first added to the converted and denatured ssDNA. After DNA extension and purification, a 5′ adapter was added. dsDNA with both 3′ and 5′ adapters were then subjected to PCR amplification, and libraries were sequenced on an Illumina Hiseq X Ten or Novaseq 6000 system. For all samples, λ-DNA (Promega, D150A) was added with a mass ratio of 1:200 before bisulfite conversion for conversion rate analysis.

All paired-end reads were first processed using cutadapt (v1.18) (Martin, 2011) with the parameter -a GGGGGGGGGGGGX -a AGATCGGAAGAG -A AGATCGGAAGAG -A GGGGGGGGGGGGX -g CTCTTCCGATCT -G CTCTTCCGATCT -n 10 -max-n 0.05 -q 20,20 -u -10 -U 10 -m 30 -e 0.2, to remove adapters and the low-quality reads before mapping. The filtered reads were then aligned to the mm9 genome using Bismark (v0.18.1) (Krueger and Andrews, 2011) with default parameters. After removing multiple-mapped reads and PCR duplicates, the unique-mapped reads were used for CpG methylation calling with bismark_methylation_extractor. CpG sites that were covered at least five times were used for downstream analyses. For 200-bp bin analyses, all reads covering CpG sites within the bin were pooled and used to estimate the methylation level for the bin. The methylation level was calculated as the ratio of methylated CpG to the total number of CpG sites within each bin. The λ-DNA was used for conversion rate calculation.

Whole-exome sequencing
Peripheral blood samples were obtained by venipuncture from family 1 (I-2 and II-2) and family 2 (I-1, I-2, II-1, II-2, and II-3). Genomic DNA was extracted following the manufacturer’s protocol of the kit (Vazyme, DC112-01). Whole-exome sequencing for family 1 (I-2 and II-2) and family 2 (I-1, I-2, and II-2) was outsourced to Berry Genomics Co., Ltd. for analysis. The GRCh38 (hg38) genome was utilized for annotation. The nomenclature for the single-nucleotide variants (SNVs) was according to guidelines by the Human Genome Variation Society (HGVS) (http://varnomen.hgvs.org). The pathogenicity of the detected SNVs was systemically evaluated based on the guidelines of the American College of Medical Genetics and Genomics (ACMG) and the Association for Molecular Pathology (AMP).

Statistical analysis
All biochemical experiments were independently repeated at least two (human materials) or three times to ensure the reproducibility. The data were presented as mean ± SEM (Standard Error of the Mean) or as the median with interquartile range using box plots. Statistical analysis was performed using the unpaired two-tailed t-test with GraphPad Prism 9 software, and P-values < 0.05 was indicated as significant difference.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript titled “TCL1A Mediates DNA Methylation Defects in NLRP7 Mutation-Associated Recurrent Hydatidiform Mole” by Gao et al. provides significant insights into the molecular mechanism underlying NLRP7 mutation-associated recurrent hydatidiform moles (RHM). The authors identified TCL1A, which is an DNA methyltransferase 3A (DNMT3A) inhibitor, as a key interactor of NLRP7, and then determined the cryo-EM structure of NLRP7-TCL1A complex. Through structural, biochemical, and cell-based analyses, they demonstrated that most pathogenic NLRP7 mutations disrupt TCL1A binding, leading to aberrant nuclear translocation of TCL1A and inhibition of DNMT3A-mediated DNA methylation. Furthermore, they identified a novel pathogenic NLRP7 variant (L766R) in RHM patients, underscoring the clinical relevance of the NLRP7-TCL1A interaction for predicting pathogenic NLRP7 variants.\nRepresenting a significant advance in reproductive and developmental biology, the manuscript is well-structured, methodologically rigorous, and clinically relevant. Nevertheless, some points need further clarification or elaboration. Below are specific questions:\n1. The cryo-EM structures reveal two distinct states of the NLRP7-TCL1A complex. What biological purpose each state serves (e.g., enhanced stability, cooperative binding, or functional switching)? Whether these states interconvert dynamically during oogenesis or early embryogenesis? Whether disease-associated mutations perturb the equilibrium between these states and contribute to pathology?\n2. Figure 5a-b shows nuclear TCL1A accumulation in human cleavage-stage embryos but not GV/MI oocytes. What could be the signals that trigger TCL1A release in embryos (e.g., degradation of NLRP7, activation of embryonic transcription factors, or post-fertilization cues)?\n3. The LRR domain harbors most RHM-linked mutations (Fig. 4b), do other pathogenic NLRP7 mutations also impair TCL1A binding. Do these mutations indirectly disrupt NLRP7 folding, or do they directly affect TCL1A binding?\nMinor points:\n1. Figure 5a: the information of each column should be labeled.\n2. Figure 6a: \"CO-IP (Mutation-WT)\" should be \"Co-IP (mutant vs. WT).\"\n3. References: Ensure all in-text citations (e.g., \"submitted to Nature Communications\" in line 399-400) are formalized.", "reviewer_2": "In this manuscript Gao and colleagues describe how NRP7 sequesters TCL1A in the cytoplasm to ensure that appropriate oocyte methylation by DNMT3A. NLRP7 mutations result in biparental hydatidiform moles (biHMs) that classically show a loss of maternal imprints. The mechanisms leading to this hypomethylation has been a mystery for years, with many groups actively trying to solve this conundrum. The manuscript is easy to follow and the figures are informative. However, in places the manuscript lacks detail for readers not so familiar with some techniques (i.e those used a better understand the influence of the genetic variants assessed) that need clarification.\nPlease note, I am not qualified to comment on the cryo-EM aspects of the manuscript.\nMajor comments\n- There are discrepancies in published co-IPs for NLRP7 interactions with SCMC members. Fig1B show lack of interaction with KHDC3L and PADI6 in native samples (same for NLRP5), but others have shown NLRP7 interacts with all members of the SCMC using over-expression (Han et al., 2023). It would be interesting to know if this is due to an experimental variable (i.e. wash conditions). Therefore, please at a table summarizing all NLRP7 co-IP results from different publications and include as a supplemental figure.\n- Has the VPICONE assay been used to assess the NLRP7-SCMC interactions or only with TLE6?\n- Results line 176: “TCL1A exclusively binds to NLRP7” is not strictly true as Fig1C-D show weak interaction with NLRP5. Please remove the word “exclusively”.\n- Results line 211: “…we could not reconstitute the well-behaved complex….”. Please add a line say why. Is this because NLRP7 is cysteine rich and thus has a propensity to form disulphide bonds and insoluble aggregates during purification?\n- The evidence from the works is compelling – mutations in NLRP7-LRR domain releases TCL1A which subsequently inhibits DNMT3A. However more detail is required to explain the quantitative nature of the transient protein expression and co-IPs. Is it not possible that reduced band intensities could be due to reasons (both valid pathological mechanisms) – reduced protein stability/expression and decreased physical interactions. How was transfection efficiency controls between experiments/mutations? It is assumed that all constructs were generated using the plasmid backbone. Did the authors consider confirming some observations using more sophisticated methods such as cycloheximide pulse-chase (for stability) and Isothermal titration calorimetry or Surface Plasmon resonance for precisely quantifying interactions?\n- In Figure 4C, please use the same colour code from 4A, B when labelling the mutations. Please add the actual PolyPhen-2 scores to Extended Data Table 3 and add a score/% scales to the bar graphs in Figure 5A.\n- NLRP7 is highly expressed in oocytes and is subject to maternal decay. However, there is a sharp rise in expression in blastocysts. The authors should speculate if this NLRP7 could regulate embryonic de novo remethylation.\n- KHDC3L mutations also cause biHMs with extreme loss of methylation in oocytes. As TCL1A does not interact with KHDC3L, do the authors propose another mechanisms is involved here. Please include a line discussing this in the manuscript discussion.\nMinor comments\n- The evidence from the works is compelling – mutations in NLRP7- LRR domain releases TCL1A which subsequently inhibits DNMT3A. However, the authors should speculate in the discussion what is the normal role of TCL1A in pre-implantation embryos and reprogramming.\n- Abstract – line 32: The first line needs amending as direct links between NLRP7 and de novo methylation in oocytes has not been proven (until this work). Please state that this is suggested.\n- Introduction line 62: Please state that RHM mutations in PADI6 and NLRP5 have mixed reproductive histories, whereas NLRP7 and KHDC3L biallelic variants normally result in biHM.\n- Introduction line 76: Please state that heterozygous mutations in NLRP7 and SCMC proteins have been observed in MLID.\n- Results line 171: When describing NLRP7 isoforms in the text, please add the accession number of the transcript.\n- NLRP2 and 7 are highly similar (due to genome duplication), but the results presented here indicate specialization for paralogs. Please state that it seems like that NLRP2 is a core SCMC subunit, whereas the main role of NLRP7 is sequestering TCL1A.\n- Results line 334: Does TCL1A increase NLRP7 expression or stabilize it?\n- Results line 398/Fig 5g: Please state this is a in vitro assay using purified DNMTs. The result show that DNMT3A relative activity was greatly increased with the addition of DNMT3L. However, human oocytes lack DNMT3L. Does the TCL1A inhibition require DNMT3A-3L complex?? Is the difference between 3A and 3A/TCL1A significant?\n- You show that NLRP7 isoform 1 does not have the same role in regulating TCL1A cytoplasmic localisation as other isoforms, so what do you propose is it function?", "reviewer_3": "The authors found that NLRP7 anchors TCL1A in the cytoplasm of oocytes via a physical interaction, and that defects in NLRP7 allow TCL1A to localize to the nucleus, resulting in inhibition of de novo DNA methylation by DNMT3A during oogenesis. Recently, it was reported that NLRP5 stabilizes UHRF1 in the cytoplasm and vice versa. Defects in NLRP5 reduce the amount of UHRF1 in both the cytoplasm and the nucleus of mouse oocytes, possibly leading to impaired maintenance of DNA methylation at gDMRs during preimplantation development. While patients with pathogenic variants in NLRP7 exhibit hypomethylation of maternally methylated gDMRs, those with pathogenic variants in NLRP5 show hypomethylation of both maternally and paternally methylated gDMRs. Thus, the mechanistic differences underlying the hypomethylation of multiple gDMRs may account for the differential patterns of hypomethylated gDMRs observed in patients or hydatidiform moles with NLRP7 versus NLRP5 pathogenic variants. It is also noteworthy that the interaction between NLRP7 and TCL1A appears to be primate-specific. The authors’ experiments using human materials provide valuable insights, and their findings are conceptually novel and important for understanding molecular pathogenesis behind hydatidiform mole and MLID. However, there are several discrepancies between the IP-MS/MS results and the Western blotting results. Additionally, despite the fact that human oocytes do not express DNMT3L, a known co-activator of DNMT3A, the enzymatic analysis in the study employed DNMT3L. Clarifying these discrepancies and removing experiments that are physiologically irrelevant would greatly improve the clarity and impact of the manuscript.\nMajor points\n1. It is reasonable to compare the interacting proteins of NLRP7 and KHDC3L to identify key factors potentially involved in de novo DNA methylation in oocytes, as pathogenic variants in these genes lead to hypomethylation of maternal gDMRs. However, it is unclear why the authors included interacting proteins of NLRP5 in this comparison, given that pathogenic variants in NLRP5 cause hypomethylation of both paternal and maternal gDMRs. Please clarify the rationale for including NLRP5 in the comparison of interacting proteins with NLRP7 and KHDC3L.\n2. The authors should provide the IP-MS/MS data with peptide counts and fold changes in Extended Data Table 1. This would help readers assess the relative binding affinities of proteins co-immunoprecipitated with these factors. Notably, UHRF1 was among the proteins co-immunoprecipitated with NLRP5—but not NLRP7—according to Extended Data Table 1. This finding may support the authors’ conclusion that there are two distinct mechanisms underlying the pathogenesis of hydatidiform mole and MLID: one involving dysregulation of UHRF1, and the other involving defects in de novo DNA methylation (line 481). The authors may consider briefly mentioning this point.\n3. There are numerous discrepancies between Fig. 1a (IP-MS/MS results) and Fig. 1b–h (Western blotting). For example, although CDK5, KHDC3L, PADI6, CCNI2, and TCL1A were co-immunoprecipitated with both NLRP7 and NLRP5 in Fig. 1a, CDK5, KHDC3L, and PADI6 were not co-immunoprecipitated with NLRP7 in Fig. 1b; PADI6, CCNI2, and KHDC3L were not co-immunoprecipitated with NLRP5 in Fig. 1c; and NLRP7 was not co-immunoprecipitated with NLRP5, PADI6, CCNI2, KHDC3L, CDK5, or OOEP in Fig. 1d. Moreover, since immunoprecipitation of FLAG-KHDC3L was unsuccessful in Fig. 1d, it is not possible to determine whether Myc-NLRP7 was co-immunoprecipitated with FLAG-KHDC3L. Nevertheless, the authors concluded that Western blot validation confirmed specific enrichment of these interactors (line 142). A more cautious interpretation is required, such as stating that “only high-affinity interactions were confirmed” and/or that “oocyte-specific proteins or post-translational modifications may be required for some interactions, which could not be detected in HEK293T cells.” Please address these discrepancies, and if necessary, consider repeating the experiments.\n4. The authors found that not only NLRP7, but also NLRP5, interacts with TCL1A (Fig. 1a and 1c.) However, they excluded NLRP5 for further analyses. The authors explained the reason as co-IP assays confirmed TCL1A specifically binds to NLRP7 and itself (line 167). This description appears inaccurate, as NLRP5 was shown to interact with TCL1A, as described above. Do the authors consider the interaction between NLRP5 and TCL1A to be a false positive? If the interaction between TCL1A and NLRP7 was preserved under stringent conditions, while the interaction with NLRP5 was lost under the same conditions (was the condition in Fig. 1h more stringent than that in Fig. 1a-c?), the authors should clarify this. Otherwise, please explain the discrepancy and the rational for focusing solely on NLRP7.\n5. The authors focused on TCL1A for further analysis due to its known role as a potential regulator of DNA methylation (line 167). While this rationale is reasonable, the functions of CNNI2 and CDK5 should also be briefly described to help readers better understand why TCL1A was prioritized.\n6. The authors reported that TCL1A interacts with NLRP7 isoforms 2 and 3 (lines 169-170). Are these isoforms expressed in human oocytes?\n7. In the legend for Figure 5, the authors state: “Notably, since DNMT3L is not expressed in human oocytes (citation #85), it remains unclear how DNMT3A is activated. One hypothesis suggests that DNMT3B3, a catalytically inactive isoform of DNMT3B, substitutes for the function of DNMT3L (citation #86).” This point is highly critical and should be described in the main text rather than in the figure legend. A previous study has reported that the inactive DNMT3B isoform is unlikely to replace DNMT3L in multiple species, including humans (Behluli et al., 2023; https://doi.org/10.1186/s13072-023-00518-2). Therefore, it is unclear why the authors examined the DNA methyltransferase activity of the DNMT3A/3L complex (Fig. 5f) and discussed the possible involvement of DNMT3B3. Since DNMT3A alone exhibits DNA methyltransferase activity, and TCL1A appears to inhibit this activity in Figure 5f, the authors should compare the activity among the blank, DNMT3A alone, and DNMT3A with TCL1A, with appropriate statistical evaluation. If the authors have evidence that DNMT3B3 is expressed in human oocytes, it would be useful to include DNMT3B3 in the analysis.\n8. The authors expressed wild-type and disease-related NLRP7 pathogenic variants in HEK293T cells (Extended Data Figure 10). They concluded that these variants impair the expression levels of NLRP7 (line 127). Since these variants likely reduce protein stability, I recommend that the authors use the term “stability” instead of “expression” for clarity. In addition, the authors should show mRNA expression levels, which are expected to be consistent across different plasmid vectors, to support this conclusion. Because transient expression analysis was performed, transfection efficiency may vary among plasmids.\n9. Line 73: In the Introduction, the authors mention that oocytes from a patient with biallelic KHDC3L variants exhibited genome-wide hypomethylation, suggesting that pathogenic variants in NLRP7/KHDC3L disrupt de novo DNA methylation during oogenesis. However, the roles of KHDC3L have not been elucidated in this manuscript. Since the interaction between NLRP7 and KHDC3L was barely detectable in Fig.1b and Fig.1d (and the experiment in Fig.1d may not have been successful), what could be the possible molecular mechanism underlying the pathogenesis in patients with pathogenic variants in KHDC3L? Please address this point in the Discussion section.\n10. Line 484: In the Discussion, the authors state that abnormal nuclear translocation of UHRF1/DNMT1—key regulators responsible for both de novo establishment and maintenance of DNA methylation—leads to genomic methylome dysregulation in 2-cell embryos with maternal disruption of PADI6 or NLRP14, which may mechanistically explain the MLID observed in patients harboring hSCMC gene mutations. However, this abnormal nuclear translocation of UHRF1/DNMT1 in these mutant 2-cell embryos has been reported to result in hypermethylation, which differs from the hypomethylation observed in patients with MLID (Giaccari et al, Genes Devs, 2024; Yan et al Nat Genet, 2023). In contrast, Unoki et al. demonstrated that defects in NLRP5 reduce the levels of UHRF1 in both the cytoplasm and the nucleus, potentially leading to impaired maintenance of DNA methylation at gDMRs during preimplantation development (Uemura et al., Life Sci Alliance, 2023; Unoki et al., Hum Mol Genet, 2024). Therefore, when discussing the involvement of UHRF1/DNMT1 in the pathogenesis of hydatidiform mole and MLID, the NLRP5 case may provide a more appropriate model.\nMinor points\n1. There is inconsistent usage of the terms “pathogenic variants” and “mutations” throughout the manuscript. Current recommendations suggest using pathogenic variants, unless referring specifically to de novo mutations detected only in the patient. In the authors’ context, pathogenic variants is more appropriate. Please revise the manuscript accordingly.\n2. In Fig. 1c, there is a visible band in lane 2 (designated as an empty lane) of the blot probed with the anti-CCNI2 antibody. What does this band represent?\n3. In Fig. 1L, the transfected plasmids should be clearly indicated and labeled consistently with the 5% input lanes.\n4. Line 396: Related to major point #7, the authors write, “DNMT3A and its activators DNMT3B or DNMT3L.” However, while DNMT3L is an established activator of DNMT3A, DNMT3B is not. Instead, the catalytically inactive isoform DNMT3B3 might act as an activator. Please revise this sentence for accuracy.\n5. In Fig. 5E, the anti-FLAG antibody detects two bands in lanes 1 and 2. What does the lower band represent? Cleaved protein? Please add an explanation of the band in the figure legend.\n6. In Extended Data Fig. 9A–C, what do the arrowheads indicate? Please include a clear explanation in the figure legend.\nTypos\nLine 94: criticalfor -> critical for\nLine 140: KHDC3 -> KHDC3L\nLine 361: Aaberrant -> aberrant\nLine 398: Fig.5g -> Fig. 5f\nLine 1159: g -> f", "reviewer_4": "The authors investigate the role of pathogenic NLRP7 variants, which are linked to 55% of RHM, in disrupting de novo DNA methylation in human oocytes. They identify TCL1A, a DNMT inhibitor, as a key interacting partner of NLRP7. The cryo-EM structure of their complex reveals that many RHM-associated mutations in NLRP7 impair this interaction. NLRP7 likely safeguards the oocyte methylome by sequestering TCL1A in the cytoplasm, preventing its nuclear entry and subsequent suppression of DNMT-mediated methylation. The study also identifies L766R as a novel pathogenic NLRP7 variant, offering insights into hypomethylation pathogenesis.\nThis study is significant due to its thorough mechanistic investigations. However, couple of major points need addressing to enhance the strength of the study.\nMajor Comments:\n1. Figure 5F uses an MTase-Glo assay to evaluate DNMT activity. Given that DNMT activity is central to the study, a more comprehensive assay is needed to assess how NLRP7-TCL1A interactions impact downstream DNMT activity. The authors should perform STEM-seq and/or WGBS.\n2. Understanding how the dysregulation of NLRP7-TCL1A interaction leads to TCL1A's nuclear accumulation would further strengthen the mechanistic insights. Does the disruption of interaction cause specific post-translational modifications that sequester TCL1A to the nucleus? The authors should perform affinity purification followed by mass spectrometry of TCL1A when interacting with NLRP7 to identify and showcase potential PTMs that block TCL1A from being accumulated in the nucleus.\nMinor Comments:\n1. In line 142, the authors claim interaction confirmation via western blotting (Fig 1b-c) for all interacting partners. However, some proteins identified in IP-MS, such as CDK5, are not confirmed in IP-WB. The authors should either provide more robust confirmation data or rephrase the sentence for specificity.\n2. In line 398, the authors refer to another study submitted to Nature Communications. They should either include relevant data from this study or remove Figure 5h altogether, or modify the phrasing.\n3. Figure legends are mislabeled.\n4. There are several minor grammatical errors, such as inconsistent sentence casing and typos, which should be corrected before submission."}, "author_response": "We would like to thank all reviewers for their thorough and insightful comments, which have not only enhanced our current research but also provided valuable insights for our future studies. We have carefully addressed the reviewers' concerns and revised the manuscript accordingly. To facilitate reviewer reading, we have also displayed the revised Figures (both main and Supplementary) as part of the Response Figures (Response Fig.xx, also shown as Fig.xx or Supplementary Fig.xx). Below we provide detailed responses to each comment in blue, and corresponding changesarehighlightedinyellowinthemanuscript.\nPoint-by-point Response\nResponse to Reviewer #1:\n[...]"}, {"round": 2, "reviewer_comments": {"reviewer_1": "I have reviewed the authors' responses to my comments and the revised manuscript. They have satisfactorily addressed all of my concerns. Therefore, I recommend acceptance of the paper for publication.", "reviewer_2": "The authors have made a great attempt addressing my queries. I have one final request before I feel the maunscript is ready for publication. The authors used CHX chase assays to look at protein stability (Suppl Fig 9), but the protocol details have not been added to the methods.", "reviewer_3": "The authors have thoroughly and sincerely addressed all my questions. I greatly appreciate their tremendous effort. I have no further requests, and I believe the manuscript is now ready for publication.", "reviewer_4": "The authors have adequately addressed my comments. I recommend adding Response Figure 11 to the supplementary data and depositing the mass spectrometry data and methods in the supplementary files. While the quality of some MS/MS sites was suboptimal, sharing this data will benefit the research community by providing novel insights that can be further validated in future studies."}, "author_response": "We would like to express our sincere gratitude to the the Reviewers for the time and effort they have dedicated to evaluating our work. Their professional guidance has been invaluable in strengthening our manuscript. Below, we provide a point-by-point response to each of their comments.\n[...]"}]

Decision letter