Germline epigenome editing identifies H3K9me3 as a mediator of intergenerational DNA methylation recovery in mice

Horii, Takuro; Morita, Sumiyo; Hino, Shinjiro; Hino, Yuko; Fukushima, Hiroto S.; Kobayashi, Ryosuke; Kimura, Mika; Nakao, Mitsuyoshi; Mizukami, Yoichi; Inoue, Azusa; Hatada, Izuho

doi:10.1038/s41467-025-67488-9

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Published: 25 December 2025

Germline epigenome editing identifies H3K9me3 as a mediator of intergenerational DNA methylation recovery in mice

Nature Communications volume 16, Article number: 11200 (2025) Cite this article

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Abstract

Inter/transgenerational epigenetic inheritance is a crucial and controversial theory that could reshape the concept of genetics. To investigate this theory directly, we invent a system for targeted reprogramming of epigenetic memory in mouse sperm. Using this system, we erase DNA methylation at the differentially methylated region of the H19 gene (H19-DMR) in sperm, which causes Silver-Russell syndrome-like phenotypes in F1 offspring. Although DNA methylation is fully lost in the sperm, it is partially restored during pre-implantation development, suggesting the existence of epigenetic memory that instructs de novo DNA methylation. Importantly, targeted removal of histone modifications in zygotes reveals that tri-methylation at lysine 9 of histone H3 (H3K9me3), which is deposited shortly after fertilization, is required for the subsequent de novo DNA methylation at the H19-DMR. Thus, our study provides a robust germline editing tool, which reveals partial intergenerational inheritance and no transgenerational inheritance at the model locus. Furthermore, we identify H3K9me3 as a mediator for DNA methylation recovery also acting at imprinted loci.

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Introduction

Phenotypic inheritance is believed to be achieved via the transmission of genomic DNA. Paradoxically, ancestral environmental exposures, such as abnormal nutritional conditions, toxins, and stress, can lead to disease and phenotypic changes in offspring^{1,2,3,4,5,6,7,8,9}, providing circumstantial evidence suggesting a non-Mendelian inheritance phenomenon, where epigenetic memory is passed to the next generation through the germline. While these data are still under discussion because of difficulties in showing the absence of genetic mutations, DNA methylation, histone modifications, and RNA may contribute to the primary molecular mechanisms underlying epigenetic inheritance^{1,2,10,11,12,13,14}. For example, paternal prediabetes induced by feeding a high-fat diet increases the susceptibility of offspring to diabetes³, while Sun et al. reported that paternal cold exposure enhances brown adipose tissue activity in offspring⁴. In these cases, altered DNA methylation was detected in both sperm and offspring; however, it does not necessarily follow that germline DNA methylation changes are passed to the next generation, as it is equally possible that epimutations are first reset after fertilization and then reestablished during development by yet undetermined factors.

To provide more direct evidence for inter/transgenerational epigenetic inheritance, it is necessary to show that epimutations induced in the germline can be transmitted to the next and later generations. In this study, we define intergenerational inheritance as the transmission of epigenetic alterations originating in the gametes to the immediate offspring (F1 generation). By contrast, transgenerational inheritance refers to the sustained presence of these modifications in subsequent generations (e.g., F2 and beyond) without continued exposure to the initial trigger. Recently, Takahashi et al. showed that insertion followed by removal of a CpG-free DNA segment at CpG island promoter regions causes DNA methylation and silencing of the surrounding loci, which are maintained over multiple generations¹⁵. Curiously, the induced DNA methylation was lost during germ cell development, but then reestablished during embryonic development by unknown mechanisms¹⁵; however, the methods used in the study could not fully rule out the influence of a small genetic change required for epimutation induction¹⁶. Moreover, other concerns, including off-target mutations during insertion of CpG-free DNA by CRISPR-Cas9 genome editing and colony selection, have been raised¹⁶. Epigenome editing using a ‘dead’ Cas9 (dCas9)^17,18, which does not introduce genetic changes and can rewrite the epigenetic information at a target locus, may be a direct method that can help solve this problem. Transmission of rewritten epigenetic information in the parental germline to the next generation, according to phenotype emergence, would directly demonstrate inter/transgenerational epigenetic inheritance.

Genomic imprinting is a process wherein a gene is differentially expressed depending on whether it has been inherited from the father or mother^19,20. An epimutation comprising DNA demethylation of the sperm-derived paternal allele in a differentially methylated region (DMR) of the imprinted gene, H19, (H19-DMR) is believed to cause Silver-Russell syndrome, which is characterized by intrauterine growth retardation due to Igf2 downregulation²¹. H19-DMR²² is located between the Igf2 and H19 imprinted genes^23,24 and regulates their expression. In somatic cells of healthy individuals, H19-DMR CpG dinucleotides, including CTCF-binding sites, are normally methylated on the paternal allele and unmethylated on the maternal allele (Fig. 1a)²²; this allele-specific CpG methylation upregulates paternal allele expression of Igf2 and downregulates that of H19. By contrast, in patients with Silver-Russell syndrome, demethylation of the paternal allele H19-DMR is expected to decrease the expression of Igf2 and increase that of H19 through epigenetic regulation of CTCF binding^25,26, resulting in fetal intrauterine growth retardation. During establishment of genomic imprinting in the male germline, H19-DMR CpGs are progressively hypermethylated through an endogenous de novo methylation mechanism until the pro-spermatogonia stage²⁷, while this region is unmethylated in oocytes (Fig. 1b). If the epimutation in H19-DMR occurring sperm is inherited, intrauterine growth retardation will be observed in offspring.

**Fig. 1: Generation of germline-specific epigenome-edited mice.**

In this study, we developed a germline-specific DNA methylation editing system and applied it to a genomic imprinting disorder disease model^28,29, to investigate inter/transgenerational inheritance of epimutations in sperm (Fig. 1b). To demonstrate this, we induced CpG demethylation in H19-DMR by germline-specific epigenome editing and investigated whether the epimutation introduced in sperm was transmitted to the next generation and led to phenotypic changes. Furthermore, we developed a targeted histone editing technique that can be used to reveal the roles of histone modifications in DNA methylation reestablishment.

Results

Germline-specific epigenome-edited mice show complete erasure of DNA methylation at H19-DMR in sperm

To induce CpG demethylation of H19-DMR, we constructed a vector that expresses epigenome editing factors driven by the Stra8 premeiotic-specific marker gene promoter (Fig. 2a)³⁰. In this system, dCas9 with a SunTag recruits the scFv-superfolder green fluorescent protein (sfGFP)-TET1CD fusion protein^31,32, which contains the catalytic domain (CD) of the ten-eleven translocation (TET) 1 hydroxylase, to the H19-DMR target locus, leading to targeted demethylation during spermatogenesis. By injecting the linearized vector into fertilized eggs, three transgenic (TG) strains were established from founder mice. A previous study showed that a transgene driven by the Stra8 promoter is activated only in the postnatal testis, and not in the ovary or prenatal gonad³³. RT-qPCR and immunohistochemistry analyses confirmed that epigenome editing factors were specifically expressed in adult testis, from spermatogonia to spermatocytes (Fig. 2b, cand Supplementary Fig. 1). As paternal imprinting in H19-DMR is established by the pro-spermatogonia stage²⁷, germline-specific epigenome editing is thought to occur after paternal imprinting establishment. H19-DMR CpG methylation levels were examined in sperm and MII oocytes from TG mice by bisulfite sequencing (Fig. 2d) and combined bisulfite restriction analysis (COBRA)³⁴ (Supplementary Fig. 2), and we found robust CpG demethylation in sperm from all TG strains. By contrast, little difference in CpG methylation levels between TG and wild-type (WT) sperm was detected on analysis of six potential off-target regions (Supplementary Fig. 3). As expected, H19-DMR was unmethylated in both TG and WT mother-derived oocytes (Fig. 2d). These data indicate that we successfully established germline-specific epigenome-edited TG mice (StrH19EpiTG), and strain 445-7 was used in subsequent experiments.

**Fig. 2: Complete DNA demethylation of *H19*-DMR in sperm derived from germline-specific epigenome-edited mice.**

Target-specific H19-DMR demethylation is partially inherited by the next generation and causes phenotypic changes

We next compared the offspring of TG mice with those of WT control mice. Offspring derived from TG mice were obtained by crossing heterozygous TG and WT mice (Fig. 3a), and those derived from TG fathers showed intrauterine growth retardation at birth (Fig. 3b, c). To exclude the possibility that the presence of the transgene might influence phenotypes, we also explored the phenotypes in offspring with and without TG alleles (TG and non-TG, respectively). Non-TG offspring derived from TG fathers exhibited a reduced birth weight (Fig. 3c), which is thought to be associated with inherited hypomethylation of H19-DMR from sperm. As expected, CpG methylation levels at the H19-DMR (CTCF binding sites m1–m4) in offspring of TG fathers were lower than those in offspring with WT fathers (Fig. 3d). Along with lowered methylation, offspring of TG fathers showed altered gene expression patterns, with increased H19 and decreased Igf2 levels (Fig. 3e). Non-TG offspring derived from TG fathers exhibited H19-DMR demethylation and altered gene expression at comparable levels to those in TG offspring (Fig. 3d, e). In addition to intrauterine growth retardation, other Silver-Russell syndrome-like phenotypes³⁵, including postnatal growth retardation, reduced food intake, and body asymmetry, were observed (Supplementary Fig. 4). By contrast, offspring derived from TG mothers did not differ from WT offspring in terms of birth weight or CpG methylation levels (Fig. 3b–d), consistent with the fact that H19-DMR is demethylated in oocytes (Fig. 2d). Thus, the developed system proved that induced loss of DNA methylation at the DMR in sperm was partially transmitted to the F1 offspring, which exhibited resulting phenotypes.

**Fig. 3: Target-specific demethylation of *H19*-DMR is inherited by the next generation and associated with phenotypic changes.**

We next investigated whether DNA methylation erasure in sperm was inherited transgenerationally (Supplementary Fig. 5a), and found that H19-DMR DNA hypomethylation was not inherited in sperm from F1 non-TG (Supplementary Fig. 5b, c) and their derivative F2 offspring (Supplementary Fig. 5d). These results indicate DMR DNA hypomethylation in F0 sperm is normalized during F1 germline cell development and not transgenerationally inherited by F2 offspring.

H19-DMR methylation is partially recovered from the 5’ end of the DMR during embryogenesis

Based on our experimental data, we surmised that DNA methylation was partially recovered in newborn mice, although it was completely erased in the sperm (Figs. 2d, 3d); we termed this phenomenon “intergenerational DNA methylation recovery”. The degree of DNA methylation recovery varied among individuals, and there was a strong positive correlation between birth weight and DNA methylation levels at the H19-DMR m3 and m4 sites and promoter regions (Supplementary Fig. 6). By contrast, DNA methylation levels at the 5’ end of the DMR (sites m1 and m2) were restored to almost WT levels (Supplementary Fig. 6), and there was no correlation between birth weight and DNA methylation levels at sites m1 and m2 (Supplementary Fig. 6). Further, time-course DNA methylation analysis across F1 development demonstrated that most DNA methylation at the 5’ end of the DMR (sites m1 and m2) was recovered before implantation, while that at 3’ end of the DMR (sites m3 and m4) was recovered after E7.5 (Fig. 4a). Furthermore, bisulfite sequencing analysis in sperm, early 1-cell zygotes (5 h post-fertilization), and morula stage embryos derived from TG fathers revealed that DNA methylation recovery at m1 and m2 sites had not yet occurred in zygotes, but had occurred by the morula stage (Fig. 4b). Allele-specific bisulfite sequencing analysis confirmed that the paternal allele underwent DNA methylation recovery (Fig. 4c). These data suggest the existence of an undetermined factor that mediates DNA methylation recovery in early preimplantation embryos.

**Fig. 4: Target-specific demethylation in sperm is partially recovered from the 5’ end of the DMR during neonatal development.**

H3K9me3 is required for DNA methylation recovery

H3K9me3 is a repressive histone mark associated with heterochromatin and gene repression³⁶, and known to co-occur with DNA methylation at the DMRs of imprinted genes³⁷. Further, genome-wide H3K9me3 loss leads to a reduction of DNA methylation at DMRs in preimplantation embryos³⁸. Nevertheless, although these studies suggest that H3K9me3 is required for DNA methylation maintenance, it is unknown whether H3K9me3 guides de novo DNA methylation in early embryos. According to public ChIP-seq data from gametes^39,40, H3K9me3 is rarely enriched around the H19-DMR in either sperm or oocytes (Fig. 5a). To determine when H3K9me3 is deposited at the H19-DMR after fertilization, we applied an ultra-sensitive chromatin immunoprecipitation sequencing (ChIP-Seq) method, termed carrier DNA-assisted ChIP-Seq (CATCH-Seq)^41,42, Interestingly, H3K9me3 CATCH-seq analysis in early 1-cell zygotes (5 h post-fertilization) and morula embryos revealed that H3K9me3 could already be detected at the paternal allele of H19-DMR in early zygotes, particularly at the 5’ end of the DMR (sites m1 and m2) (Fig. 5a). H3K9me3 was further accumulated by the morula stage (Fig. 5a).

**Fig. 5: Target-specific removal of H3K9me3 around *H19*-DMR.**

To examine the role of H3K9me3 in DNA methylation recovery, we attempted to erase H3K9me3 at the H19-DMR in early embryos. To achieve this, we conducted target-specific histone editing using an H3K9me3 demethylase, KDM4D⁴³. Mixtures of dCas9-SunTag mRNA, scFv-GFP-Kdm4d mRNA, and two guide RNAs (gRNAs) complementary to sequence upstream of H19-DMR were injected into zygotes derived from TG fathers (Fig. 5b). Since the GFP-labeled epigenome editing factor (scFv-GFP-Kdm4d) was strongly expressed for at least 3 days until the morula stage (Supplementary Fig. 7), the H3K9me3 status of epigenome-edited embryos was examined at this stage. H3K9me3 CATCH-seq analysis of TG sperm-derived, non-injected morula embryos showed that H3K9me3 was detected around H19-DMR and upstream regions (Fig. 5c). We noted that levels of H3K9me3 at this locus in TG sperm-derived embryos were lower than those in WT embryos (see Discussion) (Fig. 5c). Nevertheless, when KDM4D was recruited to the upstream of H19-DMR by target-specific histone editing, H3K9me3 around the DMR was almost completely lost over an extensive region (approximately 10 kb). This extensive effect beyond the target sites is consistent with a recent report showing that an edited region per gRNA is comparatively broad in histone modification-edited embryonic stem cells⁴⁴. By contrast, H3K9me3 was retained in DMRs of other imprinted genes (Igf2r, Rasgrf1) (Fig. 5d). Genome-wide comparative analysis demonstrated that H3K9me3 was lost at only 18 regions, including H19-DMR, in epigenome-targeted morula embryos (Fig. 5eand Supplementary Data 1). Thus, H3K9 demethylation by dCas9-SunTag histone editing did not affect H3K9me3 globally.

Next, these manipulated embryos were developed to embryonic stage E11.5, and their H19-DMR DNA methylation levels were examined (Fig. 6a). Importantly, target-specific H3K9me3 removal using dCas9-SunTag-Kdm4d revealed that DNA methylation recovery at H19-DMR is inhibited (Fig. 6b). Conversely, DNA methylation recovery was observed when a KDM4D mutant (H189A, lacking enzymatic activity)⁴⁵ or another histone demethylase (KDM6B, H3K27 demethylase)⁴⁶ was recruited (Fig. 6b). The extent of recovery in these negative controls was less than that in the non-treated control group (Fig. 6b), which could be due to steric hindrance of the epigenome editor complexes against DNA methyltransferases. In partial support of this hypothesis, a greater degree of recovery was observed when KDM4D was recruited to a different target locus, Rosa26⁴⁷ (Fig. 6b). Finally, dissection of E15.5 and E18.5 embryos derived from TG sperm-derived zygotes injected or not with target-specific KDM4D revealed that H3K9me3 removal results in more severe intrauterine growth retardation than in the non-injected controls (Fig. 6c). These data indicate that H3K9me3 around H19-DMR is involved in DNA methylation recovery during embryogenesis and is physiologically relevant to the offspring’s phenotype.

**Fig. 6: H3K9me3 is required for DNA methylation recovery.**

To further confirm the role of H3K9me3 in DNA methylation recovery, we also removed H3K9me3 globally in zygotes by injecting Kdm4d mRNA⁴⁵. Similar to target-specific removal, global loss of H3K9me3 during early preimplantation development inhibited DNA methylation recovery, whereas control embryos injected with catalytic point-mutant Kdm4d mRNA, Egfp mRNA, or buffer alone led to DNA methylation recovery (Supplementary Fig. 8). Regardless of global or target-specific editing, the inhibitory effect of H3K9me3 removal on DNA methylation recovery was more prominent at the m1 and m2 sites than that at m3/m4, possibly because DNA methylation recovery occurs more robustly at the 5’ end of the DMR. Thus, both target-specific and global removal experiments confirmed that H3K9me3 is required for DNA methylation recovery at H19-DMR.

Discussion

The germline-specific epigenome editing system developed in this study enabled us to validate the inter/transgenerational inheritance theory directly, because it involves editing of the germline epigenome alone and not the genome. Our system can be used to introduce epimutations at various loci, facilitating the study of the effects of an inter/transgenerational epimutations on offspring. Using this system, we demonstrate that artificial DNA demethylation of H19-DMR in F0 sperm can be partially transmitted to the F1 generation and mediate offspring phenotypes. Further, we used our system to demonstrate that DNA demethylation of H19-DMR is reprogrammed and normalized in F1 germline cells, and not transgenerationally inherited by F2 offspring. There have been several previous reports that epigenetic reprogramming of sperm induced by abnormal nutritional conditions or stress can be transmitted to offspring^{3,4,5,6,7,8,9}; however, these studies were based on circumstantial evidence, and the germline-specific epigenome editing system we have developed could contribute to assessing whether there are causal relationships in these contexts.

Notably, despite successful removal of DNA methylation in sperm, we found that it was partially recovered from the 5’ end of H19-DMR (sites m1 and m2) after fertilization. We propose that H3K9me3 mediates the DNA methylation recovery that occurs after fertilization (Fig. 7). (1) In WT embryos, the paternal allele of H19-DMR is marked with H3K9me3 immediately after fertilization, which accumulates further by the morula stage. The coexistence of DNA methylation and H3K9me3 marks likely ensure robust genomic imprinting throughout development. (2) In TG father-derived embryos, the DNA hypomethyled state in sperm is transmitted to zygotes. DNA methylation is then partially restored by the morula stage, until which H3K9me3 is deposited. The level of H3K9me3 accumulation in these embryos was slightly lower than that detected in WT embryos, suggesting that there is also a mechanism by which DNA methylation recruits enzymes that mediate H3K9me3. Interestingly, the extent of DNA methylation recovery varied among F1 offspring, and there was a strong correlation between birth weight and DNA methylation level at the 3’ end of H19-DMR (sites m3 and m4) and in promoter regions. This variation could be dependent on the amount of H3K9me3, which may cause DNA methylation variation in post-implantation embryos. (3) In addition to the absence of DNA methylation, H3K9me3 erasure blocks DNA methylation recovery, resulting in persistence of the hypomethylated DNA state in postimplantation embryos.

**Fig. 7: Schematic model illustration of inheritance and recovery of epigenetic marks at the paternal *H19*-DMR.**

Target-specific and global H3K9me3 removal experiments in zygotes revealed that H3K9me3 mediates DNA methylation recovery at the H19-DMR. By contrast, H3K9me3 is nearly absent at the H19-DMR in sperm, suggesting that H3K9me3 is not the primary intergenerational memory directly transmitted across generations, but rather that some other factor(s) upstream of H3K9me3 is involved. Identification of the upstream factors, which could be genetic or epigenetic, awaits further investigations. Future studies will be directed toward uncovering the mechanism of these upstream signals to further advance our understanding of epigenetic inheritance. Candidates include those factors known to participate in de novo H3K9me3 and subsequent DNA methylation during transposable element silencing⁴⁸: Krüppel-associated box (KRAB) domain-containing zinc-finger proteins recognize transposable elements and recruit the cofactors KAP1/TRIM28, SETDB1 (H3K9 methyltransferase), and HP1, leading to formation of repressive heterochromatin and de novo DNA methylation by DNMT1, DNMT3A/B, and DNMT3L^{49,50,51,52,53,54,55}. These factors may contribute to de novo H3K9me3 formation and DNA methylation in preimplantation embryos. Notably, Tanimoto et al. reported that a transgene encoding H19-DMR is de novo methylated after fertilization when it is paternally transmitted⁵⁶. Although the underlying mechanism remains unknown, these authors showed that an 118 bp sequence upstream of H19-DMR is required for de novo DNA methylation⁵⁷. Importantly, the 118 bp sequence is located in the region where H3K9me3 is deposited in zygotes. It will be interesting to examine what factors bind to this sequence, which will assist in the identification of key molecules responsible for H3K9me3 deposition and DNA methylation recovery in early embryos.

It remains unclear how H3K9me3 guides de novo DNA methylation at H19-DMR during embryogenesis. Burton et al. reported that de novo H3K9me3 catalyzed by SUV39H2 (H3K9 methyltransferase) occurs in the paternal genome immediately after fertilization⁵⁸, which is consistent with our observation that H3K9me3 emerges on the paternal allele of H19-DMR in early zygotes. While sperm-derived genomes undergo widespread DNA demethylation after fertilization, Richard et al. reported that some regions, including several dozen CpG islands, are paradoxically de novo methylated by the 2-cell stage in a maternal DNMT3A-dependent manner⁵⁹. Our study suggests that de novo H3K9me3 in zygotes is a prerequisite for the subsequent de novo DNA methylation, although the degree to which this mechanism is shared at loci other than the H19-DMR is currently unknown. Although H3K9me3 is generally recognized for its role in supporting DNA methylation, further research is necessary to determine whether this H3K9me3-mediated DNA methylation recovery is a general phenomenon or specific to imprinted loci.

DNA methylation recovery occurred specifically at the paternal, but not the maternal, DMR after fertilization, which raises the question of how the maternal allele resists de novo DNA methylation. The most plausible responsible factor is CTCF^25,26, which binds to the maternal DMR before fertilization. Indeed, DNA methylation levels at the maternal DMR are increased by CTCF inhibition or knockdown in oocytes^60,61, suggesting that the CTCF-bound maternal DMR is protected from DNA methylation. In the offspring derived from TG fathers, DNA methylation at the DMR was gradually restored pre- and post-implantation. During this time, CTCF could bind to the hypomethylated paternal DMR and inhibit DNA methylation recovery. Indeed, our previous study demonstrated that stable DNA hypomethylation of the paternal DMR by continuous expression of epigenome editing factors allows CTCF binding to the paternal DMR in newborn mice³².

Interestingly, hypomethylation of the H19-DMR and other imprinted loci has been observed in sperm from infertile patients^62,63. The causal relationship between DNA methylation and sperm phenotypes could be directly elucidated in future studies using the sperm-specific epigenome editing system presented here. Further, various DNA methylation mutations occur during spermatogenesis under specific circumstances, such as aging⁶⁴, which could be repaired after fertilization through the H3K9me3-mediated mechanism identified in this investigation. Advances in DNA sequencing technologies have led to the generation of an enormous number of candidate disease-causing epigenetic changes associated with various diseases, including genomic imprinting disorders, cancers, obesity, diabetes, and autism. Our epigenome editing system is a powerful tool that can be applied to determine how epimutations in sperm are transmitted or recovered across generations, as well as how they affect phenotypes.

Methods

Animals

B6D2F1 mice were purchased from CLEA Japan (Kawasaki, Japan). ICR, C57BL/6NCrl (B6N) mice were purchased from Charles River Japan (Yokohama, Japan). PWK/PhJ (PWK) mice (RBRC00213, RIKEN BioResource Research Center) originated from 003715 (The Jackson Laboratory). The mouse facility was kept at 21–25 °C and 40–60% humidity with a 12 h light–dark cycle. Animals had ad libitum access to food and water. All animal procedures were approved by the Animal Care and Experimentation Committee at Gunma University (No. 15-045) and carried out in accordance with approved guidelines.

Vector construction for germline-specific demethylation

The all-in-one epigenome editing vector, including dCas9 fused with five copies of GCN4 and an anti-GCN4 peptide antibody (scFv)-sfGFP-TET1CD fusion protein (pPlatTET-gRNA2, Addgene plasmid 82559), was previously reported by this laboratory³¹. An all-in-one epigenome editing vector, including nine gRNAs (Supplementary Data 2) targeting H19-DMR (pPlatTET-gRNA2-H19DMRx9), was also constructed as previously reported³². In this study, the CAG promoter of the pPlatTET-gRNA2-H19DMRx9 vector was replaced with the promoter of the Stra8 gene, which is specifically expressed during gametogenesis (pStrPlatTET-gRNA2-H19DMRx9). The vector was linearized by digestion with ApaLI before microinjection.

Generation of StrH19EpiTG mice

B6D2F1 female mice aged 8–10 weeks were induced to superovulate by injecting 7.5 units of pregnant mare’s serum gonadotropin (SEROTROPIN; ASKA Pharmaceutical, Tokyo, Japan), followed 48 h later by 7.5 units of human chorionic gonadotropin (hCG; GONATROPIN, ASKA Pharmaceutical). After administration of hCG, females were mated with B6D2F1 males and zygotes isolated from the oviduct 21 h later. After treatment with M2 medium (Sigma-Aldrich, St. Louis, MO, USA) supplemented with 0.1% hyaluronidase (Sigma-Aldrich) for 3 minutes, zygotes were washed with M2 medium and then transferred to drops of M16 medium (Sigma-Aldrich) supplemented with penicillin and streptomycin at 37 °C. Pronuclear injection was performed at 24–27 h post-hCG injection³². Linearized pStrPlatTET-gRNA2-H19DMRx9 vector (35 ng/μl) was injected into the pronuclei of zygotes. Injected embryos were cultured in M16 medium at 37 °C under 5% CO₂ in air. The next day, embryos that had developed to the 2-cell stage were transferred into the ampulla of the oviduct (20–25 embryos per oviduct) of pseudopregnant ICR females. For vector integration analysis, genomic DNA was extracted from tail tips of obtained mice using a DNA extraction kit (DirectPCR Lysis Reagent, Mouse Tail; Viagenbiotech, CA, USA). PCR analysis using the primer set for dCas9 (Supplementary Data 2) demonstrated that 9 of 41 founder mice possessed the transgene. Three reproducible TG lines (441-2, 445-7, and 445-14) and their sub-lines were finally established; the 445-7 line was used in the present study and maintained by backcrossing WT B6D2F1 females with TG males. A TG male mouse that had been backcrossed twice with a PWK strain was used for allele-specific analysis.

Collection of sperm, oocytes, embryos, and newborn mice

Sperm (Fig. 2) were isolated from the cauda epididymis of TG or WT mice aged 10–21 weeks by dissecting tissue in 500 μl modified Human Tubal Fluid (mHTF) medium (Kyudo, Saga, Japan). Sperm were allowed to swim up for 1 h at 37 °C under an atmosphere of 5% CO₂ in air. The upper 200 μl fraction was collected, and the collected sperm were then resuspended and incubated with hypotonic buffer to eliminate red blood cells for 3 min at room temperature (RT), followed by washing with phosphate-buffered saline (PBS). Newborn (P0) mice (Fig. 3) were generated by natural mating of TG or WT (B6D2F1) parents. Blastocyst, E7.5, and E11.5 embryos (Fig. 4) were generated by in vitro fertilization followed by embryo transfer. Briefly, sperm were obtained from the caudal epididymis of adult TG or WT male mice, and capacitated by 1 h incubation in mHTF medium. Cumulus–oocyte complexes were harvested from B6D2F1 female mice aged 8–10 weeks 15–17 h after hCG administration, and then inseminated with activated sperm in mHTF medium. At 4 h post-insemination, zygotes with two pronuclei were subjected to in vitro culture in KSOM medium (ARC Resource, Kumamoto, Japan) at 37 °C under 5% CO₂ in air. The next day, a proportion of embryos that had developed to the 2-cell stage were transferred into the ampulla of the oviduct of pseudopregnant ICR females and harvested at E7.5 and E11.5. Wild-type preimplantation embryos (Fig. 5a) were generated by in vitro fertilization using PWK sperm and B6N oocytes^65,66. Preparation of histone-edited embryos (Figs. 5b–e,6) is described below.

DNA methylation analysis

Sperm pellets were re-suspended in RSB (10 mM NaCl, 10 mM Tris pH 7.5, and 25 mM EDTA) containing 2% sodium dodecyl sulfate (SDS), 2% 2-mercaptoethanol, and 1 mg/ml Proteinase K. Embryos were re-suspended in TNES (10 mM Tris-HCl pH 7.5, 10 mM NaCl, 25 mM EDTA, and 1% SDS) containing 0.2 mg/ml Proteinase K. After incubation overnight at 55 °C, DNA was isolated by phenol/chloroform extraction followed by ethanol precipitation. Genomic DNA from whole bodies of newborn mice was extracted using an AllPrep DNA/RNA Mini Kit (QIAGEN, Venlo, The Netherlands). Purified DNA samples (500 ng) were processed with an Epitect Plus DNA Bisulfite Kit (QIAGEN) according to the manufacturer’s instructions. Modified DNA was amplified using TaKaRa Taq (TaKaRa, Kusatsu, Japan) and the PCR primers described in Supplementary Data 2. Percentages of demethylated CpG sites were determined by COBRA³⁴. Amplified fragments were cleaved with restriction enzymes (Supplementary Data 2) with recognition sequences located in CpG sites. PCR products were separated and quantified using capillary and microchip electrophoresis (MCE-202 MultiNA, Shimadzu, Kyoto, Japan). Methylation level was calculated as the percentage of cleaved PCR products (mV･μm) among total PCR products (mV･μm). Alternatively, PCR products were cloned using the TOPO TA Cloning Kit (Invitrogen, Carlsbad, CA, USA). Around 24 positive clones were sequenced for each sample using the BigDye terminator method (ABI PRISM 3500 x L, Applied Biosystems, Foster City, CA, USA). Alignment, visualization, and quantification of bisulfite sequence data were performed using QUantification tool for Methylation Analysis (QUMA; http://quma.cdb.riken.jp/top/index.html). Single-nucleotide polymorphisms (SNPs) of PWK and B6N strains were utilized for allele-specific DNA methylation analysis.

Bisulfite amplicon sequencing analysis

For comprehensive CpG methylation analysis of H19-DMR, genomic DNA samples treated with the Epitect Plus DNA Bisulfite Kit (QIAGEN) were amplified using specific primer pairs (Supplementary Data 2). A fragment library was prepared from pooled PCR products using the NEBNext Ultra Ⅱ DNA Library Prep Kit for Illumina (NEB). Briefly, 100 ng of PCR product was sheared using the Covaris^® S220 System (Covaris Inc, MA) at 5 °C for 2 cycles, treatment time 150 sec, with duty factor 10%, and 200 cycles/burst. To generate a fragment library, NEBNext Adapters (NEB) were ligated to both ends of the DNA. Barcoded libraries were amplified by PCR for 5 cycles using NEBNext Multiplex Oligos for Illumina (NEB). Prepared libraries were then sequenced on the Illumina NovaSeq 6000 platform (Illumina, San Diego, CA). Raw sequence data generated in FASTQ format were imported into CLC Genomics Workbench 24.0.1 (QIAGEN), trimmed using the Trim reads 3.0 tool, and mapped to 15 kb reference sequence regions (NC_000073, Mus musculus strain C57BL/6 J chromosome 7, GRCm38.p6 C57BL/6 J) using the Map Bisulfite Reads to Reference tool. For CpG methylation analysis, trimmed libraries were mapped to the reference sequence described above using Map Bisulfite Reads to Reference, and 5-mC percentages calculated using Call Methylation levels 1.4.

RT-qPCR

Total RNA was isolated from the whole bodies of newborn mice using an AllPrep DNA/RNA Mini Kit (QIAGEN). Isolated RNA (8.5 μl, 2 μg) was treated with DNase I (50 μ ml⁻¹) in a total volume of 10 μl at 37 °C for 20 min. DNase I was inactivated by adding 0.8 μl 25 mM EDTA and incubating samples at 75 °C for 10 min. cDNA was produced from each RNA sample using random primers (Invitrogen, Carlsbad, CA) and SuperScript II (Invitrogen), according to the manufacturer’s instructions. Total RNA extraction and cDNA synthesis from zygotes and preimplantation embryos were performed using the superPrep Cell Lysis & RT Kit for qPCR (TOYOBO, Osaka, Japan). Reverse transcription reactions were diluted 10-fold with water before qPCR. Gene expression levels of Igf2, H19, and dCas9 were measured using a LightCycler 96 (Roche) with TB Green Premix Ex Taq II (TakaRa), according to the manufacturer’s instructions. Expression levels were normalized against those of Gapdh. Primer sequences are listed in Supplementary Data 2.

Immunohistochemistry

Testes specimens were fixed in 4% paraformaldehyde diluted in PBS, embedded in paraffin, and 4 μm sections prepared. Sections were deparaffinized, autoclaved (121 °C, 5 min) in 10 mM sodium citrate buffer pH 6.0 for antigen retrieval, and treated with methanol containing 0.3% H₂O₂ for 10 min. After blocking with 3% bovine serum albumin diluted in PBS containing Tween 20, an anti-GFP antibody (1:500 dilution, ab290, Lot No. GR3184825-1; Abcam, Cambridge, UK) was applied and incubated overnight at 4 °C. Sections were reacted with Histofine Simple Stain MAX-PO (R) (Nichirei, Tokyo, Japan) for 30 min at room temperature, signals visualized using 3-3’-diaminobenzidine, and samples counterstained with hematoxylin.

Preparation of RNA

Kdm4d and its enzymatic mutant, Kdm4dMut (H189A)⁴⁵, Kdm5b, EGFP, dCas9, dCas9-SunTag (L15aa), scFv-Kdm4d, scFv-Kdm4dMut, and scFv-Kdm6b were amplified from template plasmid using Q5 Hot Start High-Fidelity DNA Polymerase (New England BioLabs (NEB), MA, USA) and primer sets for in vitro transcription (Supplementary Data 2). Amplified PCR products were gel-purified and subjected to in vitro transcription with an mMESSAGE mMACHINE T7 ULTRA kit (Life Technologies). In vitro transcribed RNA was eluted into RNase-free water, and the quality was checked by gel electrophoresis. Single guide RNAs (sgRNAs) for H19-DMR and Rosa26 (Supplementary Data 2) were purchased from Integrated DNA Technologies (IDT).

Histone editing by RNA injection

Zygotes derived from TG fathers were generated by in vitro fertilization. Sperm were obtained from adult TG male mouse caudal epididymis and capacitated by 1 h incubation in mHTF medium (Kyudo, Saga, Japan). Cumulus–oocyte complexes were harvested from B6D2F1 female mice 15–17 h after hCG administration, and then inseminated with activated sperm in mHTF medium. At 4 h postinsemination, zygotes with two pronuclei were subjected to cytoplasmic injection of RNA⁶⁷ for histone editing. For global histone editing, in vitro transcribed Kdm4d/Kdm4dMut (1300 ng/μl) and EGFP (1300 ng/μl) mRNA samples were injected into the cytoplasm of zygotes in M2 medium. For target-specific histone editing, in vitro transcribed dCas9-SunTag mRNA (239 ng/μl), scFv-GFP-Kdm4d/Kdm4dMut mRNA (161 ng/μl), or scFv-GFP-Kdm6b mRNA (172 ng/μl), and two sgRNAs targeting sites upstream of H19-DMR (97 ng/μl each) or an sgRNA targeting Rosa26 (194 ng/μl) were injected into the cytoplasm of zygotes in M2 medium. Injected embryos were cultured in KSOM medium at 37 °C under 5% CO₂ in air. The next day, a proportion of the embryos that had developed to the 2-cell stage were transferred into the ampulla of the oviduct of pseudopregnant ICR females and harvested at E11.5–E18.5 for DNA methylation analyses or weight measurement.

H3K9me3 Carrier Assisted ChIP-seq (CATCH-seq)

H3K9me3 CATCH-seq libraries were prepared as previously described^41,42, with some modifications. Zygotes (n = 200–250) and morula embryos (n = 30–45) were permeabilized using Nuclei EZ lysis buffer supplemented with 0.1% Triton X-100, 0.1% deoxycholate, complete EDTA-free protease inhibitor cocktail and 1 mM phenylmethanesulfonyl fluoride on ice. Chromatin was fragmented in situ using 2 μ/μl MNase (M0247S, NEB) at 37 °C for 7.5 min; the reaction was stopped by adding 1/10 volume of 100 mM EDTA and diluted with freshly prepared immunoprecipitation buffer. Then, 30 ng of annealed I-SceI carrier DNA was added to each sample. The forward and reverse strand sequences of the carrier DNA were as follows: /5AmMC6/Gtagggataacagggtaattagggataacagggtaattagggataacagggtaattagggataacagggtaattagggataacagggtaattagggataacagggtaat*c/3AmMO/ and /5AmMC6/Gattaccctgttatccctaattaccctgttatccctaattaccctgttatccctaattaccctgttatccctaattaccctgttatccctaattaccctgttatcccta*c/3AmMO/, respectively, where asterisks represent phosphorothioate bonds. Oligos were synthesized by IDT. For each immunoprecipitation reaction, 1.0 μl of rabbit polyclonal anti-H3K9me3 antibody (Active Motif, #39162, Lot No. 30220003) conjugated to precleared Dynabeads Protein A and G mixture was used. After immunoprecipitation at 4 °C overnight, chromatin-Dynabeads were washed in low and high-salt wash buffers, and chromatin eluted in freshly prepared ChIP elution buffer at 65 °C for 1 h. DNA was recovered by phenol-chloroform extraction followed by ethanol precipitation. Adapter ligation was conducted using an NEBNext Ultra II DNA Library Prep Kit for Illumina (E7645, NEB) at half scale relative to the manufacturer’s instructions, and libraries purified using 1.8× SPRIselect beads (B23318, Beckman Colter). DNA was amplified using KAPA Hifi 2 × master PCR mix (KK2605) with indexing primers for 13–14 PCR cycles. After purification with 0.9 × SPRIselect beads, samples were digested with I-SceI (5 μ/μl, NEB, R0694) at 37 °C for 2 h, followed by heat inactivation at 65 °C for 20 min, then purified using 0.9 × SPRIselect beads. A second amplification was not performed. Libraries were sequenced on the NextSeq500 (Illumina, CA, USA) or NovaSeq (Illumina) platforms.

CATCH-seq data analysis

Allele-specific analysis of hybrid embryos (B6N × PWK) was performed as described previously⁶⁶, with some modifications. Reads were trimmed using fastp⁶⁸ and mapped using bowtie2⁶⁹ with the options “-N 1”. The modified mm10 reference genome, in which PWK strain single-nucleotide polymorphisms were masked as N, was used for mapping. Multiply mapped reads were retained for downstream analysis in this study. PCR duplicates were removed using Picard MarkDuplicates (https://broadinstitute.github.io/picard/). SNPsplit⁷⁰ was used to assign reads to their parental origin. For analysis of previous data^39,40, reads were aligned to the original mm10 reference genome, and subsequent processes performed as described above. For analysis of TG embryo CATCH-seq data, reads were extended to 250 bp and processed as single-end reads to minimize the difference in sequencing conditions among samples, and subsequent processes were performed as described above. To visualize H3K9me3 enrichment, reads from biological replicates were pooled, and bigwig files generated using bamCoverage from deepTools⁷¹, with the options “--normalizeUsing RPKM --binSize 50”. Scatter plots showed a strong correlation between biological replicates of H3K9me3 CATCH-seq (Supplementary Fig. 9).

To confirm the specificity of epigenome editing by KDM4D, biological replicates of WT_control were pooled for peak calling using bdgpeakcall from macs2⁷² with options “-l 3000 -c 4”. H3K9me3 enrichment peaks were compared using DiffBind⁷³; differences of p < 0.05 and |log₂ fold change | > 2 were considered significant changes induced by KDM4D (Fig. 5eand Supplementary Data 1).

Micro-CT and 3D reconstruction

Eight-week-old mice were scanned using a computed tomography instrument (DELPetμCT100, DELBio, Taoyuan County, Taiwan (R.O.C.)). Three-dimensional cross sections were generated with a resolution of one cross-section per 22.5 μm. Images were transformed into DICOM format and landmarks placed within the 3D representation at the endpoints of limb bones in the forelimb (humerus and ulna) and hindlimb (femur and tibia) using VivoQuant2020 software (Invicro, MA, USA). Linear measurements were obtained for each pair of landmarks.

Statistical analysis

Box plots show the median (center line), the 25th and 75th percentiles (box edges), and the minimum and maximum values (whiskers). Data shown in bar and line graphs are presented as mean and standard deviation (s.d.) values. DNA methylation, mRNA expression, body weight, and food intake were analyzed by Welch’s t test (two-tailed) for pairwise comparisons or one-way analysis of variance (ANOVA) followed by Tukey’s post-hoc HSD test for multiple comparisons. To analyze correlations between these variables, Pearson’s correlation coefficient (r) values were calculated. Fisher’s exact probability test was used to compare the frequency of body asymmetry. A p-value < 0.05 was considered significant.

Reporting summary

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

Data availability

The accession number for the raw sequencing data reported in this paper is DDBJ: PRJDB19200. Detailed sample information is shown in Supplementary Data 3. Sperm and oocyte ChIP-seq datasets were obtained from previous publications (DDBJ: DRA006537 and GSE97778)^39,40. The source data and exact p-values underlying Figs. 2c, 3c–e, 4a, b, 6b, c, and Supplementary Figs. 1b, 2c, 4a–c, 5b, d, 6, and 8 are provided as a Source Data file. Source data are provided in this paper.

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Acknowledgements

We thank Ms. Junko Shima, Ms. Yuko Okazaki, Ms. Emi Hosoya, Ms. Nanase Yamazaki, Ms. Rie Fukuda, Ms. Marie Ishikawa and Ms. Eriko Suetomo (Gunma University) for technical support with PCR and bisulfite sequencing analyses, and Dr. Yuhkoh Satouh (Gunma University) for evaluation of testis sections. We also thank Ms. Makiko Nakagawa, Ms. Yuko Nakatani, and Dr. Kenji Watanabe (Yamaguchi University) and Ms. Mami Kumon (RIKEN) for technical support with next-generation sequencing analyses. We appreciate the technical support provided by the Yamaguchi University Science Research Center. This work was supported by funding from the Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Numbers 24H00679, Scientific Research (A) and 23K23802, Scientific Research (B) to T.H., 23K05727, Scientific Research (C) to S.M., and 24K02582, Scientific Research (B) to I.H.; the Japan Agency for Medical Research and Development (AMED) under Grant Number 22gm6310009h0004 to S.M.; the Platform Project for Supporting Drug Discovery and Life Science Research (Basis for Supporting Innovative Drug Discovery and Life Science Research (BINDS)) from AMED under Grant Numbers JP21am0101120 and JP24ama121049 to I.H.; the Tokyo Biochemical Research Foundation to I.H.; the Takeda Science Foundation to I.H.; the Practical Research Project for Rare/Intractable Diseases from AMED under Grant Number 21ek0109489h0002 to I.H.; the Uehara Memorial Foundation to I.H.; the Joint Usage/Research Center for Developmental Medicine, IMEG, Kumamoto University to T.H.; the Life Science Institute, Inc. (LSII) to I.H.; Grant-in-Aid for Transformative Research Areas (A) under Grant Number 25H01355 to A.I.; JST FOREST Program, Grant Number JPMJFR2335 to A.I.; and the JSPS postdoctoral fellowship PD, Grant Number 24KJ0236 to H.F.

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Authors and Affiliations

Laboratory of Genome Science, Biosignal Genome Resource Center, Institute for Molecular and Cellular Regulation, Gunma University, Maebashi, Japan
Takuro Horii, Sumiyo Morita, Ryosuke Kobayashi, Mika Kimura & Izuho Hatada
Department of Medical Cell Biology, Institute of Molecular Embryology and Genetics, Kumamoto University, Kumamoto, Japan
Shinjiro Hino, Yuko Hino & Mitsuyoshi Nakao
Laboratory for Epigenome Inheritance, RIKEN Center for Integrative Medical Sciences, Yokohama, Japan
Hiroto S. Fukushima & Azusa Inoue
Institute of Gene Research, Yamaguchi University Science Research Center, Yamaguchi, Japan
Yoichi Mizukami
Tokyo Metropolitan University, Hachioji, Japan
Azusa Inoue
Viral Vector Core, Gunma University Initiative for Advanced Research (GIAR), Maebashi, Japan
Izuho Hatada

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T.H. and I.H. designed the experiments. T.H., S.M., S.H., Y.H., H.F., R.K., M.K., M.N., Y.M., and A.I. performed the experiments and analyzed the data. T.H., S.M., H.F., R.K., A.I., and I.H. wrote the manuscript. I.H. supervised the project. All authors read and approved the final manuscript.

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Correspondence to Takuro Horii or Izuho Hatada.

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Competing interests

The authors declare that a patent application related to this work has been filed by Gunma University (Japanese Patent Application No. 2023-522326; currently under examination). The inventors are I.H., T.H., and S.M. The patent pertains to methods for germline epigenome editing. This research was funded by the LSII, Inc. The funder provided financial support to I.H. and was involved in the data analysis presented in Supplementary Fig. 4; however, the decision to publish and the preparation of the manuscript were made independently by the authors. S.H., Y.H., H.F., R.K., M.K., M.N., Y.M., and A.I. declare no competing interests.

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Horii, T., Morita, S., Hino, S. et al. Germline epigenome editing identifies H3K9me3 as a mediator of intergenerational DNA methylation recovery in mice. Nat Commun 16, 11200 (2025). https://doi.org/10.1038/s41467-025-67488-9

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Received: 19 December 2024
Accepted: 02 December 2025
Published: 25 December 2025
Version of record: 25 December 2025
DOI: https://doi.org/10.1038/s41467-025-67488-9