The timing of pronuclear transfer critically affects the developmental competence and quality of embryos

Abstract

Pronuclear transfer has been successfully used in human-assisted reproduction to suppress the adverse effects of a defective oocyte cytoplasm or to bypass an idiopathic developmental arrest. However, the effects of the initial parental genome remodelling in a defective cytoplasm on the subsequent development after pronucleus transfer have not been systematically studied. By performing pronuclear transfer in pre-replication and post-replication mouse embryos, we show that the timing of the procedure plays a critical role. Although apparently morphologically normal blastocysts were obtained in both pre- and post-replication pronuclear transfer groups, post-replication pronuclear transfer led to a decrease in developmental competence and profound changes in embryonic gene expression. By inhibiting the replication in the abnormal cytoplasm before pronuclear transfer into a healthy cytoplasm, the developmental potential of embryos could be largely restored. This shows that the conditions under which the first embryonic replication occurs strongly influence developmental potential. Although pronuclear transfer is the method of choice for mitigating the impact of a faulty oocyte cytoplasm on early development, our results show that the timing of this intervention should be restricted to the pre-replication phase.

Introduction

Pronuclear (PN) transfer, also called cytoplasmic donation/replacement therapy, is used in several countries in human-assisted reproduction, including the UK, Mexico, and Ukraine, and has been successfully implemented to mitigate the effects of a defective cytoplasm (Palacios-González and Medina-Arellano, 2017; Ishii and Hibino, 2018). During the PN transfer procedure, the parental genomes are first converted into parental pronuclei upon fertilization. These can then be safely manipulated and transferred into a recipient cytoplasm. However, this process means that the initial phases of parental genome remodelling and pronucleus assembly take place in a defective cytoplasm.

PN transfer is most frequently used in the treatment of mitochondrial diseases where the maternal/oocyte mitochondria carry a specific detrimental mutation(s). It has also been used to treat conditions caused by other cytoplasmic factors, which might or might not be mitochondria-related. These include an insufficient ATP content or an idiopathic developmental arrest of embryos obtained after fertilization of a particular patient’s oocytes (Cohen et al., 1997; Craven et al., 2010).

Although we may presume that successful development requires a pristine sequence of events, which lead to the formation of fully functional parental genomes, it is unclear to what extent the healthy recipient cytoplast can suppress or even reverse the unfavourable conditions set during the first hours post-fertilization. These questions are particularly hard to answer in human patients for legal reasons and because of the limited number of oocytes that can be obtained from a single patient. To obtain more insight and to investigate the prerequisites of the PN transfer procedure with respect to the developmental rates of the resulting embryos, in this study, we tested whether the timing of PN transfer represents an important parameter by using a mouse model of idiopathic developmental arrest.

Materials and methods

Unless stated otherwise, all reagents were purchased from Sigma-Aldrich, now part of Merck, Prague, Czech Republic. All experiments were repeated at least three times.

Animals

All animals in this study were purchased from Janvier Labs (Le Genest-Saint-Isle, France) and housed under standard conditions: temperature was maintained at 21–22°C with a 12-h light cycle; water and food were provided ad libitum. Ethical approval of the study was obtained from the Animal Care and Use Committee of the Institute of Experimental Medicine and The Academy of Sciences of the Czech Republic (approval number: 28/2019).

Hormonal stimulation and isolation of germinal vesicle oocytes

Female B6D2F1 mice, aged 8–12 weeks, were injected with 7–8 IU of pregnant mare serum gonadotropin (PMSG; Bioveta, Ivanovice na Hane, Czech Republic) at noon on the first day and were sacrificed by cervical dislocation ∼44 h later, i.e. in the morning 2 days later. Ovaries were isolated and transferred to EmbryoMax M2 manipulation medium supplemented with 2.5 µM milrinone to suppress germinal vesicle breakdown (GVBD). Cumulus oocyte complexes (COCs) were retrieved from follicles with 27G needles and then transferred to culture medium (Eagle’s α-MEM supplemented as described previously (Fulka and Langerova, 2014)) containing 2.5 µM milrinone. The oocytes were kept in a 5% CO₂ incubator at 37°C. High-quality oocytes were selected as described (Inoue et al., 2008). Germinal vesicle (GV) oocytes were subsequently used for selective enucleation (see below). Alternatively, they were matured in vitro to the metaphase II stage (see below).

Selective enucleation

To prepare cytoplasts of abnormal composition, i.e. reduced levels of nuclear structural proteins and tightly bound chromatin factors, GV-stage oocytes were subjected to selective enucleation as described (Greda et al., 2006; Fulka et al., 2019). Briefly, GV-stage oocytes were placed in M2 medium supplemented with 5 µg/ml cytochalasin D and 2.5 µM milrinone to prevent GVBD. Next, the GV nuclear envelope was attached to the enucleation pipette with a firm grip and pulled until the nuclear envelope ruptured. The resultant cytoplasts were washed to remove cytochalasin D and placed back in the culture medium supplemented with 2.5 µM milrinone.

Oocyte in vitro maturation and preparation of pronucleus-formation-competent selectively enucleated cytoplasts

To obtain in vitro-matured metaphase II oocytes and selectively enucleated (SE) cytoplasts, which are competent to form pronuclei upon fertilization, GV-stage oocytes or SE cytoplasts were placed in culture medium (Fulka and Langerova, 2014) without milrinone to increase maturation-promoting factor activity. Next, the metaphase II oocytes were activated to obtain haploid parthenogenetic (hPA)-recipient embryos or were subjected to in vitro fertilization to generate control embryos. The SE cytoplasts were subjected to in vitro fertilization to obtain SE-androgenetic donor pronuclei. The release from the maturation block was timed with respect to the planned procedure above: 17:00–18:00 on the day of GV oocyte isolation or selective enucleation for in vitro fertilization and the generation of pre-replication hPA-recipient embryos. For the experiments using the reversible inhibitor of DNA replication, aphidicolin, the release of GV oocytes was around 5:00 on the day following GV oocyte isolation to obtain pre-replication parthenogenetic haploid-recipient embryos for aphidicolin-treated donors.

Hormonal stimulation and in vivo embryo generation

Eight- to 12-week-old B6D2F1 females were first injected with 7–8 IU of PMSG (noon). After 48 h, they were injected with 7–8 IU hCG and immediately placed into cages with NMRI males. The next morning, the females were sacrificed by cervical dislocation, and oviducts were isolated and placed in EmbryoMax M2 medium supplemented with hyaluronidase. Embryos containing pronuclei were transferred to EmbryoMax KSOM and cultured at 37°C under a 5% CO₂ atmosphere. The embryos were used for aphidicolin concentration optimization.

Hormonal stimulation and isolation of ovulated metaphase II oocytes

To obtain in vivo-ovulated oocytes, B6D2F1 mice were first injected with 7–8 IU PMSG (17:00–18:00 on Day 1), followed by 5 IU hCG 48 h later (Day 3). After 14–16 h, i.e. on the morning of Day 4, the animals were sacrificed by cervical dislocation and the oviducts were placed in EmbryoMax M2 medium supplemented with hyaluronidase. The COCs were freed from the ampullae and transferred to EmbryoMax KSOM. Next, the oocytes were used for in vitro fertilization to generate control embryos or parthenogenetic activation to obtain hPA-recipient embryos.

In vitro fertilization

In vitro fertilization was performed essentially as described (Takeo and Nakagata, 2011). Briefly, sperm was obtained from NMRI males aged 8–12 weeks and capacitated for 1 h in TYH medium supplemented with 0.75 mM methyl-beta-cyclodextrin, as described (https://www.infrafrontier.eu/emma/cryopreservation-protocols/#Cryopreservation%20and%20IVF). Fertilization was performed in HTF medium supplemented with freshly prepared reduced glutathione for 5 h.

Parthenogenetic activation and hPA-recipient embryo preparation

To prepare recipient zygotes and to determine the replication timing, ovulated metaphase II oocytes were parthenogenetically activated as described (Kishigami and Wakayama, 2007).

Pronuclear transfer

PN transfer was performed either 6- to 7-h or 10- to 11-h post-fertilization/activation using a piezo drill-equipped micromanipulator in EmbryoMax M2 medium supplemented with cytochalasin B at 5 µg/ml. Fusion of the pronucleus karyoplast was induced by HVJ-E (inactivated Sendai virus) diluted 1:10 in the supplied buffer as recommended by the manufacturer (Cosmo Bio USA, Carlsbad, CA, USA). The reconstructed zygotes were moved to KSOM culture medium and incubated at 5% CO₂, 37°C. Fusion was confirmed after 30–60 min. Embryos were either used for analysis or were allowed to develop to the blastocyst stage (96-h post-fertilization).

Labelling of newly synthesized DNA

Labelling of newly synthesized DNA was performed by incubating embryos with 10 µM 5-ethynyl-2′-deoxyuridine (EdU; SiChem, Bremen, Germany). The duration of the EdU incubation was either 1–2 h for experiments mapping the replication timing or 8 h when the effectiveness of different aphidicolin concentrations was examined. Next, embryos were fixed in 4% formaldehyde supplemented with 0.1% Triton X-100 (TX-100, Calbiochem, part of Merck) in phosphate buffer saline (PBS) for 45 min at room temperature. EdU incorporation was assessed using the ClickIT Cell Reaction Buffer Kit and Alexa 488-azide, as recommended by the manufacturer (ThermoFisher Scientific, Prague, Czech Republic). Typically, the EdU assay was performed in conjunction with immunofluorescence analysis (see next section).

Immunofluorescence

Samples were fixed as above, washed three times in PBS, then blocked in 1% bovine serum albumin (BSA) and 0.1% TX-100 in PBS for 1 h at room temperature. Next, the samples were incubated with one of the primary antibodies as follows: anti-RPA32 (1:200, Cell Signaling Technology Inc., Danvers, MA, USA, clone 4E4, rat mAb No. 2208), anti-Rad51 (1:500, Abcam, Cambridge, UK, clone EPR4030(3), rabbit mAb No. ab133534), anti-phosphoserine 139 H2AX (1:200, Abcam, clone 9F3, mouse mAb No. ab26350), anti-Oct4 (1:200, Santa Cruz Biotechnology, Dallas, TX, USA, clone C-10, mouse mAb No. sc-5279), and anti-Gata3 (1:200, Abcam, clone L50-823, mouse mAb No. ab282110). Incubations were carried out in a humidified chamber at 4°C overnight. The samples were then washed three times, 10 min each wash, in PBS/1% BSA/0.1% TX-100 and incubated with a secondary antibody for 1 h at room temperature. The secondary antibodies, purchased from Jackson ImmunoResearch (Ely, UK), were goat anti-mouse immunoglobulins or donkey anti-rabbit immunoglobulins conjugated to Alexa 594 or Alexa 488, diluted 1:1000 in PBS/1% BSA/0.1% TX-100.

Western blots

For optimization of the immunoblotting procedure, somatic tissue lysates were prepared in 1× RIPA buffer (Cell Signaling Technology Inc., Danvers, MA, USA) as recommended by the manufacturer, diluted in BioRad Laemmli buffer supplemented with 2-mercaptoethanol and denatured for 5 min at 95°C. Various volumes were loaded onto a 10% SDS PAGE gel (BioRad TGX FastCast kit, BioRad, Prague, Czech Republic). After separation, the samples were wet-transferred to a PVDF membrane in Towbin buffer with 20% methanol for 2 h at 70 V. Next, the membrane was blocked for 1 h in 2% ECL Prime Blocking Reagent in Tris-buffered saline (TBS) supplemented with 0.05% Tween 20 (TTBS). Antibodies at various concentrations were tested by incubating the membrane for 24 h at 4°C with constant shaking. Then, the membranes were washed six times in TTBS and incubated with peroxidase-conjugated secondary antibody (goat anti-rabbit or goat anti-mouse, Merck-Millipore) diluted 1:100 000 in the blocking solution. Finally, the antibodies were visualized by incubating the membrane with ECL Select detection reagent and imaged using the Azure c400 system (Azure Biosystems Inc. Sierra Trinity, CA, USA). After this initial optimization, 92 and 100 in vitro-matured SE cytoplasts and metaphase II oocytes, respectively, were collected and washed in PBS supplemented with 0.1% polyvinyl alcohol. Next, the samples were transferred to Eppendorf Protein LoBind tubes and frozen at −80°C until use. Prior to use, the samples were resuspended in BioRad Laemmli buffer supplemented with 2-mercaptoethanol and denatured for 5 min at 95°C, then run on a 10% SDS PAGE gel and transferred as above. The subsequent steps were as for the somatic tissue lysates, except that the membrane was cut into individual sections and incubated separately with selected antibodies. Due to the need for a large number of oocytes, this step was performed in compliance with the 3Rs (replacement, reduction, and refinement) principles of animal welfare. Finally, the membrane was stained with Coomassie blue R250 to verify successful and equal transfer. Supplementary Fig. S1 includes uncropped images of western blots of testis lysates showing antibody specificity. The primary antibodies used were anti-polymerase delta (1:2500, Abcam, clone EPR15118, rabbit mAb No. ab186407), anti-MCM7 (1:1000, SantaCruz Biotechnology, clone 141.2, mouse mAb No. sc-9966), anti-PCNA (1:1000, Abcam, clone PC10, mouse mAb No. ab29), anti-MCM4 (1:10 000, Abcam, rabbit pAb No. ab84153-100), anti-polymerase Beta (1:1000, Abcam, clone EPR12722(B), rabbit mAb No. ab175197).

Aphidicolin treatment

A 2-mM aphidicolin stock solution was prepared in DMSO. Concentrations of 2, 0.2, and 0.1 µM aphidicolin were first tested on in vivo-generated embryos to inhibit the progression of replication by incubating the zygotes simultaneously with aphidicolin and 10 µM EdU for 1–10 h in KSOM medium. The 2 and 0.2 µM concentrations were re-tested on the SE-androgenetic samples to ascertain that the preselected concentration inhibited replication in this setting. In the control group, aphidicolin was omitted, and the embryos were cultured only in the presence of DMSO. Next, the samples were fixed and EdU incorporation was assayed as described above. In some cases, the embryos were labelled with the anti-phosphoserine 139 H2A.X antibody following EdU labelling to assess the level of DNA damage caused by aphidicolin and the inhibition of replication. Finally, the effect of 2, 0.2, and 0.1 µM aphidicolin on development was also tested by incubating in vivo-generated zygotes with the drug for 8 h in KSOM medium. In the control embryos, aphidicolin was omitted. Embryos in all groups were allowed to develop for 96 h in KSOM medium. The concentration of aphidicolin used in the PN transfer experiments was 0.2 µM for 10 h in KSOM medium.

RNA-seq library preparation and sequencing

Three samples of five two-cell embryos from each experimental condition were collected, i.e. for sham pronuclear transfer (SPN), early pronuclear transfer (EPN), and late pronuclear transfer (LPN), 34-h post-fertilization. RNA-seq libraries were generated using the Smart-seq2 v4 Kit (Clontech/TaKaRa Bio USA Inc., San Jose, CA, USA) following the manufacturer’s instructions and pooled. Briefly, embryos were lysed and mRNA was captured and amplified using kit components. After purification using AMPure XP beads, the resulting high-quality amplified RNAs were subject to library preparation using a Nextera XT DNA Library Preparation Kit (Illumina, San Diego, CA, USA) and multiplexed using Nextera XT Indexes (Illumina). The concentration of sequencing libraries was determined using the Qubit dsDNA HS Assay Kit (Life Technologies/Thermo Fisher Scientific, Waltham, MA, USA), while the size of the libraries was determined using Agilent D5000 ScreenTape with the Tapestation 4200 system (Agilent, Santa Clara, CA, USA). Pooled indexed libraries were sequenced on the Illumina NovaSeq 6000 platform with 150-bp paired-end reads.

RNA-seq data processing

Multiplexed sequencing reads that passed filters were trimmed to remove low-quality reads and adaptors by TrimGalore (version 0.6.7) (-q 25 --length 20 --max_n 3 --stringency 3). The filtered reads were passed through quality control using FastQC3 (v0.11.5), followed by alignment to the mouse genome (GRCm39) by HISAT2 (version 2.2.1) using default parameters. The output SAM files were converted to BAM files and sorted using SAMtools6 (version 1.14). Read counts of all samples were quantified using Feature Count (Version 2.0.1) with the mouse genome as a reference and were adjusted to provide CPM (counts per million mapped reads). Clustering analysis was performed using R (version 3.3.3), and principal component analysis (PCA) was used to visualize the clustering results. Differential gene expression analysis was performed using the R package edgeR set to default parameters. The genes were deemed differentially expressed if they provided a false discovery rate of <0.01 and a fold change >2. Biological pathways (denoted as Gene Ontology terms) over-represented among up- and downregulated genes were identified using the R package ClusterProfiler. A P-value cut-off of 0.05 was used to select significant pathways.

Statistical analysis

Statistical analysis was performed in RStudio. The tests applied were Pearson’s chi-squared test with Yates’ continuity correction for the developmental rates and the Mann–Whitney U-test for the number of inner cell mass (ICM) cells.

Results

Using selective enucleation to model an idiopathic developmental failure

First, we developed a system that mimics an idiopathic developmental failure. We have previously used cytoplasts generated by selective enucleation as a milieu to induce defective PN formation and likely aberrant parental genome remodelling (Mohammed et al., 2008; Fulka et al., 2019). The SE cytoplasts can produce seemingly normal zygotes when supplemented with metaphase II spindles and later fertilized in vitro (Supplementary Fig. S2). However, the reconstructed embryos mostly fail to reach the blastocyst stage as might be expected (5/46; 11%). Our initial intention was to transfer both parental pronuclei remodelled under abnormal conditions into a healthy recipient cytoplasm. However, when we attempted this, the pronuclei were too fragile and often suffered mechanical damage (data not shown). Furthermore, since the pronuclei and all the associated proteins are removed during the preparation of a recipient cytoplast, the recipient cytoplast might not contain sufficient nuclear building blocks to normalize both SE pronuclei. We therefore decided to use a system where only one pronucleus is prepared using the SE cytoplast. To select the parental genome/pronucleus to be transferred, SE-parthenogenetic or SE-androgenetic embryos were prepared by spindle transfer and activation or by IVF, respectively. We observed the most profound effect on the paternal genome, as the SE-androgenetic embryos rarely developed beyond the two-cell stage and showed signs of cytoplasm fragmentation. In comparison, the SE-parthenogenetic embryos typically arrested around the eight-cell stage (data not shown). In other words, a more pronounced developmental arrest was observed in the SE-androgenetic embryos. Next, we asked whether the adverse effects on development could be mitigated by transferring the SE/remodelled male pronucleus to a healthy cytoplasm and complementing it with a haploid female pronucleus (Fig. 1A). We reasoned that this experimental scheme should mostly normalize the deficiencies inflicted on the nuclear component by the selective enucleation procedure.

Figure 1.

Experimental scheme showing pronuclear transfer, characterization of the DNA replication machinery content, and timing of DNA synthesis in the donor embryos obtained by fertilizing selectively enucleated cytoplasts and recipient haploid parthenogenetic mouse embryos. (A) The scheme shows an outline of the experimental setup used to model an idiopathic embryonic developmental arrest and the time of pronuclear (PN) transfer. (B) New DNA synthesis was detected with the same timing, i.e. 6- to 7-h post-fertilization (hpf) or activation, in the embryos generated by IVF in the cytoplasts prepared by selective enucleation (SE) as well as those prepared by parthenogenetic activation (hPA). No marked differences between the intensities were noted. (C) A summary showing that the majority of SE and hPA embryos initiate replication at 6–7 h and complete replication by 10-h post-fertilization/replication. (D) Although the timing and the intensity do not seem to differ in SE and hPA embryos, the SE cytoplasts showed lower levels of key components of the DNA replication machinery, i.e. DNA polymerases delta (POLD) and beta (POLB), which are involved in DNA replication as well as DNA repair, and PCNA, proliferating cell nuclear antigen, which serves as a processivity factor during replication. By contrast, replication licensing, as indicated by the level of MCM4 (minichromosome maintenance complex component 4), seemed to be unaffected.

To obtain more insight into the possible cause of developmental arrest and what needed to be corrected in SE-androgenetic embryos, we performed a more detailed analysis. As mentioned, these embryos typically cleaved to the two-cell stage, but in most cases, they contained micronuclei as well as pronounced chromosome bridges, indicating defective chromosome segregation (Supplementary Fig. S3). Both chromosome bridges and micronuclei can arise due to a defective mitotic spindle, an abnormal first embryonic replication, or both (Mashiko et al., 2020; Palmerola et al., 2022). As the PN transfer is performed in interphase, i.e. prior to the formation of the mitotic spindle, we focused on the first embryonic interphase for further experiments.

A compromised cytoplasm is associated with persistent replication stress

Although mouse embryos can tolerate a certain degree of asynchrony between parental pronuclei, we decided to time-match the donor embryos, i.e. SE-haploid androgenetic embryos, to recipients, i.e. hPA embryos, to limit the variables in our experimental setup. To map the beginning and end of the first embryonic replication, we incubated the embryos with EdU at 1- to 2-h intervals around the time of the suspected replication start and end, to set the timepoint for the PN transfer procedure. In both groups, the beginning of replication was observed ∼6- to 7-h post-activation or fertilization. At 4- to 6-h post-activation/fertilization, both hPA and SE-androgenetic embryos were mostly unlabelled, with only 19% (6/32) of the hPA and 14% (4/28) of the SE-androgenetic embryos showing some EdU incorporation. However, 6- to 7-h post-activation or fertilization, this pattern changed, and EdU incorporation was detected in 81% (25/31) of hPA and in 78% (35/45) of SE-androgenetic embryos (Fig. 1B and C). This is in contrast to biparental embryos containing maternal and paternal genomes, which incorporated EdU as early as 4- to 5-h post-IVF (21/28, EdU positive/total; data not shown). At 10- to 12-h post-activation/fertilization, the embryos from all groups were in late S-phase or had already ceased replication, which is consistent with previous reports (Palmerola et al., 2022). Here, only 12.5% (3/24) of the hPA embryos and none of the SE-androgenetic embryos (0/21) were classified as EdU positive, i.e. showing incorporation outside of the peri-nucleolar precursor body region (Fig. 1C). The comparable replication timing between SE-androgenetic and hPA embryos is somewhat surprising, given that lower levels of several factors involved in replication were detected in the SE cytoplasts compared to ovulated metaphase II oocytes (Fig. 1D, Supplementary Fig. S1). This indicates that while replication occurs in both the donor and recipient embryos, the quality of replication might be impacted in the SE-androgenetic embryos.

For this reason, we analysed SE-androgenetic as well as hPA embryos for the presence of replication stress and DNA damage. Not surprisingly, the SE-androgenetic embryos exhibited DNA damage markers just prior to cleavage to the two-cell stage, i.e. in G2 phase (Fig. 2). These include RPA32, which binds single-stranded DNA intermediates and under-replicated DNA (Maréchal and Zou, 2015; Saxena and Zou, 2022), histone H2A.X phosphorylated on serine 139 (Moeglin et al., 2019; Saxena and Zou, 2022), a general marker of replication stress, and RAD51, which is involved in replication fork stability (Feu et al., 2022; Saxena and Zou, 2022). Because these markers were also present in the nuclei of early pre-replication G1-stage SE two-cell-stage embryos, the SE cytoplasts lacked the factors necessary for DNA repair, and the damage was likely permanent (Fig. 2). By contrast, the hPA embryos exhibited no signs of these markers (Fig. 2). This shows that DNA damage in the donor pronuclei is caused neither by our culture conditions nor by altered replication timing of the SE-androgenetic embryos.

Figure 2.

Immunostaining of markers of DNA damage in the male pronuclei of donor embryos and the female pronuclei of recipient embryos in relation to the first and second embryonic replications. The labelling was performed 13- to 14-h post-fertilization or activation, i.e. very late S phase/G2 phase. The top panel shows the male pronuclei formed in compromised selectively enucleated cytoplasts (SE). The bottom panel shows the female pronuclei in haploid parthenogenetic embryos (hPA) at the same time point. In the SE embryos, markers of DNA damage (γH2A.X) and replication stress (RPA32 and Rad51) appear following the first embryonic replication and persist up to the G1 phase of the two-cell stage, indicating that the cytoplasts have a reduced DNA repair capacity. The central scheme depicts new DNA synthesis and EdU incorporation. By contrast, haploid parthenogenetic embryos (hPA, bottom) do not show DNA damage markers.

A healthy recipient cytoplasm can alleviate replication-induced DNA damage, but this is not sufficient to warrant full developmental potential following PN transfer

Next, we decided to test to what extent a healthy recipient cytoplasm can mitigate the adverse effects of abnormal post-fertilization genome remodelling. We performed PN transfer between SE-IVF embryos and hPA embryos obtained from ovulated oocytes. The PN transfer was carried out either 6- to 7-h or 10- to 11-h post-fertilization; these times coincide with the pre-replication and post-replication phases in our system. As controls, we performed a sham PN transfer of IVF-generated embryos 10- to 11-h post-fertilization (post-replication). In all three groups, the PN transfer embryos developed well to the two-cell stage, but a significant decrease in the overall developmental rate was noted in the post-replication SE-PN transfer group (Fig. 3A and Supplementary Table S1). As the blastocyst stage is important from the clinical perspective and as embryo transfers are most frequently performed at this stage, the embryo quality was investigated further. Although the post-replication SE-PN transfer blastocysts seemed to have less-well-defined ICMs following staining with anti-POU5F1 (Oct3/4), and a slightly reduced number of ICM cells compared to the pre-replication SE-PN transfer group, the difference in number of ICM cells between these groups was not statistically significant (post-replication SE-PN transfer group: 7 ± 1.41, n = 9; pre-replication SE-PN transfer group: 11.45 ± 8.25, n = 11; P-value = 0.2079, Mann–Witney U-test) (Fig. 3B, Supplementary Table S2). These results indicate a link between the first embryonic replication and the developmental potential of embryos.

Figure 3.

Developmental rate and blastocyst morphology of the reconstructed embryos formed by transfer of the male pronuclei generated in selectively enucleated cytoplasts to haploid parthenogenetic recipients in relation to the timing of the first embryonic replication. (A) When the pronuclear transfer is performed prior to the first embryonic replication (pre-replication SE-PN transfer), the resulting embryos develop at the same rate as control embryos obtained by IVF (Sham post-replication PN transfer). Between 82% and 83% of embryos reach the blastocyst stage. However, when the PN transfer is performed after the first embryonic replication, the developmental rate drops to 61% (post-replication PN transfer). These results were statistically significant (Pearson’s chi-squared test with Yates’ continuity correction, P-value = 0.001804). (B) Although both SE-PN transfer groups of embryos can develop to the blastocyst stage, the post-replication SE-PN transfer embryos typically exhibit fewer cells in the inner cell mass—ICM (Oct3/4; in green), and slightly more scattered ICMs (right image). In contrast, the pre-replication SE-PN transfer embryos show overall better morphology (left image). However, the difference in the number of ICM cells was not statistically significant (Mann–Whitney U-test, P-value = 0.2079).

The timing of PN transfer impacts the embryonic gene expression

To further investigate the possible cause of the reduced developmental competence observed above, we assessed all manipulated embryos for DNA damage and abnormal chromosome segregation. However, none of the three groups showed micronuclei or signs of DNA damage or replication stress (Fig. 4). Therefore, it seemed that healthy recipient embryos can repair the damaged DNA arising from defective zygotic replication and suppress abnormal chromosome segregation. For this reason, we asked whether the reduced developmental rates in post-replication SE-PN transfer embryos could be linked to more subtle changes, which might not be uncovered by standard preimplantation genetic diagnosis. We therefore performed RNA-seq analysis to look for changes in embryonic gene expression that might be caused by the manipulations (Fig. 5). As controls, we again used sham PN transfer embryos. Overall, more than 18 000 transcripts were identified in our samples. PCA analysis of transcriptome data showed that biological replicates in each of the experimental groups clustered together (Fig. 5A). The majority of genes were expressed at comparable levels in both the control sham PN transfer and SE-PN transfer groups. Nevertheless, a clear trend in the number of dysregulated genes became apparent depending on the timing of the PN transfer. While only 641 transcripts were found to be dysregulated in the embryos when SE-PN transfer was performed prior to the first replication, the number of dysregulated genes increased to 1991 in the post-replication SE-PN transfer group. This is more than a 3-fold increase, indicating that the longer the pronuclei are allowed to form and function in suboptimal conditions, the larger the effect on embryonic gene expression (Fig. 5B). Gene ontology analysis showed that the time-dependent differentially expressed genes are involved in various cellular processes, such as the regulation of actin filaments, RNA processing, and mitochondrial function (Fig. 5C).

Figure 4.

The dramatic reduction in developmental rate cannot be explained by persistent DNA damage in the transferred male pronuclei. The DNA damage markers initially detected following the first embryonic replication were absent by the next embryonic stage when the pronuclei were transferred to a healthy recipient cytoplast. Here, the G1 phase nucleus of the two-cell stage is shown for the post-replication SE-PN transfer embryo.

Figure 5.

RNA-seq analysis shows a more pronounced dysregulation of embryonic gene expression when pronuclear transfer is performed after the first embryonic replication. (A) Principal component analysis shows clustering of individual experimental groups of embryos. (B) While in the pre-replication pronuclear transfer group (early PN transfer, EPN) only 641 genes were differentially expressed compared to the control group (Sham PN transfer, SPN), following replication, the number of differentially expressed genes (DEG) in PN transfer embryos (late PN transfer, LPN) increased to 1991 in comparison to controls. (C) The top five Gene Ontology pathways downregulated in the LPN transfer embryos compared with the EPN transfer embryos are mainly involved with the metabolism of various RNAs and mitochondrial function.

The first embryonic replication is vital for the developmental competence of embryos

As many factors can cause the observed changes to major genome activation and to developmental competence, we wanted to ask whether there is a substantial effect on developmental competence if the replication occurs under suboptimal conditions. To test this, we introduced aphidicolin, a reversible inhibitor of DNA replication, into the system. Aphidicolin is known to block cell division at the border between the G1 and S phases of the cell cycle when added prior to replication (Wang, 1991; Borel et al., 2002). However, it also causes DNA damage and noticeably reduces developmental competence in embryos in a dose-dependent manner (Glover et al., 1984; Spindle et al., 1985; Wang et al., 2014). To minimize these adverse effects, we decided to optimize the concentration of this drug. In the first set of experiments, we used in vivo-generated zygotes and incubated them in the presence of 2, 0.2, and 0.1 µM aphidicolin. In the control group, aphidicolin was omitted. Simultaneously, the embryos were pulsed with 10 µM EdU for 1 h. Unlike 0.1 µM aphidicolin, both 2 and 0.2 µM aphidicolin markedly reduced the incorporation of EdU (Fig. 6A). No difference in the EdU incorporation of controls and 0.1 µM aphidicolin was observed (not shown). In a second experiment, 2 µM aphidicolin noticeably reduced the developmental competence of embryos with only 2/20 embryos developing to the blastocyst stage. By comparison, 19/20 formed blastocysts when aphidicolin was omitted (representative embryo images are shown in Supplementary Fig. S4). This result is in agreement with previously published results (Wang et al., 2014).

Figure 6.

Inhibition of DNA replication by aphidicolin significantly increases the pronuclear transfer success rate. (A) In the first set of experiments, the effects of different concentrations of aphidicolin on DNA replication in embryos obtained in vivo were tested. Both 2 and 0.2 µM aphidicolin reduced EdU incorporation, with 2 µM aphidicolin having a stronger effect. (B) However, when re-tested on donor male pronuclei, 2 µM aphidicolin also resulted in marked levels of phosphorylated H2A.X, indicating pronounced DNA damage. For this reason, 0.2 µM aphidicolin was used, although EdU incorporation was not completely abolished. (C) To better understand the effect of the first embryonic replication on developmental competence, we incorporated aphidicolin treatment into the experimental scheme of pronuclear transfer. (D) The developmental rates of embryos obtained in a series of PN transfers using in vitro-matured hPA recipients under different conditions. The use of hPA recipients obtained from in vitro-matured oocytes allowed us to synchronize various donor and recipient embryos. Overall, very few differences were observed early in development, and the majority of embryos developed to the two-cell stage regardless of the procedure. In contrast to the experiments where hPA-recipient embryos originated from ovulated oocytes, reduced developmental rates were recorded for the in vitro-produced embryos. Indeed, only 8% of embryos were able to develop to the blastocyst stage when post-replication SE-PN transfer was performed, although 72% of embryos in the pre-replication group developed to the blastocyst stage (Pearson’s chi-squared test with Yates’ continuity correction, P-value = 4.161e-08). When aphidicolin was used to block replication, the developmental rate dramatically increased. In this case, 63% of embryos reached the blastocyst stage. This is not statistically different from the pre-replication group (Pearson’s chi-squared test with Yates’ continuity correction, P-value = 0.5327). The actual developmental rates and a more detailed statistical comparison are given in Supplementary Table S3. (E) Morphologically normal blastocysts were obtained when replication was first blocked by low concentrations of aphidicolin prior to PN transfer.

Next, the 2- and 0.2-µM aphidicolin concentrations were tested in SE-androgenetic embryos. These were simultaneously incubated with both aphidicolin and 10 µM EdU for 10 h from the time of fertilization. Since SE-IVF embryos do not develop well, as already mentioned, we evaluated the presence of phosphorylated H2A.X instead of following the developmental rate to assess the DNA damage caused by aphidicolin. As expected, 2 µM aphidicolin induced a much higher level of phosphorylated H2A.X than 0.2 µM, while EdU incorporation was negligible at both concentrations (Fig. 6B). Integrating these results, we opted for 0.2 µM aphidicolin to inhibit the replication in the SE-IVF-generated pronuclei before transfer. As we wanted to mimic the original conditions as closely as possible, the total duration of the aphidicolin treatment was 10 h to cover the whole period of replication.

Instead of using hPA embryos derived from in vivo-ovulated oocytes, we opted for in vitro-matured hPA recipients as the synchronization of multiple groups and procedures was necessary. The experimental scheme, including the aphidicolin treatment, is shown in Fig. 6C. Early and late SE-PN transfers to in vitro-matured hPA recipients were also performed to obtain parallel groups for comparison. Overall, slightly lower developmental rates were observed in all groups when compared to the hPA recipients obtained by ovulated oocyte activation (Supplementary Table S3). This indicates that lower-quality oocytes are produced overall by in vitro maturation. Rather surprisingly, compared to the in vivo hPA recipients, a much more pronounced negative effect was observed in the post-replication SE-PN transfer group than in the pre-replication SE-PN transfer group. Here, only 8% of embryos developed to the blastocyst stage when the replication occurred in the SE environment; in contrast, 72% of embryos reached the blastocyst stage when the transfer was performed by 6- to 7-h post-fertilization and activation (Fig. 6D and Supplementary Table S3). Most importantly, when 0.2 µM aphidicolin was used to block replication in the SE-IVF group, and the male pronuclei were transferred to pre-replication in vitro-matured hPA recipients 6-h post-activation, the developmental rate increased to 63% and normal-looking blastocysts were obtained (Fig. 6E). As this is only 9% lower than the developmental rate of the pre-replication SE-PN transfer group, we conclude that the conditions in which the first embryonic replication takes place strongly influence developmental competence, and that the effects might be irreversible.

Discussion

Here, we focused on PN transfer, a procedure approved and used in human-assisted reproduction, and assessed whether the timing of this intervention influences developmental potential and embryo quality. Our results using a mouse model of idiopathic developmental arrest show that the first embryonic replication represents an important landmark.

In the context of research on reproductive biology, the main advantages of using inbred strains of laboratory mice as a model are the genetically uniform biological material and the relatively large number of oocytes that can be obtained from individual females. Another considerable advantage is that culture systems, including the in vitro maturation of oocytes, and assisted reproduction techniques are well-established. These advantages allowed us to perform a detailed analysis of the factors that might be important for successful PN transfer. We examined two different time points for the PN transfer, corresponding to the G1 and G2 phases of the first embryonic cycle. Our initial experiments indicated that the longer the male pronucleus resides and functions in the abnormal environment of the SE-cytoplast, the lower its developmental potential. Using aphidicolin, a reversible inhibitor of DNA replication, we showed that the reduced developmental potential can be attributed to the compromised replication in the abnormal environment.

The first embryonic S phase is extremely important as any mutations and unresolved DNA integrity issues threaten the viability of the whole embryo. Our results indicate that mouse embryos possess a robust DNA repair machinery, because multiple markers of DNA damage that were detectable in the SE-male pronuclei, likely an immediate consequence of compromised DNA synthesis, disappeared by the time the embryos entered the next S phase, which took place after the male pronuclei have been moved to a healthy cytoplasm. It is currently unclear whether the bulk of DNA repair occurred during the zygotic G2 phase, i.e. directly following the PN transfer, or during the G1 phase of the two-cell embryonic stage. To what extent human embryos, where PN transfer is used in clinical practice, would be capable of repairing the introduced DNA is unclear and remains to be determined. It should be mentioned that human embryos are intrinsically prone to replication stress, and this has been suggested as the leading cause of their developmental failure (Palmerola et al., 2022). There are additional differences between humans and mice. For example, mouse embryos exhibit a higher zygotic mutation rate than human embryos (Milholland et al., 2017), but they are less prone to chromosome missegregation (Currie et al., 2022). Whether this indicates a higher fidelity of DNA duplication processes, but a less efficient DNA repair system in human embryos, or simply a higher sensitivity to DNA lesions and replication stress in humans, is unknown.

Because markers of DNA damage were not observed beyond the first cell cycle in our model, we assume that DNA damage is unlikely to be the main reason for the reduced developmental rates in the post-replication PN transfer group. Post-fertilization genome remodelling might also be affected. Indeed, the first embryonic replication serves not only to duplicate the DNA, but is also crucial for reprogramming the terminally differentiated gametes to the totipotent state and for setting up the embryonic chromatin landscape (Nakatani et al., 2022, 2024). Our RNA-seq analysis of the two-cell PN transfer embryos showed that there is a correlation between the time the parental genomes spend in the abnormal cytoplasm and the number of dysregulated genes at the time of the major genome activation. Although we did not determine to what extent this is directly linked to the first embryonic S phase, it is possible that the quality of the first replication is indeed responsible. In agreement with our results, it has been shown that the first round of replication can impact gene expression during the major genome activation. Already in 1997, Davis and Schultz (1997) identified a subset of replication-sensitive peptides when zygotic DNA synthesis was inhibited in mouse embryos, indicating a link between zygotic replication and embryonic gene expression. Although the mouse is quite unique from the perspective of the main wave of embryonic genome activation, which occurs particularly early following fertilization (for review, see Lee et al., 2014; Jukam et al., 2017), the same effect of zygotic DNA synthesis inhibition was also observed in bovine embryos (Memili and First, 1999). Whether the same is true in human embryos is unclear, but bovine embryos are often used as a model instead of human samples, as they share many similarities, including the timing of the genome activation (Graf et al., 2014; Jiang et al., 2014); for review, see Ménézo and Hérubel (2002).

The major limitation of our work is that our system allowed us to analyse the effect of abnormal post-fertilization remodelling and replication on only one of the parental genomes. We selected the paternal genome because, although the experimental scheme is technically challenging, it is feasible to demonstrate the principle. Analysis of the maternal genome would require additional micromanipulations, such as the enucleation of metaphase II oocytes and the transfer of spindles to the SE cytoplasts, coupled with IVF of the metaphase II cytoplasts, prior to PN transfer. This is overly complex and challenging to synchronize. Although the maternal genome is considered to undergo a less dramatic remodelling during the post-fertilization phase, and the epigenetic changes seem less dynamic than in the paternal genome (for review, see Burton and Torres-Padilla (2010)), this is probably not relevant because replication is a process common to both parental genomes. Nevertheless, the effect, if any, of a compromised post-fertilization remodelling or replication on the maternal genome remains an open question.

In summary, this study demonstrates that the first embryonic replication can have a profound effect on the developmental competence of embryos. Our findings support the notion that the transfer of genetic material should be performed prior to the first embryonic replication, particularly in the clinical setting.

Supplementary data

Supplementary data are available at Molecular Human Reproduction online.

Data availability

The raw FASTQ files are available at Gene Expression Omnibus (GEO) (https://www.ncbi.nlm. nih.gov/geo/) under the accession numbers: GSM8028245-GSM8028253.

Author roles

H.F.: conceptualization of the study and design, data acquisition, interpretation of the data, manuscript preparation; T.Z.: experimental work; N.D.: data analysis and interpretation, manuscript preparation; H.M. and Z.J.: data acquisition, analysis and interpretation, conceptualization.

Funding

Czech Science Foundation (GACR 20-04465S); NIH Eunice Kennedy Shriver National Institute of Child Health and Human Development (R01HD102533 to Z.J.); USDA National Institute of Food and Agriculture (2019-67016-29863 to Z.J.).

Conflict of interest

The authors declare no conflict of interest.