Two candidate single guide (sg) RNAs targeting exon 2 (E2-1 and E2-2) and one targeting exon 5 (E5) of the porcine IGF1R gene were designed using BE-Designer online software (http://www.rgenome.net) and commercially synthesized by Synthego (Figure 1A; Supplementary Table S1). The BE4 mRNA was generated by in vitro transcription of pCMV-BE4-RrA3F plasmid (a gift from Nicole Gaudelli (Addgene plasmid #138340; http://n2t.net/addgene:138340; RRID:Addgene_138340)) using the mMESSAGE mMACHINE T7 kit (Invitrogen) according to the manufacturer’s instructions. Following transcription, the mRNA was purified by ethanol precipitation. Candidate sgRNAs were co-transfected with BE4 mRNAs into porcine fetal fibroblasts (PFF) by nucleofection (Lonza), followed by culture in 10-cm dishes at a seeding density of 5 cells/cm² to generate single-cell colonies as per established and published protocols (Simpson et al., 2024; Park et al., 2022; Park et al., 2017). After 10–14 days, single-cell colonies were isolated and genotyped by Sanger sequencing (Supplementary Table S2).

Genomic DNA from candidate PFF clones was isolated by lysing cells using lysis buffer (40 nM Tris-HCl, 1% Triton X-100, 1% NP-40, and 0.4 ng/mL proteinase K) at 55 °C for 1 h and 95 °C for 10 min. PCR amplification was performed using either KOD Hot Start master mix (Novagen) or Bioline Taq with the following cycling conditions: initial denaturation and polymerase activation at 95 °C for 3 min, followed by 35 cycles of 95 °C for 20 s, 60 °C for 10 s, and 70 °C for 10 s, with a final extension at 70 °C for 5 min. PCR products were purified using the NucleoSpin gel and PCR clean-up kit (Machery Nagel) and subjected to Sanger sequencing. The primers used were provided in Supplementary Table S2. Targeted genomic sites were amplified using Phusion polymerase (Thermo Fisher Scientific). Paired-end sequencing of PCR amplicons was conducted on an Illumina MiSeq platform by Novogene Bioinformatics Technology Co. Ltd. Potential off-target sites for each sgRNA were predicted using Cas-OFFinder (http://www.rgenome.net/cas-offinder/) to analyze site-specific edits. For the off-target assay, up to three base mismatches and no DNA or RNA bulges were permitted.

Cumulus-oocyte complexes (COCs) were obtained from De Soto Biosciences (Seymour, TN, United States). We generated IGF1R wild-type (WT) and knockout (KO) embryos by somatic cell nuclear transfer (SCNT) using WT and KO PFFs. The WT and KO PFFs were synchronized to the G1/G0 phase by serum deprivation (DMEM with 0.1% FBS) for 96 h. Oocytes were enucleated by aspirating the polar body and MII metaphase plates with a micropipette (Humagen, Charlottesville, VA, United States) in 0.1% DPBS supplemented with 5 μg/mL cytochalasin B. Following enucleation, a single donor cell was placed into the perivitelline space of each enucleated oocyte. Cell-oocyte couplets were fused by applying two direct current pulses (1-s interval) of 2.0 kV/cm for 30 microseconds using an ECM 2001 Electroporation System (BTX, Holliston, MA). After fusion, reconstructed zygotes were activated by a direct current pulse of 1.0 kV/cm for 60 microseconds, followed by post-activation in 2 mM 6-dimethylaminopurine for 3 h (Park et al., 2021). After overnight culture in PZM3 medium containing the histone deacetylase inhibitor Scriptaid (0.5 μM), reconstructed zygotes were cultured in PZM3 under low oxygen conditions (5% O₂ and 5% CO2 in 90% N2) (Park et al., 2021).

Apoptotic activity in embryos was assessed using a live caspase-3/7 detection assay (Invitrogen). Embryos were incubated in culture medium supplemented with the caspase-3/7 reagent at a final dilution of 1:400 for 30 min at 38.5 °C under standard culture conditions. Following incubation, embryos were washed briefly in fresh medium and immediately imaged using fluorescence microscopy.

To determine dynamic range and a non-toxic working concentration of OSI-906 (OSI), a dual IGF1R/INSR tyrosine kinase inhibitor (Mulvihill et al., 2009), presumptive zygotes were cultured continuously from the one-cell stage to the blastocyst stage in PZM-3 medium supplemented with OSI-906 at final concentrations of 0, 0.001, 0.01, 0.1, 1, or 10 µM. In the second experiment, embryos that reached the morula stage were randomly assigned to treatment groups and cultured in PZM-3 medium supplemented with OSI-906 at concentrations of 0, 0.1, 1, 2, 5, or 10 µM. Media were refreshed daily with freshly prepared inhibitor. Embryos were maintained under standard culture conditions, and media containing freshly prepared OSI-906 were replaced every 24 h. Cleavage rates were assessed at the 2-4 cell stage (48 h post-activation), and blastocyst formation was evaluated on day 7 (168 h).

RNA isolation and cDNA synthesis were performed as previously described (Sheets et al., 2018). Primers listed in Supplementary Table S3 were used for qRT-PCR. Early embryos were rinsed three times with 0.1% PBS/polyvinylpyrrolidone and fixed with 4% paraformaldehyde in PBS for 5 min. Samples were permeabilized with 0.5% Triton X-100/PBS for 30 min, then blocked for 2 h with a buffer containing 5% bovine serum albumin and 0.1% Triton X-100 in PBS. Samples were incubated with primary antibodies overnight at 4 °C, followed by treatment with secondary antibodies for 1 h. Nuclear DNA was counterstained with DAPI (Thermo Fisher Scientific) for 5 min. Samples were mounted and imaged using a Leica DM 4000B microscope and the Leica Application Suite Advanced Fluorescence software (Leica Microsystems, Wetzlar, Germany). All antibody information is provided in Supplementary Table S4.

All quantitative data are presented as mean ± SEM unless otherwise indicated. Statistical analyses were performed using GraphPad Prism 9 (GraphPad Software, San Diego, CA, United States). For comparisons between two groups, unpaired two-tailed Student’s t-tests were applied when data satisfied assumptions of normality and equal variance. For comparisons among multiple groups, one-way ANOVA followed by Dunnett’s multiple comparisons test was used to assess differences relative to the control group, as indicated in the figure legends. Developmental rates and proportions (e.g., blastocyst formation rate and percentage of caspase-3 positive embryos) were analyzed using the same statistical framework. Each experiment was performed using at least three independent biological replicates unless otherwise stated, and the number of embryos analyzed per group is provided in the corresponding figure legends. Differences were considered statistically significant when P < 0.05.

To elucidate the developmental context of IGF1R function, the expression pattern was examined during porcine preimplantation development. Quantitative RT-PCR analysis revealed a canonical maternal-to-zygotic expression pattern: IGF1R transcripts were abundant in oocytes, decreased progressively through the 4- to 8-cell stages, increased again at the morula stage, and declined again in blastocysts. This temporal pattern coincided with compaction and blastocoel formation, suggesting stage-specific regulation of IGF1R expression (Figure 1A). Immuno-fluorescence analysis demonstrated dynamic changes in IGF1R localization (Figure 1B). During early cleavage stages, IGF1R protein was predominantly cytoplasmic. From the late 8-cell stage onward, the IGF1R signal became detectable within the nucleus, and by the blastocyst stage, a subset of embryos exhibited punctate intranuclear enrichment consistent with nucleolar localization. In mid-to late-stage blastocysts, IGF1R was further concentrated within a subset of nuclei as distinct punctate foci. These observations indicate that IGF1R undergoes regulated spatial redistribution during preimplantation development, although the functional significance of this redistribution at this stage remains unresolved.

To investigate the potential role of IGF1R, a functional KO of the porcine IGF1R gene was generated using the fourth-generation cytidine base editor (BE4). Candidate sgRNAs were designed to target the extracellular domain of IGF1R within exon 2 (E2-1, E2-2) and exon 5 (E5-1), positioning the catalytic cytidines within the BE4 editing window (N13-N17 upstream of the PAM). BE4-mediated conversion of C-to-T at these positions was expected to introduce premature termination codons (PTCs) and functionally inactivate the gene (Figure 2A) (Yu et al., 2020). Transfection of porcine fetal fibroblasts (PFFs) with BE4 mRNA and sgRNAs resulted in efficient cytidine deamination, as confirmed by targeted amplicon sequencing (Amp-seq). Editing efficiencies varied by sgRNA, yielding 17% (E2-1), 56% (E2-2), and 73% (E5-1) conversion, with E5-1 showing the highest efficiency (Figure 2B; Supplementary Table S5). Other editing outcomes, including imperfect and minimal indel formation (0.69%–1.03%), were also detected (Figure 2C; Supplementary Figure S1). For comparison, SpCas9 editing with the same sgRNAs yielded lower overall efficiencies (13%, 37%, and 37%, respectively) (Figure 2B) and resulted in a heterogeneous outcome, including both in-frame and frameshift mutations (Supplementary Table S1; Supplementary Figure S5). In contrast, BE4 editing predominantly yielded precise PTC-inducing substitutions, demonstrating higher accuracy, albeit guide-dependent (Figure 2C). Single-cell colonies derived from the edited PFFs were expanded and sequenced, revealing that 9 of 24 clones carried the intended point mutations introducing the PTC. Specifically, 2 of 12 (16.7%) E2-1 colonies harbored the Q44* mutation, 2 of 5 (40%) E2-2 targeted colonies carried the W157* mutation, and 5 of 7(85.7%) E5-1 targeted colonies contained the Q407* mutation (Figure 2D). All edited colonies were homozygous for the introduced PTCs, except for a single heterozygous E2-1 clone (Supplementary Figure S1B). Sanger sequencing confirmed precise C-to-T substitutions generating the corresponding stop codons, with no off-targeting events detected. A homozygous IGF1R KO clone (E2-4) and the precursor PFF were selected as nuclear donors for SCNT to generate KO and WT control embryos, respectively.

The functional requirement for IGF1R during early development was assessed by comparing developmental kinetics between IGF1R WT and KO SCNT embryos. Cleavage rates and overall blastocyst formation rates were indistinguishable between the two groups, indicating that IGF1R is dispensable for early mitotic divisions and blastocyst formation (Figure 3A). Despite similar blastocyst rates, KO blastocysts contained significantly fewer total cells than WT controls (p < 0.01; Figure 3B). Time-course analysis showed that while 85.4% of WT morulae had progressed to early blastocysts by day 5 (Figure 3C), the majority (73.4%) of KO embryos remained at the morula stage at this time point. By day 6, 91% of WT embryos predominantly reached early to mid-blastocyst stages, whereas only a minority (32.2%) of KO embryos did. Although most KO embryos (82.2%) eventually formed blastocysts by day 7, fewer (32.2%) progressed to expanded or hatching stages (Figures 3C,D), indicating that KO embryos displayed a pronounced delay in developmental kinetics.

The impact of growth delay on early lineage specification was assessed by examining the expression of key transcription factors associated with TE, epiblast (EPI), and primitive endoderm (PrE) identities. Immunofluorescence confirmed the presence of CDX2-positive TE, SOX2-positive EPI, and SOX17-positive PE in both groups, with no detectable differences in lineage proportions (Figure 4A). Despite the defects in blastocyst development, lineage allocation remained largely intact. Quantitative analysis revealed more subtle shifts consistent with developmental delay rather than fate disruption (Figure 4B). SOX2 transcripts were reduced in KO blastocysts relative to WT embryos, whereas CDX2 expression was modestly elevated. GATA6 expression was largely preserved (Figure 4C). Modest reductions in hypoblast-associated transcripts (SOX17) were observed in some KO samples (Figure 4C). Collectively, these data indicate that IGF1R primarily regulates blastocyst growth kinetics and expansion, while lineage specification and spatial segregation remain intact in its absence.

The association between impaired blastocyst development in KO embryos and increased cell death was evaluated using live caspase-3/7 staining and transcript analysis. During early cleavage stages, Occasional caspase-3/7-positive (CS+) signals were detected; however, these were spatially restricted to small punctate structures consistent with the extruded polar bodies and were not associated with delayed or degenerated embryos. Importantly, caspase-positive blastomeres were not detected during cleavage stages, indicating that IGF1R loss does not induce apoptosis during early embryonic divisions (Supplementary Figure S2A). However, CS + cells were detected more frequently in KO blastocysts compared with WT controls, particularly in embryos exhibiting delayed development or abnormal morphology (Figure 4D). Quantitative analysis showed a higher percentage of CS + blastocysts in the KO group, although this increase did not reach statistical significance (p = 0.1397; Supplementary Figure S2B). At the transcriptional level, KO blastocysts exhibited reduced expression of the anti-apoptotic gene BCL2 and increased expression of pro-apoptotic genes BAX and CASPASE8, consistent with heightened apoptotic susceptibility (Figure 4C). These findings indicate that loss of IGF1R compromises blastocyst survival capacity, particularly under conditions of delayed developmental progression.

To account for potential compensatory signaling through INSR, OSI-906 (OSI), a dual IGF1R/INSR tyrosine kinase inhibitor, was used to inhibit IGF1R and INSR simultaneously across distinct developmental windows and dose ranges. Zygotes were treated continuously with OSI from the one-cell stage through blastocyst formation using concentrations ranging from 0.001 to 10 μM (Figure 5A; Supplementary Figure S3). Across this range, early cleavage to the 2-4 cell stage was largely unaffected at doses up to 1 μM, whereas blastocyst formation was modestly reduced only at the highest concentration tested (10 μM), indicating a broad tolerance to dual receptor inhibition during early cleavage stages. To assess the requirement for IGF1R/INSR signaling during blastocyst maturation more precisely, OSI was next applied specifically during the morula-to-blastocyst transition using a refined dose range (0.1, 1, 2, 5, and 10 μM; Figure 5B). Under these conditions, OSI treatment resulted in delayed blastocyst formation, reduced progression to expanded stages (Figure 5C), and increased apoptosis (Figure 5D), closely resembling the phenotype observed in IGF1R KO embryos. Increasing OSI concentrations did not induce more severe developmental impairment beyond this delay, nor did they cause widespread developmental arrest at doses ≤2 μM. Dual IGF1R/INSR inhibition did not worsen developmental outcomes compared with IGF1R genetic ablation alone, suggesting that INSR signaling does not play a dominant role during porcine preimplantation development under these conditions.

IGF-1 supplementation was used as a functional probe to test whether increased ligand input could compensate for impaired IGF1R/INSR-PI3K pathway capacity during the morula-to-blastocyst transition by treating morula-stage embryos with increasing concentrations of IGF-1 alone or in combination with OSI. IGF-1 supplementation alone exerted a dose-dependent effect on blastocyst formation (Supplementary Figure S4). Low to moderate IGF1 concentrations (10–50 ng/mL) modestly increased developmental efficiency, whereas high-dose IGF-1 (100 ng/mL) significantly reduced blastocyst formation compared with control embryos (Figure 6A). Combined treatment with IGF-1 (50 ng/mL) and OSI (2 µM) failed to restore normal blastocyst development. Morula-to-blastocyst rates in the combined treatment group remained significantly lower than those of controls. They were not improved relative to OSI alone (Figure 6A), indicating that ligand supplementation cannot bypass receptor-level inhibition. Consistent with prior work showing that IGF1-driven blastocyst responses are PI3K-dependent and blocked by IGF1R neutralization (Green et al., 2015), ligand supplementation did not overcome receptor-level inhibition in our system.

Apoptosis was assessed by caspase-3/-7 staining, which revealed a significant increase in apoptotic embryos following OSI treatment (p = 0.0113), whereas IGF-1 treatment alone did not differ significantly from controls (Figures 6B,C). The combined IGF-1 + OSI group exhibited intermediate levels of CS + embryos, not statistically different from the control, indicating partial attenuation rather than rescue of OSI-induced apoptotic stress. Apoptotic CS + cells were predominantly confined to small, poorly expanded, or developmentally delayed embryos. At the same time, well-expanded blastocysts at D7 across all groups were largely CS negative (Figure 6B) (Green et al., 2015). Gene expression analysis further supported these phenotypic findings. OSI-906 treatment was associated with marked suppression of SOX17 and dysregulation of cell-cycle regulators, including increased CDKN1B and altered CCND1 expression, whereas IGF-1 supplementation alone did not restore normal expression profiles (Figure 6D). IGF-1 exposure in the context of receptor inhibition does not compensate for impaired signaling and may exacerbate pathway imbalance. Collectively, these findings indicate that IGF-1 supplementation at the morula stage does not confer developmental benefit and may exacerbate stress responses when receptor signaling is compromised.