Targeted hypoxia-inducible factor 1-alpha (HIF1A) stabilization during in vitro maturation of bovine cumulus-oocyte complexes increases blastocyst rates
Martina Gübeli, Ulrich Bleul, Mariusz P. Kowalewski
TL;DR
Stabilizing HIF1A during in vitro maturation of bovine oocytes improves blastocyst formation rates, suggesting a potential method to enhance embryo development.
Contribution
The study demonstrates that low-dose Roxadustat treatment during in vitro maturation improves bovine blastocyst formation through HIF1A stabilization.
Findings
Low-dose Roxadustat (25 µM) significantly enhances blastocyst formation (p < 0.01).
HIF1A mRNA decreases in treated groups, but protein levels remain stable, indicating a feedback mechanism.
PHD inhibition benefits are specific to the maturation phase, not fertilization or cleavage.
Abstract
In vitro production (IVP) of bovine embryos frequently results in varying blastocyst yields, partly because culture conditions fail to support optimal oocyte and embryo development. In vivo, cumulus-oocyte complexes (COCs) mature under hypoxic conditions, and while in vitro protocols try to mimic the natural conditions, standard in vitro maturation (IVM) is typically performed under normoxia, guided largely by empirical outcomes rather than a biological understanding. We hypothesized that IVP efficiency would be improved by stabilization of hypoxia-inducible factor 1-alpha (HIF1A), via pharmacological inhibition of prolyl hydroxylase domain (PHD) activity, during IVM. To test this, we treated COCs with varying concentrations of Roxadustat, a PHD inhibitor. Low-dose treatment (25 µM) stabilized and significantly enhanced blastocyst formation (p < 0.01), while higher doses (100 µM)…
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Figure 5- —https://doi.org/10.13039/501100001711Schweizerischer Nationalfonds zur Förderung der Wissenschaftlichen Forschung
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TopicsReproductive Biology and Fertility · Pluripotent Stem Cells Research · Ovarian function and disorders
Introduction
Final oocyte maturation involves tightly coordinated nuclear and cytoplasmic changes that are essential for successful fertilization and early embryogenesis. These events are mediated by the surrounding follicular epithelial granulosa cells, whose oocyte-associated fraction forms the cumulus oophorus cells which, together with the oocyte, become the cumulus-oocyte complex (COC)^1,2^. The COC supports oocyte development via metabolic support, paracrine cues, and direct communication via gap junctions^3–5^. In addition to local cellular interactions, oocyte maturation is governed by a dynamic hormonal environment shaped by gonadotropin signaling. Following the FSH (follicle-stimulating hormone)-driven phase of follicular development, rising preovulatory levels of granulosa cell-derived estradiol (E2) trigger a surge of luteinizing hormone (LH), stimulating the developmental transition from primary to secondary oocytes, which then acquire the capacity to undergo fertilization^2^. In parallel with these important hormonal events, the physical environment within the follicle undergoes critical changes. Follicular vascularization increases with growth, but is limited to the theca cell layers^6,7^, resulting in an avascular intrafollicular environment surrounding the COCs, with the oxygen (O_2_) supply relying on diffusion from the theca vasculature. Thus, the maturation of COCs, including the steroidogenic activities of the granulosa cells, takes place under reduced O_2_ tension (hypoxia, referred to as physoxia). As shown in cattle and pigs^8,9^, ovarian blood flow decreases as ovulation approaches, further decreasing the ovarian O_2_ levels. Several studies have attempted to measure O_2_ tension in follicular fluid, but the findings were varied due to methodological limitations, including contamination by air and blood^10–15^. Nevertheless, mathematical modeling by Redding et al.^16^ supports the existence of a hypoxic niche around the maturing oocyte.
In vitro production (IVP) of embryos is a widely used assisted reproductive technology, but its overall efficiency remains limited, with widely varying blastocyst rates that average around 30% under standard in vitro conditions relative to the COCs used for in vitro maturation^17–21^. One factor potentially influencing the limited outcomes of IVP is the O_2_ tension during oocyte maturation. Although oocyte maturation occurs within physoxia in vivo, established standardized IVP protocols typically utilize atmospheric pO_2_ during in vitro maturation and fertilization (IVM and IVF)^22,23^. Interestingly, evidence regarding the optimal O_2_ tension during IVM remains inconclusive, with several reports indicating improved blastocyst rates under reduced O_2_ compared to atmospheric levels^19,24,25^, while others indicate detrimental effects of low O_2_ tension on oocyte developmental competence^26–28^. Notably, the addition of glucose to the maturation medium as an energy source under low O_2_ conditions restores the developmental competence of oocytes to levels comparable to those under atmospheric normoxia (21% O_2_)^29,30^. This suggests that the composition of the maturation medium plays a critical role in supporting oocyte development, further contributing to the variable IVP outcomes. The underlying mechanisms remain veiled, and so the commercial protocols are based on empirical values.
Hypoxia-inducible factor 1-alpha (HIF1A), a transcription factor that regulates a multitude of genes, including those involved in energy metabolism, erythropoiesis, and angiogenesis, is a key, and well-established, mediator of cellular adaptation to low O_2_ concentration^31,32^. While controlled HIF1A activation is essential for normal physiological processes, its excessive stabilization can be detrimental. Within the reproductive system, HIF1A plays essential roles in ovulation and early embryogenesis, as demonstrated in mouse models^33–35^. HIF1A directly promotes the function of steroidogenic cells, including ovarian granulosa cells. Among its important targets is the steroidogenic acute regulatory protein (STAR), a rate-limiting compound of the mitochondrial cholesterol transport machinery, directly linking O_2_ availability to steroidogenesis^36–38^. Consistent with this, our previous work demonstrated that pharmacological inhibition of HIF1A during IVM of bovine COCs reduces the transcriptional and translational availability of STAR, impairs progesterone (P4) production and cumulus expansion, and affects the expression of oocyte- and cumulus maturation-associated factors^39^. Finally, the detrimental effects of HIF1A suppression led to significantly reduced blastocyst formation rates^39^. Together, these findings position HIF1A as an important regulator and a key functional node linking hypoxia signaling to oocyte developmental competence.
The activity of HIF1A is regulated by both O_2_-dependent and O_2_-independent mechanisms. Under normoxic conditions (high O_2_ availability), HIF1A undergoes hydroxylation by prolyl hydroxylase domain enzymes (PHDs), specifically, the three isoforms PHD1, PHD2, and PHD3, and by factor inhibiting HIF (FIH), targeting it for proteasomal degradation via the von Hippel–Lindau (VHL) complex^40–43^. As these enzymes require O_2_ for their activity, hypoxic conditions reduce HIF1A hydroxylation, resulting in its stabilization. The accumulated HIF1A then dimerizes with its constitutively expressed dimer partner HIF1B and binds to hypoxia-responsive elements (HREs) of target genes^44^. Beyond O_2_-dependent regulation, HIF1A expression in granulosa cells is also modulated in an O_2_-independent manner, including its responsiveness to gonadotropins such as FSH and hCG or immune system-related factors^45–47^. In the ovary, these overlapping regulatory inputs allow HIF1A to contribute to the coordination of follicular responses to varying O_2_ levels and hormonal cues, supporting gene expression and functional pathways relevant to the development and function of COCs.
Targeted stabilization of HIF1A can be achieved by modulating PHD activity. In previous studies, Roxadustat^48,49^, a specific blocker of PHDs, achieved targeted PHD-mediated stabilization of HIF1A in murine granulosa cells^36^. A biphasic effect of HIF1A stabilization on STAR protein expression has been observed, where excessive HIF1A levels were associated with reduced STAR expression, similar to what has been observed upon STAR expression under severe hypoxia (1% O_2_) in the same study^36^. However, the underlying molecular mechanisms remain unclear. When investigating the expression of the three PHD isoforms, PHD2 emerged as the only O_2_-sensitive isoform, positively responding to decreasing O_2_ concentrations^36,50^. More recently, PHDs - particularly PHD2 - were found to be ubiquitously distributed across various cellular compartments in the bovine corpus luteum, indicating their spatial and temporal involvement in regulating luteal function^51^. Based on this, it is reasonable to assume that PHDs may also play a role in regulating HIF1A in COCs, with particular interest in the function of PHD2.
Based on existing evidence that HIF1A is essential for COC maturation and the acquisition of oocyte developmental competence for blastocyst formation^39^, and considering the biphasic HIF1A-dependent regulation of granulosa cells^36^, we hypothesized that PHD-dependent stabilization of HIF1A during IVP may positively influence the IVF outcomes. To address this hypothesis, we stabilized HIF1A by treating either bovine COCs or early embryos with Roxadustat to block PHDs.
Materials and methods
Collection of cumulus-oocyte complexes (COCs)
Ovaries were sourced from a local slaughterhouse, in compliance with national regulations on animal welfare and meat quality standards (not requiring ethics permits for use in research). For each experiment, 50–60 ovaries exhibiting small follicles (up to 8 mm) were selected from healthy animals. Upon retrieval, the ovaries were immersed in a 0.9% NaCl solution supplemented with 30 mg/L penicillin and 50 mg/L streptomycin and maintained at approximately 38.5 °C in a flask.
Prior to COC harvesting, the ovaries were washed in a warm 0.9% NaCl solution containing 60 mg/L penicillin and 100 mg/L streptomycin. The ovaries were then sliced with a sterile disposable scalpel and the follicular fluid collected into a beaker containing warm BO-WASH medium (IVF Bioscience, Cornwall, UK) supplemented with 0.2 mL of Heparin-Na (5000 IU/mL, B. Braun, DE). The COCs were filtered through a 100 μm cell strainer (Corning Falcon, Corning, NY, USA) and recovered by back-flushing the strainer with 15 mL of warm BO-WASH medium using an 18G cannula.
They were then observed under a stereo microscope (SZX7, Olympus, Tokyo, JAP), and those COCs characterized by at least two layers of compact cumulus cells and an evenly granulated ooplasm were selected and washed four times in warm BO-WASH medium prior to use in subsequent experiments. All processing steps were completed within five hours of ovary collection.
In vitro production (IVP)
The IVP protocol used in this study was based on the standardized, commercially available system provided by IVF Bioscience (Cornwall, UK), previously applied in our laboratory^39^. This protocol served as the foundation for all experiments and was modified only where explicitly stated.
Briefly, four morphologically homogenous groups of 40 COCs were matured in 0.5 mL oocyte maturation medium (BO-IVM, IVF Bioscience, Cornwall, UK) for 24 h at 38.5 °C in a humidified incubator (Incubator C16, Labotect, Göttingen, DE) with 5% CO_2_ and atmospheric O_2_ (21%).
IVF was carried out using sperm processed via density gradient centrifugation with BoviPure and BoviDilute (Nidacon International AB, Gothenburg, SE). Pre-warmed gradients were prepared by layering 40% (v/v) BoviPure over 80% (v/v) BoviPure, followed by the addition of 0.25 mL thawed cryopreserved bull semen of proven fertility. The sample was centrifuged at 350 × g for 15 min, and the supernatant was discarded. The resulting pellet was washed twice with 750 µL of Semen Preparation medium (IVF Bioscience, Cornwall, UK) by centrifugation at 350 × g for 3 min.
Sperm concentration was determined using a Neubauer chamber, and sperm were added to each well containing matured COCs in 0.4 mL of pre-warmed IVF medium (BO-IVF, IVF Bioscience, Cornwall, UK) at a final concentration of 1 × 10^6^ sperm/mL. Fertilization proceeded for 19 h in a humidified incubator (Incubator C16, Labotect, Göttingen, DE) at 38.5 °C with 5% CO_2_ and 21% O_2_.
Following fertilization, zygotes were manually denuded using The Stripper™ pipettor with 135 μm and 125 μm tips (CooperSurgical, Måløv, DK), transferred to 0.5 mL of pre-warmed embryo culture medium (BO-IVC, IVF Bioscience, Cornwall, UK), and overlaid with 0.4 mL paraffin oil (BO-OIL, IVF Bioscience, Cornwall, UK). Embryo culture was then conducted continuously in the same medium and well for 8 days at 38.5 °C in a humidified incubator with 6–7% CO_2_, and 6% O_2_ (Direct Heat Incubator SMA, ASTEC, Fukuoka, JAP).
All procedures were carried out using sterile, non-treated 4-well plates (Nunc IVF Multi Dish, Thermo Scientific, DK).
Targeted blocking of PHDs during in vitro maturation (IVM)
To stabilize HIF1A during IVM, we inhibited PHDs by supplementing the maturation medium (BO-IVM) with increasing dosages of Roxadustat (FG-4592, MedChemExpress, NJ, USA). The prepared media were dispensed into a 4-well plate, which was pre-incubated at 38.5 °C for at least two hours to ensure temperature equilibration. Treatment concentrations (0, 25, 50, or 100 µM) were selected based on previous studies^36^. Homogenous groups of immature COCs were randomly assigned to one of the treatment conditions. Apart from the addition of Roxadustat, the IVP protocol was as described above (Fig. 1a). A total of six biological replicates were performed.
Fig. 1. Schematic overview of the experimental design for (a) Roxadustat treatment during IVM, and (b) Roxadustat treatment during IVC.
Targeted blocking of PHDs during in vitro culture (IVC)
An additional set of experiments was conducted to block PHDs during the 8-day IVC phase. Similar to the treatment applied during IVM, the IVP protocol was as described, except for the addition of Roxadustat to the culture medium (BO-IVC). Following IVF, embryos were randomly assigned to one of four wells containing culture medium supplemented with Roxadustat (Fig. 1b). A total of six biological replicates were performed.
Assessment of maturation, cleavage and blastocyst rates
Maturation rates were determined after 24 h of IVM by assessing the presence of the first polar body. To enable visualization, COCs were enzymatically dissociated using 80 IU/mL hyaluronidase (GM501, Gynemed, Lensahn, DE) at 38 °C for 5 min, followed by manual denudation using The Stripper™ pipettor, equipped with 135 μm and 125 μm tips (CooperSurgical, Måløv, DK), as described in Turhan et al.^39^. Denuded oocytes were examined under a stereo microscope (SZX7, Olympus, Tokyo, JAP) and a bright field microscope (IX71, Olympus, Tokyo, JAP) to determine polar body extrusion. A total of nine biological replicates were analyzed. In experiments involving IVF and IVC, cleavage rates (2-cell stage or later) and blastocyst formation rates were assessed on days 2 and 8 post-fertilization, respectively. All rates are calculated based on the initial number of COCs used for IVM (100%) and are expressed as percentages relative to that number.
RNA isolation and semi-quantitative real-time (TaqMan) PCR
Gene expression profiling was conducted on cumulus cells from the control and Roxadustat-treated groups, as well as from an additional group of immature COCs included as a reference^39^.
The immature group was treated with hyaluronidase, as described above, to separate the cumulus cells from the oocytes. The cumulus cells were collected by centrifugation at 1000 × g for 10 min. The resulting cell pellet was washed in PBS 1 × (137 mM NaCl, 2.7 mM KCl, 10 mM Na_2_HPO_4_, 1.8 mM KH_2_PO_4_), centrifuged again (1000 × g, 10 min), and stored in 500 µL TRIzol (Invitrogen, Carlsbad, CA, USA) at – 80 °C until further analysis. The remaining four groups underwent the same treatment 24 h later, post-maturation.
Total RNA extraction was performed using TRIzol reagent following the manufacturer’s instructions and established methodologies^52^. The quantity and purity of RNA were assessed with a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific AG, Reinach, Switzerland). The stock solutions of isolated total RNA samples were diluted to working concentrations of 200 ng/µL, which were then used in downstream protocols. To eliminate potential genomic DNA contamination, samples were treated with RNase-free recombinant DNase I (Roche Diagnostics, Basel, CH), as per the manufacturer’s instructions. Reverse transcription (RT) was then performed using reagents from Applied Biosystems (Thermo Fisher, Waltham, USA), with random hexamers as primers, in accordance with the manufacturer’s protocol. The resulting cDNA, synthesized from DNase-treated RNA, was used to perform semi-quantitative real-time TaqMan PCR (RT-qPCR). cDNA corresponding to 100 ng DNase-treated total RNA was used for each target gene using the ABI PRISM 7500 Sequence Detection System (Applied Biosystems, Thermo Fisher, Waltham, USA). All samples were analyzed in duplicate (technical replicates), with negative controls included by replacing cDNA with autoclaved water. Custom-designed primers and FAM/TAMRA-labelled probes were obtained from Microsynth (Balgach, CH), while commercially available systems were sourced from Thermo Fisher. Detailed information regarding the primers is listed in Table 1. Relative quantification of target gene expression was calculated using the comparative Ct (ΔΔCt) method, following the ABI PRISM 7500 guidelines and previously established protocols^52^. Gene expression levels were normalized to two reference genes: GAPDH and ACTB showing stable expression among biological replicates. Their stability was assessed by using an online tool RefFinder that incorporates several major computational programs (BestKeeper, Normfinder, geNorm and the comparative Delta-Ct method)^53^. Average Ct of the group with the lowest expression (i.e. the highest Ct values) served as calibrator (accounting for biological and technical replicates).
Table 1. List of primers and TaqMan systems used for RT-qPCR.GenePrimers used for RT-qPCR (Taqman system)Product (bp) HAS2 Commercially available:Applied Biosystems,* prod. no. Bt03212695_g1.Accession number: NM_174079.3119 TNFAIP6 Commercially available:Applied Biosystems, prod. no. Bt03210224_m1**Accession number*: NM_001007813.2124 TMSB4
Forward 5’-GCG CCC TCT GCA ACC ATG T-3’
Reverse 5’-CGA AGG CAG TGG ATT TCT CTC T-3’
TaqMan probe 5’-CCC GAT ATG GCT GAG ATT GAG AAG TTC G-3’ Accession number: XM_005222037.4108 HIF1A Commercially available:Applied Biosystems,* prod. no. Bt03259341_m1**Accession number*: NM_174339.3109 GAPDH
Forward 5’-GCG ATA CTC ACT CTT CTA CCT TCG A-3’
Reverse 5’-TCG TAC CAG GAA ATG AGC TTG AC-3’
TaqMan probe 5’-CTG GCA TTG CCC TCA ACG ACC ACT-3’ Accession number: NM_00103403482 ACTB Commercially available:Applied Biosystems,* prod. no.* Bt03279174_g1Accession number: NM_173979.3141
Protein preparation and western blotting
Following the collection and centrifugation of cumulus cells, as described for the RNA isolation, the cell pellet was washed with PBS 1x, centrifuged again (1000 × g, 10 min), and then resuspended in 10 µL of NET-2 lysis buffer (50 mM Tris-HCl, pH 7.4, 300 mM NaCl, 0.05% NP-40) supplemented with protease inhibitor (10 µL/mL, Sigma-Aldrich Chemie). Homogenization was performed by sonication using a Vibra-Cell sonicator (75186, Weilburg, USA) at 75 W for 10 s.
Protein concentrations were measured using the Bradford assay with a Smart Spec Plus spectrophotometer (Bio-Rad Laboratories, Munich, DE). Samples were solubilized in 4x sample buffer (25 mM Tris-Cl, pH 6.8, 1% SDS, 5% β-mercaptoethanol, 10% glycerol, 0.01% bromophenol blue). Due to limited protein availability, sample concentrations were standardized across groups based on the group with the lowest protein yield, resulting in final concentrations ranging from 0.5 to 1 µg/µL across experiments. Proteins (10 µL) were separated on 10% polyacrylamide gels and then transferred to methanol-activated polyvinylidene difluoride (PVDF) membranes at 100 V. Membranes were cut prior to blocking to allow for simultaneous probing with different primary antibodies. Each section was blocked in 5% skimmed milk in PBST 1 × (137 mM NaCl, 2.7 mM KCl, 10 mM Na_2_HPO_4_, 1.8 mM KH_2_PO_4_, 0.1% (w/v) Tween^®^ 20) for 1 h at ambient temperature, followed by overnight incubation at 4 °C with the primary antibodies listed in Table 2. Membranes were stripped with 0.1 M glycine, pH 2.6 for 1 h at ambient temperature and reblotted overnight at 4 °C with different antibodies, including the anti-ACTB antibody to normalize protein loading. After washing, membranes were incubated for 1 h at ambient temperature with HRP-labelled secondary antibodies: donkey anti-rabbit IgG (Pierce Biotechnology, IL, USA; catalog number 31460) or anti-mouse IgG (Promega, Dübendorf, CH; catalog number W402B), both diluted 1:7500. Protein detection was performed using SuperSignal West chemiluminescent substrate (Pierce Biotechnology, Rockford, IL, USA) and visualized using the ChemiDoc XRS + System and Image Lab software (Bio-Rad, CA, USA). Precision Plus Protein™ Standard (10–250 kDa, Bio-Rad, CA, USA) was used as a molecular weight marker. A Glow Writer phosphorescent pen (MIDSCI, Valley Park, USA) was used to annotate membranes for chemiluminescent exposure as the ladder is not fluorescent. Signals were recorded using Image Lab Software with automated exposure settings optimized for appropriate signal intensity and dynamic range. Semi-quantitative analysis of band intensities was performed using ImageJ software (US National Institutes of Health, MD, USA). A total of five biological replicates were used for western blots. Technical replicates could not be performed due to limited material availability. Data were normalized to the control in each experiment to reduce variability arising from biological replicates of primary cells. Uncropped blots are presented in Supplementary Fig. 1. The apparent overexposure of the Glow Writer signal seen in some images in Supplementary Fig. 1 results from the amount of pen applied to the membrane and does not correspond to the amount of protein loaded. The imaging system tends to overexpose the marker while still appropriately capturing the sample signals.
Table 2. List of primary antibodies used for Western blot analysis.ProteinAntibodies used for Western blotSize (kDa)HAS2Polyclonal Rabbit anti-HumanLSBio, Seattle, WA, USACatalog number: LS-C411405-20Dilution: 1:100064PCNAMouse monoclonalAbcam, Cambridge, UKCatalog number: AB29Dilution: 1:100029PHD2Rabbit monoclonalCell Signaling TechnologyCatalog number: 4835Dilution: 1:100050HIF1ARabbit polyclonalNovus Biologicals, Abington, UKCatalog number: NB100-134Dilution: 1:25093ACTBMouse monoclonalSanta Cruz Biotechnology, Santa Cruz, CA, USACatalog number: sc-69879Dilution: 1:200042
Statistical analysis
Statistical analyses were conducted using BioRender Graph (based on R version 4.2.2; BioRender, Toronto, CA). For Western blot results, data were normalized to the control in each experiment. For qPCR, the average Ct of the group with the lowest expression (i.e. the highest Ct values) served as the calibrator. Data were assessed for normality and homogeneity of variances using Saphiro-Wilk and Levene’s tests, respectively. For normally distributed data with equal variances, one-way ANOVA was applied. In cases of unequal variances, Welch’s ANOVA was used. Nonparametric data were analyzed using the Kruskal-Wallis test. Post hoc analyses were performed as follows: Dunnett’s test (following one-way ANOVA), Games-Howell test (following Welch’s ANOVA), or Dunn’s test (following the Kruskal-Wallis test), to compare the control group with the other groups. The specific statistical tests used for each analysis are provided in the corresponding figure legends. A p-value ≤ 0.05 was considered statistically significant. Data are presented as mean ± SEM.
Results
HIF1A stabilization during IVM increases blastocyst rates
To evaluate the impact of PHD-mediated HIF1A stabilization during IVM on oocyte development, we assessed maturation, cleavage, and blastocyst formation rates. Dosage-dependently, Roxadustat treatment significantly influenced maturation and blastocyst formation with, however, opposite effects (one-way ANOVA p < 0.05, details in Fig. 2); whereas the highest concentration (100 µM) was associated with lower rates of oocyte maturation (p < 0.05, Fig. 2a), the lowest concentration (25 µM, R25) enhanced and stabilized blastocyst formation (p < 0.01, Fig. 2c). No significant difference (p > 0.05) was observed between the groups for cleavage rates (Fig. 2b). Although maturation rates were similar between the 25 µM and control groups, numerically, R25 oocytes showed apparently lower developmental arrest after maturation, resulting in higher blastocyst rates (Fig. 2d).
Fig. 2. Effects of Roxadustat-mediated HIF1A stabilization during IVM on oocyte maturation and embryo development. COCs were matured in vitro in the presence of Roxadustat at 25 µM (R25), 50 µM (R50), or 100 µM (R100), or without treatment (Ctrl). (a) Maturation rates, (b) cleavage rates, and (c) blastocyst rates were analyzed. One-way ANOVA showed a significant effect of treatment on maturation (p = 0.02) and blastocyst rates (p = 0.01), but not on cleavage (p = 0.33). Dunnett’s post hoc test was used for comparisons to the control. (d) Developmental distribution of 40 COCs per group based on observed maturation, cleavage, and blastocyst rates. Data are presented as mean ± SEM. *p < 0.05, **p < 0.01.
Assessment of cumulus cells: expression of PHD2, HIF1A/HIF1A and PCNA
The expression of PHD2 and PCNA was assessed by Western blot, while HIF1A expression was determined at the transcript and protein levels (Fig. 3). All factors were detectable in immature as well as in matured cumulus cells. At the transcript level, the expression of HIF1A increased significantly in matured cumulus cells (p < 0.01, Fig. 3a) but was markedly reduced in the Roxadustat-treated groups (p < 0.01, details in Fig. 3). Despite this reduction at the transcript level, the HIF1A protein was unchanged (p > 0.05) in treated groups (Fig. 3b), indicating the stabilizing effect of Roxadustat upon the protein content. PHD2 seemed to increase gradually in response to Roxadustat and showed a significant response to its highest concentration (100 µM, p < 0.05, Fig. 3c). The protein content of PCNA, a marker of proliferation, was significantly suppressed by higher dosages of Roxadustat (50 and 100 µM, p < 0.01, Fig. 3d).
Fig. 3. Effects of Roxadustat treatment on HIF1A/HIF1A (a,** b**), PHD2 (c), and PCNA (d) expression in cumulus cells. Five groups were analyzed: immature (I), control-matured (Ctrl), and Roxadustat-treated (R25, R50, R100). (a) HIF1A mRNA expression was analyzed by one-way ANOVA (p < 0.001), followed by Dunnett’s post hoc test. (b-d) Representative Western blots and quantification of HIF1A, PHD2 and PCNA. The Kruskal-Wallis test revealed significant differences between the groups for PHD2 (p = 0.02) and PCNA (p = 0.003), but not for HIF1A (p = 0.41). Dunn’s multiple comparison test was performed as a post hoc analysis for the significant results. Data were normalized to the control to reduce variability arising from biological replicates of primary cells. Data are presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. SOD = standardized optical density related to ACTB.
Assessment of cumulus cells: morphology and expression of HAS2/HAS2, TNFAIP6 and TMSB4
To determine whether the increased blastocyst rates could be associated with modified expression of some of the known markers associated with cumulus expansion and meiotic resumption, the expression of HAS2 and TNFAIP6, as well as of TMBS4, was assessed. Due to the limited availability of species-specific or cross-reacting antibodies, the expression of HAS2 was assessed at both the mRNA and protein levels, whereas TNFAIP6 and TMSB4 were evaluated based on their transcript levels (Fig. 4). As expected, there was a significant increase in all markers after cumulus cell maturation (immature (I) vs. control-matured (Ctrl))^39^. Morphological assessment revealed cumulus expansion across all groups compared to immature COCs, though COCs treated with 100 µM Roxadustat seemed to have more compact (less expanded) cumulus structures (Fig. 4a). Although blocking of PHD activity had no significant effect at the gene expression of HAS2 (Fig. 4b), the respective protein levels were suppressed (p < 0.05, Fig. 4c). Both TNFAIP6 and TMSB4 were significantly affected by Roxadustat treatment (p < 0.05, Fig. 4d and p < 0.0001, Fig. 4e).
Fig. 4. Effects of Roxadustat-mediated PHD inhibition on COC morphology and expression of HAS2/HAS2, TNFAIP6, and TMSB4 in cumulus cells, and representative micrographs of COCs. Five groups were analyzed: immature (I), control-matured (Ctrl), and Roxadustat-treated (R25, R50, R100). (a) COCs post-IVM (Ctrl, R25, R50, R100) and immature COCs as a reference. (b, d, e) qPCR analysis of HAS2, TNFAIP6, and TMSB4 mRNA expression. (c) Relative protein expression of HAS2 with a representative Western blot (SOD = standardized optical density related to ACTB). Data were normalized to the control to reduce variability arising from biological replicates of primary cells. The Kruskal-Wallis test revealed significant differences in HAS2 mRNA (p = 0.007) and HAS2 protein (p < 0.001) expression and was followed by Dunn’s multiple comparison test. TNFAIP6 mRNA expression was assessed using Welch’s ANOVA (p = 0.002), with the Games-Howell post hoc test. TMSB4 mRNA expression was analyzed by one-way ANOVA (p < 0.001), followed by Dunnett’s post hoc test. Scale bars: 200 μm. Data are presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.
HIF1A stabilization during IVC decreases blastocyst rates
The effects of HIF1A stabilization during IVC were evaluated by assessing cleavage and blastocyst rates. Maturation rates were not included in the analysis, as all groups underwent an identical IVM protocol. Although cleavage rates did not differ significantly between control and Roxadustat-treated embryos (p > 0.05) (Fig. 5a), blastocyst formation was dramatically reduced by treatment with 100 µM Roxadustat (p < 0.01, Fig. 5b).
Fig. 5. Effects of HIF1A stabilization during IVC. COCs were cultured in vitro in the presence of Roxadustat at 25 µM (R25), 50 µM (R50), or 100 µM (R100), or without treatment (Ctrl). (a) Cleavage rates, and (b) blastocyst rates were evaluated. One-way ANOVA revealed no significant differences in cleavage (p = 0.8), but a significant effect on blastocyst formation (p < 0.001). Dunnett’s post hoc test was used to compare each treated group to the control. Data are presented as mean ± SEM. **p < 0.01.
Discussion
As indicated by functional studies, including our own, HIF1 activity, driven by its O_2_-sensitive subunit HIF1A, is essential for steroidogenesis^37,38,54^, ovulation^34^, oocyte development^39^ and luteal function^55^. Here, we explored whether, and to what extent, PHD-dependent stabilization of HIF1A during IVP could enhance oocyte developmental competence, with particular emphasis on IVM, the pivotal stage for the acquisition of competence.
The most important findings from our results show a dose-dependent dual effect of the PHD blocker, Roxadustat, on the IVP process. While impaired oocyte maturation was observed at higher concentrations, lower doses consistently resulted in increased blastocyst yields. Interestingly, the enhancement of blastocyst rates was not accompanied by a significant change in cleavage rates, suggesting that PHD-dependent stabilization of HIF1A does not directly promote fertilization or early embryonic development, but rather improves the developmental competence of oocytes. It is important to note that, due to the need for denudation to assess polar body extrusion, maturation rates were assessed in a separate cohort of COCs from those used to evaluate cleavage and blastocyst rates. Thus, direct statistical comparisons across developmental stages within same cohorts of oocytes were not possible.
The cumulative representation of developmental dynamics shown in Fig. 2d enables a clear numerical comparison across groups. Interestingly, the R25 group exhibited a distinct developmental pattern compared to the control, which is especially relevant given its improved blastocyst rates. Although the proportion of oocytes failing to mature was similar, and the cleavage rates were not significantly affected, the greatest loss in the control group appeared to occur during early embryonic development, resulting in lower blastocyst rates. Due to the similar maturation and cleavage rates, these effects seem to depend on the acquisition of oocyte developmental competence required for blastocyst formation. This is supported by the lack of difference in cleavage rates in experiments evaluating the effects of HIF1A stabilization during the IVC process of COCs matured using the standardized (non-blocked) protocol. The accumulation of HIF1A appeared to have a detrimental effect at the highest dosage of the blocker. Although the molecular and biological basis of these effects requires further evaluation, it needs to be mentioned that the O_2_ tension is routinely reduced (5–6% O₂) in standard protocols^22,23^. Under these hypoxic conditions, HIF1A degradation is naturally limited^31^; thus, additional pharmacological stabilization must lead to excessive HIF1A activity, potentially amplifying signaling beyond physiological levels and disrupting normal developmental processes. Cumulatively, our findings suggest that PHD-mediated HIF1A stabilization during IVM may not alter cumulus maturation per se, but instead supports the developmental competence of oocytes, ultimately enhancing their progression to the blastocyst stage. Combined with the effects we observed in the IVC blocking experiments, the impact of HIF1A-mediated cumulus-oocyte dialogue in this process cannot be excluded - particularly since HIF1A inhibition has been shown previously to drastically reduce the developmental competence of COCs, reflected in lowered blastocyst rates^39^. Our findings suggest that moderate HIF1A stabilization during IVM may enhance developmental competence, whereas excessive or prolonged activation during IVC disrupts the balance of HIF1A signaling, ultimately impairing embryonic progression to the blastocyst stage. Indeed, the IVP outcomes are inherently variable and influenced by numerous biological and technical factors, such as oocyte source, operator skill, and sperm quality^56,57^. This intrinsic variability often results in a wide range of reported blastocyst rates. Therefore, the improvements achieved here resulting in stabilization of blastocyst rates under tightly controlled experimental conditions are meaningful.
When we looked more closely into the responses of the cumulus in our experimental design, we confirmed that PHD2, the main oxygen-sensitive PHD isoform^36,50^, is expressed in bovine cumulus cells. This adds to a recent report of its ubiquitous luteal expression, including in steroidogenic luteal cells^51^. The increased levels of PHD2 in response to higher concentrations of Roxadustat suggest that it is HIF1A-sensitive, corroborating the previous findings in murine granulosa cells that showed increased PHD2 levels in response to reduced O_2_ levels^36^. Interestingly, while HIF1A protein levels remained stable across groups, its mRNA was consistently downregulated by Roxadustat. This transcriptional repression may represent a compensatory mechanism to prevent excessive HIF1A accumulation. Indeed, such a mechanism aimed at limiting prolonged or exaggerated cellular exposure to HIF1A activity has been demonstrated, and involves the REST (Repressor Element 1-Silencing Transcription factor) complex, which was shown to inhibit HIF1A transcription in human embryonic kidney cells under prolonged hypoxic conditions^58^. While this remains speculative in bovine cumulus cells, it may provide a plausible explanation for the observed mRNA suppression alongside stable HIF1A protein levels. Furthermore, at higher concentrations, Roxadustat led to reduced expression of PCNA, a marker of cell proliferation. Given its association with HIF1A expression and granulosa cell proliferation^59^, this downregulation suggests that excessive HIF1A stabilization may impair cumulus cell function.
Cumulus expansion depends on cytoskeletal remodeling and the formation of a mucoelastic extracellular matrix (ECM) composed primarily of long hyaluronan (HA) chains and HA-binding glycoproteins^60–62^. Inhibition of HA synthesis during IVM of bovine COCs has been shown to decrease cumulus expansion and maturation rates^63^, suggesting that HA production plays a supportive role in oocyte nuclear maturation.
HAS2 (hyaluronan synthase 2) catalyzes HA synthesis, whereas tumor necrosis factor alpha-induced protein 6 (TNFAIP6) contributes to ECM stabilization and organization, and thymosin beta-4 (TMSB4) regulates actin polymerization and cytoskeletal remodeling^61,64–66^. All three genes were upregulated after IVM, consistent with their established roles and our previous findings^39^. Their reduced availability at higher doses of Roxadustat occurred concomitantly with the decreased proliferation of cumulus cells. While these molecular alterations are suggestive of impaired ECM remodeling, a visible reduction in cumulus expansion was only apparent at the highest dosage of the PHD blocker, indicating that functional consequences may only manifest at elevated levels of HIF1A stabilization.
This study shows that controlled stabilization of HIF1A during IVM enhances bovine oocyte developmental competence, likely through PHD-dependent stabilization of HIF1A and activation of its responsive pathways, leading to increased blastocyst rates. Therefore, low-dose Roxadustat (25 µM) significantly improved and stabilized blastocyst yield, highlighting a promising approach to improve, or at least stabilize, IVP outcomes. These effects possibly involve cumulus-oocyte dialogue, without affecting the process of fertilization and early embryonic cleavage, which is supported by the lack of effect when low doses of Roxadustat were added during IVC. Although the precise molecular mechanisms remain incompletely understood, they suggest a complex and temporally dynamic regulation that warrants further investigation. Inhibition of the HIF1A- and O_2_-dependent PHD2 activity provides a targeted and controlled stabilization of HIF1A, in contrast to the multidirectional stabilization caused by CoCl₂^41,67,68^, which is often associated with detrimental effects on ovarian cell functionality due to exaggerated and uncontrolled HIF1A expression^69–71^.
Similarly, as previously shown for murine granulosa cell function^36^, the present study clearly suggests that the balanced availability of HIF1A during the IVP process is essential. Its moderately stabilized levels during IVM appear to support oocyte developmental competence, whereas excessive levels during IVC interfere with embryonic progression to the blastocyst stage. In this context, the compensatory downregulation of HIF1A mRNA alongside stable protein levels is an important observation, suggesting active feedback regulation and reflecting the complex control of HIF1A activity in cumulus cells, as suggested recently for the bovine corpus luteum^51^.
Future research should incorporate broader molecular profiling to capture the regulatory mechanisms underlying the beneficial effects of targeted, PHD2-dependent stabilization of HIF1A on IVP outcomes. It should include the assessment of downstream effects, as well as the long-term developmental competence of embryos following transfer.
Supplementary Information
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Supplementary Material 1
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